^
Enzyme
and Metabolic
Inhibitors
Volume II
Malonate, A nalogs, Dehydroacetate,
Sulfhydryl Reagents, o-Iodosobenzoate, Mercurials
Volume I General Principles of Enzyme Inhibition
Volume III lodoacetate
Maleate
A^-Ethylmaleimide
Alloxan
Ouinones
Arsenicals
Volume IV
in preparation
Uncouplers of Oxidative Phosphorylation
Dinitrophenol
Arsenate
Cyanide
Carbon Monoxide
Azide
Sulfide
Antimycin
Fluoroacetate
Parapyruvate
Urethanes
Chelating Agents
Volume V
in preparation
Protein Group Reagents
Heavy Metals (Copper, Zinc, Cadmium, Silver, etc.)
Dyes
Aldehydes
Dimercaprol
Antienzymes
Phlorizin
Selenite and Tellurite
Naturally Occurring Inhibitors
Cholinesterase Inhibitors
Monoamine Oxidase Inhibitors
Drugs as Inhibitors
Carbonic Anhydrase Inhibitors
Borate
Enzyme
and Metabolic
Inhibitors
Volume II
Malonate, Analogs, Dehydroacetate,
Sulfhydryl Reagents, o-lodosobenzoate. Mercurials
J. LEYDEN WEBB
School of Medicine
University of Southern California
Los Angeles, California
1966
ACADEMIC PRESS New York and London
Copyright © 1966, by Academic Press Inc.
all rights reserved.
no part of this book may be reproduced in any form,
by photostat, microfilm, or any other means, without
writt'^n permission from the publishers.
ACADEMIC PRESS INC.
Ill Fifth Avenue, New York, .s'ew York 10003
United Kingdom Edition published by
ACADEMIC PRESS INC. (LONDON) LTD.
Berkeley Square House, London W.l
Library of Congress Catalog Card Number: 62-13126
PRINTED IN THE UNITED STATES OF AMERICA
This volume is dedicated with sincere gratitude to the medical librarians
— Vilma, Ruth, Lilian, Clara, Esther, Michelle, Rose, Shari, Nancy, Nahida,
and others — who have not only helped me to the limit, hut have made each
visit to the library a pleasure and often the most delightful experience of
the day.
PREFACE
For those rare readers who may feel inclined to pursue their way through
Volumes II and III from beginning to end, I have tried to arrange the
chapters and sections in a logical and interdependent order. Malonate has
been approached first because its actions so well illustrate some of the
general principles covered in Volume I, and, indeed, malonate is discussed
in greater detail than any other inhibitor in order to suggest how one
would like to deal with all inhibitors if one had either the time or space.
Inasmuch as malonate is the classic substrate analog, the next chapter
takes up various types of analogs and here we are able to obtain some
rough idea of the energies involved in the interactions of inhibitors with
enzyme surfaces, as well as study some of the factors which determine
specificity. Some readers may feel that too much attention has been given
to these analogs, but I believe they represent a very important group of
inhibitors and illustrate many principles — competitive behavior, group
specific interactions, protection and reversal, and even nuituil depk Lion
kinetics since some analogs are extremely potent inhibitors — and, in
addition, contribute to our understanding of feedback inhibition and me-
tabolic regulation. Most of the remainder of the volumes is devoted to sub-
stances considered to react with SH groups, certainly one of the most
commonly used and important classes of inhibitors, about which it is sur-
prisingly difficult to find adequate and comprehensive treatment. Certain
aspects of inhibition have been treated in detail, not necessarily because
of any intrinsic importance, but because of the information which is
provided to help us comprehend the general phenomena of inhibition.
There are many ways of writing about inhibitors, and I have tried to alter
the approach according to what I believe to be the most interesting aspects
of each inhibitor. These aspects may not happen to be those which would
have been chosen by the reader, but it is impossible to cover any inhibitor
completely and present it from all viewpoints. On the other hand, there
are certain sections which I have been unable to make very interesting,
to organize into a coherent picture, sometimes because the data are insuf-
ficient or too heterogeneous, but nevertheless some worthwhile material
vu
Vlll PREFACE
can often be included in these areas. One finds much of the subject to be
somewhat disconnected and it is seldom possible to present an orderly-
clear version of any inhibition because of the gaps in our knowledge, but
one must consider that these isolated strands may some day be woven
into a durable fabric. Each chapter has in general been organized so that
the treatment proceeds from the simplest system to the higher levels of
organization, since this generates progressive understanding it may be
hoped, although one occasionally wishes that the effects on the simpler
systems could be appreciated against a background of the actions on tissues
and animals. Perhaps to some extent the historical introductions at the
beginning of most chapters may serve as provisional backgrounds. As in
many fields of science one is confronted with the problem of vertical or
horizontal presentations. Efforts, however inadequate, have been made to
correlate the results at the different levels, and it is hoped that unlike
bacteria under certain conditions this volume does not too much exhibit
the phenomenon of accumulation without synthesis, or suffer from an even
worse danger, that in the psychosynthesis of concepts and over-all pictures
some abnormal or spurious units have been lethally incorporated.
This periplus of the field of enzyme inhibition presents a rather large
and often heterogeneous group of information, but everything has been
selected for some reason; the reasons may be debatable, since different
readers come to a book for different purposes, but occasionally one detects
something in a report, perhaps intuitively, which others would not, and
hence includes it for reasons difficult to express. The half-life for the general
use of a book is, indeed, determined in part by the ability or good fortune
of the author to select that which will have the most value or pertinence
in the pseudopodal fronts of science. One has no time for justifications,
since some decisions have to be made, and only the naive think they can
please or help everyone, but there is perhaps one justification I feel impelled
to make. Certainly there will be those who ask why the effects of an inhib-
itor on the blood pressure or the central nervous system have been pre-
sented when there is little or no obvious correlation with any metabolic
inhibition, or why I have made up tables of tolerated or lethal doses, and,
in general, some may criticize the discussion of inhibitor actions which are
likely to be unrelated to enzyme inhibition, or at least for which there is
no direct evidence. In defense of this, I can only say that I believe we
should not so rigorously categorize the actions of inhibitors. The refusal
to consider the nonmetabolic actions has led many investigators to very
biased interpretations of their data. If we are interested in the mercurials,
we are, I assume, interested in all their possible actions, whether they are
PREFACE IX
based on metabolic disturbances or not. To be narrow here would be like
discussing only the beneficial effects of drugs and omitting the toxic actions.
Of course, space limitations make it impossible to treat all these actions
equally, and I have tried to emphasize those actions in which a disturbance
of metabolism is the most likely mechanism. But we must never ignore the
possibilities of other mechanisms with any inhibitor, particularly those
reacting with groups on proteins and other cell components. The mercurials
offer an especially clear example of enzyme inhibitors producing character-
istic effects on many tissues (e.g. kidney, heart, central nervous system,
liver, muscle, etc.) and where not a single action can be definitely corre-
lated with a mechanism involving enzyme inhibition. Nevertheless, with
further improvements in techniques and more knowledge, it is quite pos-
sible that in the future at least some of these actions will be related to
effects on enzymes. To be perfectly honest, at the present time we cannot
say in the majority of cases just how substances called enzyme inhibitors
act to produce their interesting and often clinically or industrially important
effects on microorganisms or tissues, and it is necessary to realize our igno-
rance so that progress in understanding may take place. Inhibitors do
produce some very intriguing effects on tissue function or in whole animals,
and many of these effects are unknown to those who look upon inhibitors
merely as biochemical tools; so by reading of these effects some may be
activated to study the ultimate causes in greater detail. Incidentally,
since inhibitors will be used more and more frequently in animals, infor-
mation on dosage ranges to produce various effects may serve a very prac-
tical purpose.
I would like to express my gratitude to those who have written to me
saying they have found the first volume of interest or of some value to
them, who have sent me unpublished manuscripts or difficultly obtainable
material, and who have given me encouragement during those periods
when I sincerely wished I were in a monastery in Kyoto.
J. Leyden Webb
November, 1965
CONTENTS
Dedication v
Preface vii
Introduction xv
Conventions xix
Symbols xix
CHAPTER 1
Malonate 1
Early Historical Development 1
Chemical Properties 3
Inhibition of Succinate Dehydrogenase 15
Inhibition of Succinate Oxidation in Cellular Preparations .... 50
Inhibitions of Enzymes Other Than Succinate Dehydrogenase ... 58
Effects of Malonate on the Operation of the Tricarboxylic Acid Cycle . 69
Accumulation of Succinate during Malonate Inhibition 90
Accumulation of Cycle Substrates Other than Succinate 104
Antagonism of Malonate Inhibition with Fumarate 112
Specificity of Malonate Inhibition in the Cycle 117
Effects of Malonate on Oxidative Phosphorylation 118
Effects of Malonate on Glucose Metabolism 122
Effects of Malonate on Lipid Metabolism 135
Effects of Malonate on Amino Acid and Protein Metabolism .... 151
Effects of Malonate on Porphyrin Synthesis 158
Effects of Malonate on Miscellaneous Metabolic Pathways 163
Effects of Malonate on the Endogenous Respiration 166
Permeability of Cells to Malonate 186
Growth, Development, and Differentiation 192
Cellular and Tissue Function 202
Effects of Malonate in the Whole Animal 217
Effects of Malonate on Bacterial Infections 221
Metabolism of Malonate 224
Inhibitors Structurally Related to Malonate 235
XI
xii CONTENTS
CHAPTER 2
Analogs of Enzyme Reaction Components 245
Terminology 246
Possible Sites and Mechanisms of Inhibition 246
Kinetics of Analog Inhibition 248
Means of Expressing Results 252
Important Types of Molecular Alteration Producing Inhibiting Analogs . 255
Development of the Concept of Inhibition by Analogs 259
Analog Inhibition of Membrane Transport 261
Analogs Which Are Isomers of Substrates 268
Fumarase 274
Inhibition of Xanthine Oxidase by Purine Analogs and Pteridines . . 279
Choline Oxidase 290
Inhibition of Nitrogen Fixation by Other Gases 291
Phenol Oxidases 296
Tyrosine Metabolism 302
Tryptophan Metabolism 321
Glutamate Metabolism 327
Arginase 335
L-Amino Acid Oxidases 338
D-Amino Acid Oxidase 340
Analog Inhibition of the Metabolism of Various Amino Acids .... 350
Diamine Oxidase (Histaminase) 360
Carboxypeptidase, Aminopeptidases, and Dipeptidases 365
Chymotrypsin and Other Proteolytic Enzymes 368
Hexokinases 376
Effects of 2-Deoxy-D-Glucose on Carbohydrate Metabolism .... 386
Effects of 6-Deoxy-6-Fluoro-D-Glucose on Metabolism 403
Various Analog Inhibitors of Carbohydrate Metabolism 405
Glycosidases 415
Pyruvate Metabolism 429
Lactate Metabolism 432
Phosphatases 439
Sulfatases 443
Adenosinetriphosphatases and Transphosphorylases 444
Hydroxysteroid Dehydrogenases 447
Nitrite and Sulfite Metabolism 450
Simple Ion Antagonisms 452
Inhibition by Macroions 453
Inhibitions by Nucleotides and Related Substances 465
Inhibitions by Coenzyme Analogs 482
Analogs of Nicotinamide and the Pyridine Nucleotides 484
Analogs of Thiamine 514
Analogs of Riboflavin and FAD 534
Analogs of Pyridoxal 561
Analogs of Pteroylglutamate (Folate) 579
CONTENTS Xni
Analogs of Other Vitamins, Coenzymes, and Their Components . . . 586
Miscellaneous Analog Inhibitions 590
CHAPTER 3
Dehydroacetate 617
Chemical Properties 618
Inhibition of Enzymes 620
Effects on Respiration and Glycolysis 623
Effects on Tissue Functions 624
Effects on the Whole Animal 627
Distribution and Metabolism 629
Antimicrobial Activity 631
CHAPTER 4
Sulfhydryl Reagents 635
Role of SH Groups in Metabolism and Function 636
Chemical Properties of SH Groups 637
Types of SH Reaction Important in Inhibition 642
Factors Determining the Reactivities of SH Groups 643
Interpretation of Inhibitions by SH Reagents 647
Protection and Inhibition Reversal by Thiols 650
General Considerations of the Uses of SH Reagents 651
CHAPTER 5
Oxidants 655
Disulfides 661
Porphyrexide and Porphyrindin 664
Ferricyanide 670
Iodine 678
Peroxides 690
Tetrathionate 696
CHAPTER 6
o-lodosobenzoate 701
Chemistry 701
Reaction with Protein SH Groups 703
Inhibition of Enzymes . 704
Inhibition of Metabolism 721
Effects on Animal Tissue Functions 723
Effects in Whole Animals 724
Effects on Sea Urchin Egg Development 726
Effects on Bacteria and Viruses 727
XIV CONTENTS
CHAPTER 7
Mercurials 729
Chemical Properties 730
Reactions with Proteins 751
Inhibition of Enzymes 768
Electron Transport and Oxidative Phosphorylation 870
Fermentation and Glycolysis 874
Tricarboxylate Cycle 877
Respiration 879
Various Metabolic Pathways 886
The Cell Membrane as a Site for Mercurial Action 892
Effects on Permeability and Active Transport 907
Effects on the Kidney 917
Effects on Tissue Functions 937
Effects Observed in the Whole Animal 950
Effects on Mitosis, Growth, and Differentiation 963
Effects on the Growth of Microorganisms 970
Development of Resistance to Mercurials 983
References 987
Author Index 1071
Subject Index 1125
INTRODUCTION
Certain principles or prejudices in the approach should be clearly stated
since these occasionally heretical opinions have been the basis for much of
the organization of this volume.
What is hoped to be pertinent information on the physical and chemical
properties of the inhibitors has been given in the belief that the proper use
of any inhibitor requires as much knowledge of its properties as possible.
Such data are often difficult to find and I am afraid that much important
material has been omitted.
The quantitative formulation of inhibitions has been stressed because
here, as in every science, progress often depends on accurate recording and
reporting of observations. It is, for example, not only not informative but
actually misleading to state that aldehyde oxidase is or is not inhibited
by p-mercuribenzoate. What is meant by " inhibited " — 10%, 50%, or
100%? What is the concentration of the inhibitor? If it is 0.01 mM it may
mean something, but if it is 10 mM probably nothing. What is the source
of the enzyme? There are many aldehyde oxidases and they differ quite
markedly according to their sources. What substrate and acceptor were
used? Is the substrate acetaldehyde, glyceraldehyde, formaldehyde, reti-
nene, or even hypoxanthine, and are electron acceptor dyes used or the Og
uptake measured? These and other factors must be made explicit. Of
course, it is impossible in a book like this to give a complete picture of
each inhibition mentioned, but I have tried to state the source of the
preparation, the substrate, the inhibitor concentration, and the per cent
inhibition in every case, as well as the pH and the incubation times when
necessary.
One can understand a phenomenon better when it can be visualized in
some manner and much of the recent work on the inhibition of enzymes
has been done with the purpose of clarifying the topography of the en-
zyme surface and the nature of the interactions occurring there. Thus I
have tried to emphasize the interpretation of data in terms of an accurate
delineation of group orientation and intermolecular forces, although in the
present state of our knowledge this can seldom be done satisfactorily.
XV
XVI INTRODUCTION
Metabolism within cells is almost always a matter of multienzyme sys-
tems and so the effects of inhibitors on such systems have been discussed
fully wherever possible, although this is even more difficult to describe
quantitatively than the behavior of single enzymes.
The importance of the specificity of inhibition was sufficiently emphasiz-
ed in the previous volume and it should be clear that this is a critical prob-
lem which has been neglected, ignored, or abused extensively. It is not
an easy matter to evaluate the specificity of an inhibitor under various
conditions, particularly when the necessary data are lacking, but it is
hoped that at least a provisional picture has been presented in some in-
stances.
Certain aspects of metabolism (e.g. glucose utilization, respiration, pho-
tosynthesis, protein synthesis, or oxidative phosphorylation) and cellular
activity (e.g. active transport, membrane potentials, movement, mitosis,
or proliferation) are obviously of general significance, and the effects of
inhibitors on these have been emphasized. This is not to say that other
pathways or functions are unimportant, and indeed where necessary they
have been treated as adequately as possible, but one cannot discuss all
the actions of each inhibitor, so that some compromises must be made.
A major use of inhibitors is in the attempt to correlate cellular functions
with particular enzymes or metabolic pathways, and for this reason, as
well as the fact that this represents one of the most fascinating aspects of
inhibitor study, these correlations have been discussed fully if the infor-
mation has been available, and the effects on certain organisms or processes
have often been given in the hope that some correlation will emerge or
further work will be stimulated. It is believed that conceiving inhibitor
actions in terms of deviations in the energy flow is of some value although
an accurate formulation of this must await the development of a new
terminology.
It is simpler to restrict the treatment of an inhibitor's action to a par-
ticular organism or tissue, but it is felt that a great deal may be learned
from comparative inhibitor enzymology. Therefore, in the tables, the ef-
fort has been made to present the results from as many sources as possible
for a particular enzyme or metabolic pathway since by doing this one is
better able to see the great extent of the variability in responses; only a
distorted view is obtained if a limited range of action is considered.
Paradoxical actions have been both the despair and delight of scientists
in many fields, and it is recognized that some of our finest theories have
originated in the observation and study of anomalies. There is an inherent
desire in most of us to eliminate anomalies and perhaps devote a good deal
INTRODUCTION XVU
of effort to this, since we feel that an anomaly really is something we would
expect if we knew the system or mechanism better, or as Henry Miller
has said in the " Tropic of Capricorn," " confusion is a word we have in-
vented for an order which is not understood." I have thus brought up
certain so-called anomalies, not only for their interest but again because
they often stimulate deeper investigation, although at present they may
to some only confuse the picture.
Many of the results have been put into tabular form, first because this
is the most efficient way of presenting certain types of data, second because
such simple observations are often the sole information on the inhibitors
provided in the reports, third because this allows a more convenient com-
parison of results (e.g., for those interested in possible phylogenetic rela-
tionships, for which reason the source organisms have usually been given
in the classic taxonomic sequence, or for studying the variability in re-
sponses on a comparative basis), fourth because this is the clearest way
to provide information from which specificity may be evaluated, and fifth
because these tables may serve as reference sources for those interested in
the actions of a particular inhibitor on a certain enzyme or organism.
There is much more in these tables than anyone can assimilate or under-
stand or interpret today, but it is these data which could possibly con-
tribute to some idea or concept if placed against the proper experience or
background. Nothing makes some data look more miserable or incomplete
than putting them in tables, but perhaps this is an asset, since it shows
what is missing, what should have been done, and what more there is to
do. A great deal of information could not be included in the tables, for,
although some of them look formidably long, they represent only a frac-
tion of what is available in reports. One tries to include only that which
is important, but the definition of this word becomes more difficult as one
applies it. There are so many very specialized and unique enzymes being
isolated and studied these days that it becomes more of a problem each
year to determine which of the enzymes are generally significant. An en-
zyme which at first sight might seem esoteric, if for no other reason than
its gargantuan name, implying a specificity of catalysis incommensurate
with anything but a very limited role in metabolism, may well be of great
importance in a particular pathway, a pathway perhaps as yet undiscover-
ed. Every enzyme is of some importance to some organism or tissue, or it
would not be there. And we often take a limited viewpoint; one of the
numerous enzymes in the pathway of steroid biosynthesis is recognized as
important in cholesterol or adrenal corticoid formation, but it may be equal-
ly important to some microorganism in producing steroids which function
XVIU INTRODUCTION
in their membranes, the inhibition of the formation of which could lead
to a suppression of growth. In view of the past history of science, anyone
is presumptuous to claim they can distinguish what is important from what
is not — we have to do this much of the time, of course, but we should
realize we are presumptuous. There are probably some errors in the tables,
since it is often difficult to determine exactly the conditions used; one is
sometimes referred to a previous report, but cannot be certain that all the
conditions have been maintained throughout the work. One must often
guess a parameter from other work the investigators have done, and some-
times calculate results from heterogeneous data. There has been a good
deal of calculation, and recalculation, and averaging, and I take full re-
sponsibility for anything right or wrong I may have done. A number of
curves have been replotted or data represented in a way that differs from
that of the original investigator, and I fully realize that this usuallj' results
in nothing but animosity.
CONVENTIONS
The naming of enzymes is not an easy task. On the one hand, there are the more
trivial names with their occasional confusions — on the other, there are the official
names in the " Report of the Commission on Enzymes " (1961) which are reasonably
precise but often unwieldy. I have usually chosen the former because I feel most
readers will recognize these more readily, but frequently I have taken an interme-
diate course which probably will not please anyone. It is much more accurate to
write NADH:menadione oxidoreductase than to use the designation NADH oxidase
or NADH dehydrogenase, since the former name indicates the substrate and acceptor
used. In addition it is cumbersome to use D-xylulose-5-phosphate D-glyceraldehyde-
3-phosphate-lyase (phosphate-acetylating) instead of phosphoketolase, yet there is no
doubt that this longer terra accurately describes the enzyme. There are also prefer-
ences in nomenclature, for various reasons. I never cared much for the term invertase;
I prefer to call it [ii-fructofuranosidase, although it is clumsier, but not as much so
as P-D-fructofuranoside fructohydrolase. In other instances the older and shorter
names are more pleasing to me and I imagine to others. I have tried to use enzyme
names which, at least, can be found in the index of the " Report of the Commission
on Enzymes," and some cross referencing of names has been included in the index.
There are certain instances of inconsistency which I do not particularly regret.
As in the first volume, concentrations have been given as millimolar (m.M) except
when designated otherwise, and in other matters the conventions given there have
been retained.
SYMBOLS
A
absorbance
DQ
ADP
adenosinediphosphate
DQH,
AMP
adenosinemonophosphate
Eo
ATP
adenosinetriphosphate
9,10-AQ
9,10-anthraquinone
Eo'
BAL
dimercaprol
ChE
cholinesterase
ED,
CoA
coenzyme A
6-DFG
6-deoxy-6-fluoro-D-glucose
EDTA
2-DG
2-deoxy-D-glucose
EI
DNA
deoxyribonucleic acid
EM
DNP
2,4-dinitrophenol
duroquinone
durohydroquinone
standard oxidation-reduc-
tion potential (pH = 0)
oxidation-reduction poten-
tial at specified pH (usually 7)
effective dose or concentra-
tion for X per cent
ethylenediaminetetraacetate
enzyme-inhibitor complex
Embden-Meyerhof (path-
way)
XX
SYMBOLS
EP
enzyme-product complex
NEM
iV-ethylmaleimide
Epi
epinephrine
1,2-NQ
1 ,2-naphthoquinone
ES
enzyme-substrate complex
1,4-NQ
1 ,4-naphthoquinone
FAD
flavin-adenine dinucleotide
pl
- log (I)
FDP
fructose- 1 ,6-diphosphate
p-MB
p-mercuribenzoate ion
FMN
flavin mononucleotide
p-MPS
j9-mercuriphenylsulfonate
GSH
reduced glutathione
ion
GSSG
oxidized glutathione
P-Q
2)-benzoquinone
i
fractional inhibition
P-QH,
p-benzohydroquinone
if
final fractional inhibition
(hydroquinone)
lA
iodoacetate
pS
- log (S)
lAM
iodoacetamide
P-XQ
p-xyloquinone
IC
intracutaneous
9,10-PAQ
9, 10-phenanthraquinone
IM
intramuscular
3-PGDH
3 - phosphogly ceraldehyde
IMP
inosinemonophosphate
dehydrogenase
IP
intraperitoneal
PM
phenylmercuric ion
ISBZ
o-iodosobenzoate
Pyr
pyruvate
IV
intravenous
SC
subcutaneous
Ka
ionization constant
SH
sulfhydryl
Ki
inhibitor constant
S/M
slice/medium ratio
Km
Michaelis constant
s— s
disulfide
Ks
substrate constant
TD
tolerated dose or concentra-
LD.
lethal dose or concentration
tion
for X per cent
T/M
tissue/medium ratio
MD
menadione
TQ
toluquinone
MHD2
menadiol
TQH3
toluhydroquinone
MLD
minimal lethal dose
V
rate
MM
methylmercuric ion
Vm
maximal rate
NAD
nicotinamide-adenine
P-XQ
p-xyloquinone
dinucleotide
£
molar extinction coefficient
NADP
nicotinamide-adenine
(p-AsO
phenylarsenoxide
dinucleotide phosphate
99-ASO2
phenylarsonic acid
Enzyme
and Metabolic
Inhibitors
Volume II
Malonate, Analogs, Dehydroacetate,
Suljhydryl Reagents, o-Iodosobenzoate, Mercurials
CHAPTER 1
MALONATE
Malonate is one of the most interesting, specific, useful, and well-known
enzyme inhibitors, and for these reasons will be discussed in detail in order
to illustrate some of the general principles delineated in the first volume.
It will be valuable perhaps to take up one inhibitor to the degree necessary
to consider many of the various problems in the application of these prin-
ciples, and to examine the pitfalls that may appear even in the use of an
inhibitor that in several ways approaches what one would want ideally.
Much of what will be said concerning malonate may be applied to the other
inhibitors. This discussion will emphasize the often overlooked fact that
the effect of relatively simple inhibitors in cells may constitute a very
complex problem and that their use in elucidating metabolic relationships in
tissues or whole organisms should not be undertaken lightly. It is a simple
matter to apply an inhibitor such as malonate but it is often very difficult
to use it properly and to interpret the results accurately. The treatment
of malonate will, furthermore, provide a foundation for the more general
discussion of competitive inhibitions produced by analogs in the following
chapter.
EARLY HISTORICAL DEVELOPMENT
The first report of the use of malonate in a biological system was made
by Heymans (1889) to the Physiological Society in Berlin. The toxicity
of oxalate to animals had been known for many years and Heymans be-
lieved that an investigation of the higher homologs of the dicarboxylate
series might be interesting. Although sodium oxalate was quite poisonous
when injected into the frog dorsal lymph sac, the sodium salts of malonate,
succinate, and glutarate were essentially without effect, sodium malonate
being nonlethal at a dose as high as about 8 g/kg. However, Pohl (1896),
working in Prague, found that the urinary excretion of oxalacetate was
increased by administering malonate to dogs and, furthermore, that only
a small portion of the malonate given could be recovered in the urine,
indicating that the dog can metabolize malonate. The first experiments
showing the metabolic inhibitory action of malonate were done by Thun-
2 1. MALONATE
berg (1909) in Lund. He had observed the inhibition of minced frog muscle
respiration by oxalate and decided to study the higher homologs. Thus the
inhibitory action of malonate on muscle respiration was demonstrated,
whereas succinate instead stimulated the oxygen uptake. Apparently this
observation went unnoticed and the inhibitory activity had to be rediscov-
ered later. Rose (1924) at Illinois showed that malonate exhibits no nephro-
toxic action, as does glutarate, when given orally to rabbits, although he
later (Corley and Rose, 1926) found that the methyl and ethyl derivatives
depress renal function. At about the same time, Momose (1925) in Japan,
continuing the work of Pohl, observed that malonate when perfused through
dog liver, gives rise to acetoacetate, acetone, and aldol. He postulated that
these substances arise from malonate after decarboxylation to acetate, but
it is more likely from our present knowledge that malonate gives rise to
these substances by a disturbance of the metabolism. During the next
few years evidence was accumulated that malonate can arise from normal
tissue metabolism — occurring in alfalfa (Turner and Hartman, 1925)
and wheat (Nelson and Hasselbring, 1931), and appearing during citrate
fermentation in the mold Aspergillus (Challenger et al., 1927) — and be
metabolized by certain microorganisms, such as Escherichia coli (Grey,
1924).
Our present concepts of the inhibitory action of malonate, however,
arose from the work on bacterial dehydrogenations by Quastel and Whetham
(1925) at Cambridge. They tested the abilities of various dicarboxylic acids
to reduce methylene blue in suspensions of E. coli and found that only suc-
cinate is active. Malonate inhibited this reduction by succinate. As stated
in their own words, "Oxalic, glutaric, and adipic acids (when mixed with
succinic acid) do not retard the reduction due to the succinic acid, but
malonic acid has a definite retarding effect. It is difficult to explain the
anomalous behaviour of malonic acid, but there is no doubt as to the reality
of the effect." They found the methylene blue reduction time with succinate
bo be tripled in the presence of 77 mM malonate. Quastel and Wooldridge
(1928) extended this work to show that the action on succinate oxidation is
rather specific in that malonate does not appreciably inhibit the oxidation
of several other substrates by E. coli. But in addition they demonstrated
that increasing succinate concentrations would counteract the malonate
inhibition, leading them to suggest that both substances are adsorbed to
the enzyme reversibly, probably competing for the same active site. Final-
ly, Quastel and Wheatley (1931), now at the Cardiff City Mental Hospital,
reported that the malonate inhibition of succinate oxidation occurs in
many bacteria, and in mammalian brain and muscle as well, the enzymes
from the mammalian tissue being even more sensitive.
The concept of the competitive inhibition of an enzyme by a substance
structurally related to the normal substrate was first clearly demonstrated
CHEMICAL PROPERTIES 6
and expressed for this inhibition of succinate oxidation by malonate in the
work of Quastel and Wooldridge, although competition specifically was not
mentioned. Cook (1930), also at Cambridge, however, stated that a "com-
petitive" mechanism had been established, presumably referring to the
work of Quastel inasmuch as Cook performed no experiments indicating
a competitive relationship. The competitive nature of the malonate inhibi-
tion has been substantiated many times and placed on a quantitative basis,
so that malonate has come to be recognized as the classical example of
inhibition by a pvirely competitive mechanism. The development and ap-
plications of this concept will be discussed in more detail in Chapter 2.
For 20 years malonate was the only available specific inhibitor of succinate
dehydrogenase, and later of the tricarboxylic acid cycle, and actually played
an important role in the elucidation of the cycle sequence. Other cycle inhib-
itors have been described recently, but no other inhibitors of the succinate
oxidation step as specific and useful as malonate have been found.
CHEMICAL PROPERTIES
Malonic acid and its salts when obtained commercially are often not
sufficiently pure for accurate work and it has been the practice in our lab-
oratory to recrystallize all material. Malonic acid may be recrystallized
from ethyl acetate and benzene (Adell, 1940), or ether and benzene con-
taining 5% light petroleum (Vogel, 1929), or simply from a hot concentrated
benzene solution by cooling to 50-10°. The sodium and potassium salts may
be dissolved in small amounts of warm water and precipitated by the ad-
dition of ethanol, as is commonly done with other dicarboxylate salts
(Potter and Schneider, 1942), or, with somewhat less yield, may be crystal-
lized by cooling hot concentrated aqueous solutions. In all cases we have
decolorized with activated charcoal in the solutions before recrystallization
and have washed the products with ether preparatory to drying. It should
be emphasized that the choice of the sodium or the potassium salt will
depend on whether the preparation to be tested is cellular or subcellular.
Stability
Malonic acid and its salts are quite stable and chemically unreactive.
Decarboxylation to acetate proceeds very slowly under ordinary conditions.
Aqueous solutions of sodium malonate heated to 125° for 48 hr show no
perceptible decomposition (Fairclough, 1938), and the half-life of sodium
hydrogen malonate in solutions 5-50 mM is at 80° around 40 days, cal-
culated from the rate constant for decarboxylation (Hall, 1949). The free
energy change for the reaction
Malonate -> acetate + CO,
4 1. MALONATE
is approximately —7 kcal/mole but the activation energy is 27.9 kcal/mole
(Gelles, 1956). Thus at physiological temperatures one may consider mal-
onate as completely stable. Nevertheless, there are enzyme systems which
catalyze the decarboxylation (see page 227) and one should be reasonably
certain in the use of malonate that it is stable in the system investigated,
since the formation of acetate might well confuse the results.
Molecular Structure
In crystals of malonic acid the molecules are arranged in zigzag chains
with the carboxyl groups linked through two hydrogen bonds (Goedkoop
and MacGillavry, 1957). The following bond parameters were observed
(the two values refer to the two carboxyl groups, since the molecule in the
crystal is apparently not symmetrical): C — C — C angle = 110^; C — C
distance = 1.54, 1.52; C— 0 distance = 1.29, 1.31; C=0 distance = 1.24,
1.22; 0— C— 0 angle = 128^, 128°; and H-bond distance = 2.68-2.71.
The malonate ion in solution would probably deviate somewhat from this
configuration but not a great deal inasmuch as malonate is fixed in a rather
rigid structure, because the C — C — C angle is determined by the electronic
tetrahedral orbitals and can be distorted only with difficulty. The carbox-
ylate groups can rotate around the C — C axis but they presumably ster-
ically interfere with each other when both lie in the plane of the molecule,
since the centers of the oxygen atoms would be 2.2 A apart and the van
der Waals' radius of the oxygen atom is 1.4 A (Goedkoop and MacGillavry,
1957). In malonic acid crystals one carboxyl seems to be in the molecular
plane and the other is at right angles; in the malonate ion it may well be
that neither is in the C — C — C plane. However, another factor must be
considered; it is possible that in solution there is intramolecular hydrogen
bonding (Gelles, 1956), at least for the hydrogen malonate ion. When the
carboxyl group ionizes, the equivalence of the structures:
R-cf R_c--°
allows a greater resonance than in the unionized state (equivalent to an
extra 8 kcal/mole energy), and this high resonance would indicate an inter-
mediate structure in which the center of negative charge lies midway be-
tween the two oxygen atoms. Although keto-enol tautomerism occurs
(Hofling et al., 1952) in the esters of malonic acid:
O OH
-CH^-C-OEt ^ -CH=C-OEt
it is probably not significant in the malonate ion because it would reduce the
electronic resonance.
CHEMICAL PROPERTIES 5
The distance between the two centers of negative charge in malonate
is of importance in the binding to succinate dehydrogenase. Calculation of
this distance (using the following values: C — C — C angle = 111.7°; 0 — C — 0
angle = 125.8°; C— C distance = 1.544, and C— 0 distance = 1.273)
leads to a value of 3.28 A. Intercharge distances for various dicarboxylates
and other compounds known to inhibit succinate dehydrogenase are given
in Table 1-1, and these values will be of interest in comparisons of inhibi-
tory activity. Dicarboxylic acids with more than one methylene group are
flexible and the intercharge distances may vary between the limits of great-
est bending and extension of the molecules. As the chain length increases
there will be greater tendency for the intercharge distance in the ions to
be less than that of the maximal extension, since the electrostatic repulsion
will decrease. The mean statistical intercharge distance in the succinate ion
will probably be closer to 4.75 A than the contracted distance of 3.81 A.
Indeed, it may be calculated that it would require at least 2.3 kcal/mole
to bring succinate from the extended to the contracted configuration,
using the dielectric constant obtained from D = 6d — 7 (Eq. 1-6-72)*;
since the dielectric constant is probably less due to the hydrocarbon groups
between the charges, this would be a minimal energy value. The mean
intercharge distance in the succinate ion may be estimated as not less than
4.20 A (Gane and Ingold, 1931; Eyring, 1932; Westheimer and Shookhoff,
1939) and possibly closer to 4.75 A. The glutarate intercharge distance is
probably around 5.2 A. These considerations are of importance in comparing
the interactions of these substances with the active center of succinate
dehydrogenase. It must be remembered that the bound ions are undoubt-
edly held in a configuration different from the statistical mean in solution.
The flexibility of the higher homologs allows them to adjust to a specified
configuration, but at the expense of the energy and entropy changes neces-
sary to bring them from their free configurations.
Acidic Ionization
The ionizations of malonic and succinic acids are important in the inter-
actions with succinate dehydrogenase and with regard to the penetration of
these substances into cells. The pKJs for the simple dicarboxylic acids, in-
cluding various derivatives of succinate and malonate, and related succinic
dehydrogenase inhibitors, are given in Table 1-2. The dissociation constants
change slightly with ionic strength; for malonic acid, dpK^ Ids = — 0.32,
and dpK^ jds = — 0.98, for ionic strengths around 0.15, where s is the
* In cross references of this type, Eq. 1-6-72, the roman number indicates the volume
of this treatise in which the equation, table, or figure (as the case may be) may be
found, the first arabic number indicates the chapter number, and the second arabic
jiumber the equation, table, or figure number.
1. MALONATE
Table 1-1
Intekchakge Distances in Dianions op Interest "
Compound
Oxalate
Malonate
Succinate
Glutarate
(c)
Adipate
Hydroxy malonate
(tartronate)
Fumarate
Maleate
Acetylene-dicar boxy late
o-Phthalate
Isophthalate
Terephthalate
Structure
'ooc-coo
'ooc coo"
\ /
CHj
"ooc -CHj
"OOC
CH,- COO"
COO"
\ /
CH2 CH2
"OOC— CHj
CH2 CH2
COO"
"OOC-CHj
"ooc coo"
/ \
CH9 CHa
^ /
CHj
"OOC-CHj
CHj— CH.
\
COO
CHj- COO"
"OOC COO
\ /
CH
I
OH
"OOC H
\ /
c=c
/ \ -
H COO
"OOC COO"
\ /
c = c
/ \
H H
"OOC-CEC-COO"
COO"
^^^^-^ COO-
COO"
Distance (A)
2.42
3.28
4.75
3.01
5.83
4.84
1.70
6.87
3.28
4.87
3.66
5.16
3.55
5.84
7.33
COO
CHEMICAL PROPERTIES
Table 1-1 (continued)
Compound
Structure
Distance (A)
Cyclobutane-dicar boxy late
Cyclopentane-dicar boxy late
COO
coo
COO'
Cyclohexane-dicarboxylate
COO'
COO'
COO
/3-Sulfopropionate <
(a)
(b)
OOC-CH,
\
CHj-SO.
"OOC SO,'
CHj— CHj
Methanedisulfonate
(methionate)
'O3S SO3'
CH,
1,2- Ethanedisulf onate <
(a)
(b)
'OjS-CHj
CHj— SO3'
'0,8 SO,'
' \ / '
CHj— CHj
/::?X^COO"
0 -Sulfobenzoate
^SO,'
Arsonoacetate
'OOC ASO3H'
CH,
^-Phosphonopropionate <
(a)
(b)
'OOC-CHj
"OOC
CH,— PO;
po:
\ /
CH2 CH2
3.39
cis-
3.52
trans-
4.91
cis -
3.05
trans -
4.51
5.05
3.13
3.40
5.38
3.25
3.72
4.05
5.09
3.14
The distances were calculated on the basis of bond lengths and angles given In Tables 1-6-12 and 1-6-13
except where direct measurements were available. It was assumed that the center of negative charge lies
midway between the resonating oxygen atoms. For o-phthalate and cyclohexane-dicarboxylate a 3° distortion
of the bond angles due to electrostatic repulsion was assumed. For cyclopentane-dicarboxylate a further
5° widening was estimated from the bending of the ring angles. The value for cyclobutane-dicarboxylate
is only approximate and is based on a 7° distortion of the bond angle in comparison to malonate. It should
be pointed out that the values in this table are smaller than generally used; one reason is that Intercarboxy-
late distances have usually been based on the distances between associating or dissociating protons , ra-
ther than between centers of negative charge.
ionic strength. Since AH for the dissociation of weak acids is quite small,
the constants do not change much with temperature; d^K^JdT = 0.0031,
and d])Kg jdT = 0.0038, approximately. Three species will be present in
any solution of malonate — HOOC— CHg— COOH, HOOC— CHg— C00-,
and -OOC — CHg — COO" — the relative concentrations being determined
1. MALONATE
Table 1-2
Ionization Constants of Dicarboxylic Acids "
Acid
pKa,
P^.,
Oxalic
Malonic
Succinic
Glutaric
Adipic
Pimelic
Methylmalonic
Ethylmalonic
Dimethylmalonic
Diethylmalonic
n.-Propylmalonic
i«o-Propylmalonic
Methylethylmalonic
Di-w-propylraalonic
Phenylmalonic
Hydroxymalonic (tartronic)
Maleic
Methylmaleic (citraconic)
Fumaric
Methylfumaric (mesaconic)
Malic
Tartaric
2,2 '-Dimethylsuccinic
2,2 '-Diethylsuccinic
Tetramethylsuccinic
Methylenesuccinic (itaconic)
Cyclopropane- 1 , 1 -dicarboxylic
Cyclobutane- 1 , 1 -dicarboxylic
Cyclopentane- 1 , 1 -dicarboxylic
Cyclohexane- 1 , 1 -dicarboxylic
<raw5-Cyclopropanedicarboxylic
cis-Cyclopropanedicarboxylic
frarw-Cyclopentanedicarboxylic
<raws-Cyclohexanedicarboxylic
Phthalic
Isophthalic
1.09
3.79
2.58
5.17
3.95
5.16
4.07
4.93
4.17
4.95
4.23
4.98
2.86
5.24
2.79
5.34
2.97
5.59
2.04
6.90
2.83
5.38
2.78
5.46
2.71
6.07
1.92
7.13
2.43
4.68
2.93
—
1.67
5.75
2.23
5.89
2.85
4.00
2.93
4.82
3.21
4.62
2.83
3.88
3.77
5.82
3.34
6.22
3.33
6.90
3.67
5.19
1.77
7.37
3.08
5.82
3.18
6.02
3.40
6.05
3.49
4.75
3.16
6.09
—
4.22
—
4.45
2.95
5.23
2.15
4.49
" These values have been obtained from a variety of sources and have been corrected
to a temperature of 37° and an ionic strength of 0.15 so as to be applicable to physio-
logical conditions. These corrections were obtained from studies on the variations of
dicarboxylic ionization constants with temperature and ionic strength (e.g. AdeU,
1940). They are not absolutely correct but probably allow a closer approximation
than the values in the literature, which are usually for 25° and extrapolated to zero
ionic strength.
CHEMICAL PROPERTIES
by the pH (see Eqs. 1-14-6 to 1-14-8). The variations of these species with
pH are shown in Fig. 1-1, and actual concentrations are given in Table 1-3
for malonate and succinate at a total concentration of 10 mM. In the
usual range of physiological pH, over 95% of these acids are in the form
of the completely dissociated doubly-charged anion; this is the active form
for the inhibition of succinate dehydrogenase. However, it is the concen-
trations of the other forms which are important in the rates and degrees
of penetration into cells, and these change appreciably with pH (e. g.,
from pH 7.4 to 6.8 there is a 4-fold increase in the singly dissociated form
and a 16-fold increase in the undissociated form).
"x
/^
,cooh\
^COOH \
^<
_ cooA
' COOH \
/ COO'
/ ^coo-
; ^
I
' Malonate
J
\
V
pH
Fig. 1-1. Concentrations of the various ionic species of malonic
acid at different values of the pH expressed as per cent as of
the total concentration.
Das and Ives (1961) on the basis of thermodynamic evidence suggested
that an internal symmetrical hydrogen bond occurs in H-malonate~, and
that this would affect to some extent the piiC^ values and reduce hydration.
However, Lloyd and Prince (1961) examined the infrared spectra of malonic
acid and its ions in DgO, compared the data with those obtained with
fumaric acid (in which no hydrogen bonds could occur), and concluded that
if hydrogen bonding exists in H-malonate~, the bond is very weak and not
symmetrical. Eberson and Wadsd (1963) determined the ionization enthal-
pies in water and ethanol, and also concluded that hydrogen bonding is
not important in the dicarboxylates when zlpiiC^ is less than 4. It would
thus appear that intramolecular hydrogen bonding is not a significant
factor in stabilizing the H-malonate~ ion.
10
1. MALONATE
>
ft
E-t
< ^
d d
o o
-H 05 -<
^ -^ oo
ft
CHEMICAL PROPERTIES 11
Metabolic studies of the substituted malonates and malonic esters wiU be
taken up after the actions of malonate have been discussed (see page 235).
It is interesting to note (Table 1-2) that the pK^ 's of the substituted
malonic acids are generally higher than for malonic acid itself. This is
mainly the result of the reduction of the dielectric constant of the region
between the interacting carboxyl groups, and is particularly evident for
the disubstituted ethyl and w-propyl derivatives. This increase in pK^
should facilitate penetration of these compounds into cells; in addition they
are more lipid-soluble, which will also favor penetration. The esters are not
active inhibitors of succinate dehydrogenase, at least by the same mechanism
as malonate, but have been used because of their ability to enter cells and
tissues readily, some hydrolysis to active malonate within the cells being
assumed. The presence of two keto groups on either side of the methylene
group makes this latter group more reactive and, indeed, imparts some
acidic character to it, malonic diethyl ester having a p^^ of approximately
5 X 10~^^ (Pearson and Mills, 1950). The rate of ionization is, however,
quite slow (A; = 1.8 X 10-^ min-^).
Chelation with Metal Cations
Malonate is able to form fairly stable complexes with various cations
normally present in media used in metabolic studies. The importance of
this in malonate inhibition will be discussed later (see page 66), and in the
present section we shall investigate the magnitudes of the effects expected.
These complexes are chelates with a six-membered ring structure, accounting
o or o. /O
for the relatively high stability compared to complexes with the monocar-
boxylates. The chelate dissociation constant is given by:
(M++) (A=)
K=- — - (1-1)
(MA)
where A= represents the anion of any dicarboxylic acid. The values for the
piii's of some of the more important chelates are given in Table 1-4. These
constants are dependent on temperature and ionic strength. The pK for
Mg-malonate is related to the temperature at zero ionic strength in the
following way: pZ = 2.92 - 0.008 (35 - «oC) (Evans and Monk, 1952).
Thus the pii's at 37° are approximately 0.1 unit higher than at 25°, the
temperature at which the constants are most commonly determined. It
was calculated from the data on the complexes of malonate with Mg++,
Ca++, Ba++, and Zn++ that the pK at an ionic strength of 0.15 is about
12
1. MALONATE
« >S
O
-M — I --
O cc -^
CI -^ — 1
(N -H rt
-H (M P-
c
2
c5
o
o
+s
P<
cS
3
3
T3
^
02
O
<Jl
P^
^ CS
a
c
CO
>>
S
^
ki
■^
'-^S
cS
43
fcT
3
O
i
^
V
.S
V
s
is
E3
3
CO
o
O
'53
CD
W
^
o
-1-i
§
ft
C
CO
M
g
S
a
c3
•r"
1
O
o
CO
a;
O
S
o
^
_C
to
o3
3
to
o
to
to
c
c3
C
o
_o
'-P
73
'-3
^
01
+3
eS
4^
^
s
S
«
OJ
s
V.
o
iH
o
e
o
o
s
o
o
o
1
a
c
ft
>.
_bi
J2
5b
'S
CO
[o
"c
c
eS
'm
o
o
C
c3
CO
73
CO
1
"s
P
<D
^
S
s
4^
,::— '
2
C
t4H
."'^
JS
o
o
II
-»^
o
^
o
+
+
§
3
03
to
3
aj
■ '
O
3
le
II
^
^
3
C
>
+3
S
a
>.
0;
'So
aj
^
J2
CO
j3
'T3
o
-t>
«
-ti
_s
Cm
C
O
o
CO
>
to
c
-C
O
S)
_o
03
CO
-C
'-5
»o
-tJ
S
C
CO
C
O
d
o
a
73
a)
C
o
o
■t^
C
(h
C
M
a
,o
_o
C
c«
"-3
2
m
w
2
(0
a>
'o
c3
CO
2
'S
J5
a;
73
.2
o
>
s
j;
'-3
c
-p
cS
eS
bl)
-i-i
H
C
e
TS
?>
13
C
3
cS
CO
a*
CHEMICAL PKOPERTIES 13
0.74 unit lower than at an ionic strength of zero. The free energy of forma-
tion of Mg-malonate is — 3.90 kcal/mole, while .1//° = 3.2 kcal/mole,
and J>S'o = 23.9 cal/degree (Evans and Monk, 1952; Chaberek and Martell,
1959, p. 139). The entropy term is large and probably results mainly from
displacement of water from the charged groups.
The importance of this chelation in inhibition work lies in the reductions
in the concentrations of the free ions it may bring about, both the metal
ions and malonate. The decrease in metal ion concentration can easily alter
enzyme activity or cellular function and such changes are apt to be attrib-
uted to a direct action of malonate. Conversely, the malonate concentra-
tion may be reduced appreciably. Examples of mutual concentration re-
duction are given in Table 1-5. The concentrations of Mg++ chosen approxi-
mate those in Tyrode solution (0.11 niM), Krebs-Ringer phosphate medium
(1.18 mM), the usual media for mitochondria (5 mM), and sea water (53.6
TCiM), while the concentrations of Ca"'""'" correspond to Krebs-Ringer phos-
phate medium (2.54 mM) and sea water (10.24 mM). It may be noted that
very significant reductions in Mg^^ and Ca++ can occur with concentrations
of malonate commonly used; e.g., malonate at 10 mM will reduce the Mg++
49% and the Ca++ 23% in Krebs-Ringer medium, and higher concentra-
tions may almost deplete these ions from the solution. When the concen-
trations of these cations are high, as in sea water, the effective malonate
concentration may be reduced markedly; e. g. malonate added to sea water
at a total concentration of 10 mM will result in a 1.5 mM solution of free
malonate ion. Such phenomena have usually been ignored or forgotten
despite their possibly large magnitudes. One way of determining the impor-
tance of cation reduction in malonate studies is to calculate the reduction
to be expected in the medium used and at the malonate concentration, and
then to test the effects of lowering the cation concentration to this extent
(Rice and Berman, 1961). Other metal cations may be reduced to a greater
extent than Mg++ and Ca++. Media initially 1 mM in Co++, Mn++, or Cu++
will in the presence of 5 mM malonate contain these ions at concentrations
of 0.65 mM, 0.34 mM, and 0.0052 mM, respectively. If such metal ions are
normally bound to enzymes, the degree of removal from the enzyme will,
of course, depend on the relative affinities of the metal ion for the enzyme
and malonate. There is also the possibility that malonate may chelate with
metal ions combined with the enzyme, inactivating them for their catalytic
role. It should also be remembered that similar phenomena may occur with
succinate, and the inhibition kinetics may be distorted when malonate is
used due to the differential reduction in the concentrations of these anions.
Detection and Determination of Malonate
Methods have been developed for the separation and identification of
organic acids from animal and plant tissues. Earlier determinations involved
14
1. MALONATE
r^ q
K
,— V
c
lO
o
CO
CO
t^
CD
00
LCl
(M
c<l
o
+*
o
o
r-
00
o:
in
LO
+
O
g
d
d
CO
t^
d
C5
Vm
O
o
'^
lO
o
CO
CO
CO
Oi
o
+
t-
CD
GO
LO
l-H
(^
+
ai
t-
00
OJ
i-O
^_
lO
"-3
O
05
05
00
r^
CD
■*
(M
cS
;-i
-fj
c
0)
o
t^
C3
g
<M
CD
c-
t-
O
o
^
Is
05
t^
CD
o
CO
LO
CD
o
GO
CD
Tt<
o
Tt<
O
„_
g
o
d
Tji
d
d
GO
oo
o3
g
-*
^
Tt<
05
'Si
lO
°3
C<l
+
CO
CD
r^
t^
i?q
CI
o
o
+
fO
r-H
o
Tt<
t^
00
o
cS
lO
Tt<
(M
05
LO
C5
CD
O
c^
■M
Csl
-'
-<
d
d
II
lO
g
c
CD
lO
t-
o
'N
_o
CD
t^
C5
'M
CD
CD
^
Is
O
d
d
00
d
00
^
^
o
CO
(M
t^
j;^
^^-
-M
»c
_o
o
^_^
'-3
IC
+
+
'M
CD
00
Ci
(M
CD
CD
h
Ci
GO
00
"*
O
(M
"S
^
Ol
C5
O'
_h'
tH
CO
t^
Tl*
^
-<*
'*
CO
'>^
o
o
"ip
c
o
t^
t^
'^^
CO
^
CC
t^
•*
GO
Tt<
00
CD
,o
+'
o
CD
CD
t~
00
00
Tj<
Qi)
+
SB
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INHIBITION OF SUCCINATE DEHYDROGENASE 15
oxidative titrations with hot acid permanganate or eerie sulfate (Willard
and Young, 1930; Christensen and Ross, 1941) and such methods have
been used for the analysis of malonate in the presence of proteins (Ross and
Green, 1941). Colorimetric tests, for example with tetrahydroquinoline-iV^-
propenal to form blue- violet compounds (sensitive to 0.01 mg malonic acid)
(Dieterle and Wenzel, 1944), have been used, and several microcolorimetric
and spot tests are available (Feigl, 1960). However, the most valuable
methods are chromatographic. A large variety of solvents and spraying
agents have been utihzed, and different techniques have been applied:
ion-exchange resin columns and partition chromatography (Stark et al., 1951;
Phares et al, 1952; Owens et al, 1953; Shkol'nik, 1954; Reinbothe, 1957),
strip paper chromatography, both one- and two-dimensional (Buch et al.,
1952; Cheftel et al, 1952; Denison and Phares, 1952; Duperon, 1956; van
Duuren, 1953; Jermstad and Jensen, 1950; Kalyankar et al, 1952; Ladd
and Nossal, 1954; Overell. 1952; Smith and Spriestersbach, 1954), and cir-
cular paper chromatography (Airan and Barnabas, 1953; Airan et al, 1953;
Barnabas, 1955). The use of chromatographic methods for the determina-
tion of succinate and malonate in animal tissues is well illustrated in the
work of Busch and Potter (1952 a, b; Busch et al., 1952). A moderately sen-
sitive method for estimating 10-100 //g of malonate by applying iodine
vapor to paper chromatograms was developed by Dittrich (1963).
INHIBITION OF SUCCINATE DEHYDROGENASE
The oxidation of succinate in cells is catalyzed by a mitochondrial
multienzyme system which is structurally organized into units that can be
isolated as a particulate suspension capable of transferring electrons from
succinate to oxygen: this complex is known as succinate oxidase. Evidence
will be presented that the inhibition of this system by malonate is related
to the binding of the malonate to the most proximal site in the sequence,
namely, the active center for the attachment and dehydrogenation of succi-
nate: this component is known as succinate dehydrogenase. It is impossible
at the present time to define the succinate dehydrogenase accurately, be-
cause it is assayed with various electron-accepting dyes and the basic mini-
mal unit has not been completely characterized. The inhibition of succinate
dehydrogenase by malonate has generally been determined with various
preparations of succinate oxidase and not with the isolated dehydrogenase,
and thus it is necessary to discuss briefly the nature of the entire system.
Properties of Succinate Oxidase
Knowledge of the components of succinate oxidase and the pathways of
electron flow has been advanced markedly in the past few years (Mahler,
16 1. MALONATE
1956; Singer, et al., 1957; Green and Crane, 1958; Singer and Lara, 1957;
Green and Flieseher 1960; Green, 1960; Redfearn, 1960). The particulate elec-
tron transport particle (ETP) of Green, obtained from heart mitochondria, is
a succinate oxidase preparation and has been shown to contain the following
components (per molecular weight of about 5 x 10^): flavin dinucleotide 2,
nonheme iron 64, heme (equal amounts of cytochrome a, cytochrome
b, and cytochromes c + Cj) 6, copper 8, ubiquinone (coenzyme Q) 10, and
lipid 34,5%. In addition, there are the several proteins with which these
substances are bound. Such preparations also oxidize NADH, ascorbate,
p-phenylenediamine, and hydroquinones, these substrates supplying elec-
trons at various sites in the electron transport chain. The use of various
electron donors and acceptors, the application of specific inhibitors, and the
fragmentation of the succinate oxidase complex have led to several postu-
lates of the pathways of electron flow. One difficulty is the probable dif-
ference in pathways between mitochondrial phosphorylating systems and
submitochondrial nonphosphorylating preparations. A second difficulty is
the possibility of alternate pathways of electron flow rather than a single lin-
ear sequence. Scheme 1-2, (Green, 1960; Redfearn, 1960) might be assumed
provisionally:
UQ UQ
cs . i'«/ ^ ^^ ^ ^^f,|
Succinate »- ASF ^ -• NADH
iFe) \^ /^ I >\ ^ lFe\
(1-2)
Fe represents nonheme iron, f, is the flavoprotein associated with succinate
dehydrogenase, f^ is the flavoprotein associated with NADH dehydrogenase,
UQ is ubiquinone (coenzyme Q), ASF is the antimycin-sensitive factor, and
the other symbols indicate the usual cytochromes. It is possible that cyto-
chrome b and ubiquinone are common to the two pathways from succinate
and NADH, but Green believes the evidence points to fusion of the chains
INHIBITION OF SUCCINATE DEHYDROGENASE 17
only at the antimycin-sensitive factor. It is also possible that ubiquinone
and cytochrome b are on a linear pathway rather than on alternate path-
ways. In nonphosphorylating systems, the role of cytochrome b is debat-
able. The sites of oxidation of p-phenylenediamine and ascorbate are distal
to ASF. It should also be pointed out that succinate oxidase from other
sources may have different ratios of components and somewhat different
pathways. Thus the oxidation of succinate and NADH by the electron
transport system of Azotobacter is not sensitive to antimycin (Bruemmer et
al., 1957). Two recent observations have extended our knowledge of suc-
cinate oxidase and related pathways. Azzone and Ernster (1961) have
demonstrated an ATP requirement for mitochondrial phosphorylating
succinate oxidation and propose reactions such as the following:
Succinate + A -l ATP -> fumarate + AH ~ P + ADP
AH ~ P + B + ADP -> A + BH2 + ATP
Succinate + B -> BHj + fumarate
A is possibly a flavoprotein, a quinone, or some other component of succinate
dehydrogenase, and B may be cytochrome b. In nonphosphorylating systems
the electrons may reach B more directly. An alternate pathway for AH -^ P
is the reduction of NAD:
AH ~ P + NAD -> A + NADH + P^
Succinate has been shown to reduce NAD in mitochondria by Chance and
Hollunger (1961 a, b, c) and this is dependent on ATP. Since cndogen^asly
formed succinate is more effective than added succinate, it is possible that
this reaction may involve succinyl-CoA. The reduction of NAD passes
through antimycin-sensitive and Amytal-sensitive links and seems to involve
a third flavoprotein component. These findings are not only important with
respect to the behavior of succinate oxidase, but also have possible bearing
on the responses to malonate in mitochondrial and cellular systems.
The succinate oxidase particles have been fragmented in various ways to
yield smaller particles or soluble preparations with different compositions
and properties. One such preparation is the succinate dehydrogenase complex
(SDC) from heart mitochondria or ETP, which perhaps represents that frac-
tion of the complex up to cytochrome c, since it oxidizes both succinate and
NADH and possesses an antimycin-sensitive step. Of more interest for
malonate inhibition are the several forms of soluble succinate dehydrogenase
that have been prepared. The purest contain no heme or lipid; four atoms
of tightly bound nonheme iron and one flavin occur in a molecule (assuming
a molecular weight of around 200,000). The flavin is apparently not ribofla-
vin but occurs in a dinucleotide form covalently attached to peptide chains
of the apoenzyme (Kearney, 1960). There is spectral evidence that both
18 1. MALONATE
flavin and nonheme iron are reduced by succinate, and the functional role
of iron in the catalysis is further indicated by the inhibition produced by
complexing the iron with 1,10-phenanthroline or /J^-globulin (Singer et al.,
1957). Electron spin resonance studies of succinate oxidase have demonstrat-
ed signals when succinate is added; these free radicals seem to be associat-
ed with the dehydrogenase and possibly reflect changes in the states of
iron or flavin (Commoner and Hollocher, 1960; Hollocher and Commoner,
1960). The high sensitivity of succinate dehydrogenase to most sulfhydryl
reagents indicates the presence of an SH group at or near the succinate-
binding site. One may, therefore, characterize succinate dehydrogenase
from our present knowledge as containing two cationic groups for the
binding of succinate, an SH group nearby, some nonheme iron, and a unique
flavin dinucleotide in a tight peptide complex.
Relationship of Malonate Inhibition to the Electron Acceptor Used
Measurement of malonate inhibition involves either the oxygen uptake of
the complete succinate oxidase system, or the determination spectroscopi-
cally of the reduction of one of the normal components (such as cytochrome
c), or the reduction of some artificial electron-acceptor dye. The complete
system can reduce a variety of substances in the presence of succinate, and
malonate has been shown to inhibit such reductions whatever the acceptor
used: methylene blue (Quastel and Whetham, 1925; Hopkins et al., 1938;
Forssman, 1941; Franke 1944 a; Kaltenbach and Kalnitsky, 1951 a; Wad-
kins and Mills, 1955), ferricyanide (Stoppani, 1948; Thorn, 1953), tetrazolium
dyes (Barker, 1953; Becker and Rauschke, 1951; Zollner and Rothemund
1954; Waterhouse, 1955), manganese dioxide (Hochster and Quastel, 1952),
2,6-dichlorophenolindophenol (Repaske, 1954; Wadkins and Mills, 1955,
Millerd, 1951), janus red (Agosin and von Brand, 1955), brilliant cresyl
blue (Agosin and von Brand, 1955), and lY-methylphenazine sulfate (Singer
et al., 1956 b). Inhibition of cytochrome c reduction has also been observed
Seaman, 1954). There is thus substantial evidence that malonate blocks
electron flow very early in the sequence, as would be expected if it prevents
the binding of succinate to the dehydrogenase. The most proximal location
of the site of malonate inhibition comes from the work of Ziegler (1961) on
electron transport particles from heart mitochondria. Some of the nonheme
iron is reduced by succinate and this is blocked by 20 raM malonate. It
may also be noted that succinate reduces NAD and NADP in submito-
chondrial particles from heart through an ATP-dependent system and this
is readily inhibited by malonate (Snoswell, 1962; Hommes, 1963; Lee et al.,
1964), which possibly indicates that succinate dehydrogenase is involved.
The addition of malonate to succinate dehydrogenase brings about changes
in the absorption spectrum in the flavin region, as do succinate, fumarate,
and other competitive inhibitors (Dervartanian and Veeger, 1962). There is
INHIBITION OF SUCCINATE DEHYDROGENASE 19
a decrease in absorption between 400 and 470 m// and an increase between
480 and 540 m//, with a maximum at 510 m// in the difference spectrum.
It is not known what this implies relative to the site of malonate binding.
The dyes may accept electrons from various sites in the succinate oxi-
dase system. The fragments obtained from the oxidase particles differ in
their abilities to reduce these dyes, and as one approaches the purest succi-
nate dehydrogenase the number of possible electron acceptors is reduced.
Indeed, soluble succinate dehydrogenase reduces only the A^-alkylphenazines
and ferricyanide at appreciable rates. Some dyes accept electrons chiefly
from the nonheme iron, some accept more efficiently from the flavin com-
ponent, and some, such as the indophenol dyes, involve ubiquinone. It may
well be that none of the commonly used dyes is completely selective. Fer-
ricyanide, for example, may react at other sites down the chain in addition
to the nonheme iron, since its reduction has a partially antimycin-sensitive
component, and even the A^-alkylphenazines, considered to be the most
reliable acceptors at the dehydrogenase level, may be able to react at other
sites.
The question of importance with respect to malonate inhibition is whether
all of the methods of measuring succinate dehydrogenase activity are equiva-
lent for the purpose of obtaining accurate kinetic results. Unfortunately,
there have been very few reliable investigations wherein different methods
or acceptors have been compared. The inhibitions of succinate oxidase
(manometric) and succinate dehydrogenase (dye acceptors) at the same
concentrations of succinate and malonate have been reported sporadically.
It has generally been found that the oxidase is inhibited somewhat more
strongly: Arum spadix (Simon, 1957), Limidus gill cartilage (Person and
Fine, 1959), and potato tubers (Millerd, 1951). But in the enzymes from
Xanthomonas, this is reversed (Madsen, 1960). The results of Millerd are
particularly difficult to understand, inasmuch as she found a 40% inhibi-
tion of the oxidase at 0.1 raM malonate whereas the dehydrogenase (using
2,6-dichlorophenolindophenol as acceptor) was not inhibited at all. The
author is not aware of any quantitative studies comparing malonate inhi-
bition with different dye acceptors. It would appear that most workers
have assumed there would be no difference.
Actually, from kinetic consideration, it is not at all necessary that the
inhibition produced by a chosen concentration of malonate be the same
when different acceptors are used. In fact, the inhibition may depend on the
acceptor concentration. Thorn (1953) showed that the inhibition of pig heart
succinate dehj^drogenase by 0.536 mM malonate increases with the con-
centration of methylene blue: at 0.15 n\M methylene blue the inhibition
is 15.9% and at 3 vaM methylene blue it is 28.8%. Succinate oxidase, is
inhibited 52.6% at the same malonate concentration. The transfer of hydro-
gen atoms from succinate to a dye acceptor always involves the interaction
20 1. MALONATE
of these two substances with the enzyme surface, undoubtedly at different
sites. At low concentrations of the acceptor, or with weak acceptors, the
over-all rate may not be determined by the rate at which the hydrogen atoms
are removed from the succinate, but may depend also on the rate of transfer
to the acceptor. The malonate inhibition will thus vary with the degree of
saturation of the enzyme with acceptor. On this basis it would seem reason-
able to use those acceptors which are the most active and react with
sites closest to the succinate site. A further consideration is the inhibition
produced by the acceptors themselves. Both ferricyanide and iV-methylphe-
nazine begin to inhibit succinate dehydrogenase as the concentration is
raised above certain levels. Such systems would then constitute examples
of multiple inhibition and the kinetics of the inhibition due to malonate
alone may be distorted. The choice of the acceptor and its concentration
is thus of some significance.
Site of Inhibition by Malonate in the Succinate Oxidase Sequence
The results discussed in the preceding section point clearly to the site
of inhibition as succinate dehydrogenase. Indeed, inhibition of soluble
succinate dehydrogenase by malonate has been demonstrated. The competi-
tive nature of the inhibition, to be treated in the following section, indicates
the inhibition to be at the active site at which succinate is bound. There is
thus no question but that the major site of inhibition is at the very begin-
ning of the electron transport sequence in succinate oxidase. The question
that now must be considered is whether malonate can inhibit at any other
step of the electron transport chain.
There are two obvious ways to examine this. One is to test the action
of malonate on the succinate oxidase system, using substrates that donate
electrons at more distal sites than succinate. The other way is to determine
the response of other oxidases that utilize most of the electron carriers
in the succinate oxidase. Quastel and Wheatley (1931) observed that ma-
lonate at 67 mM does not inhibit the oxidation of p-phenylenediamine
and hence concluded that the cytochrome region of the sequence is immune
in their preparations. Actually, many oxidases, comprising varying segments
of the electron transport chain, have been found to be insensitive to malo-
nate at concentrations from 25 mlf to 50 mM. All this evidence points to
a rather specific action on the dehydrogenase. However, there are data in
the literature which indicate that, at least in some species and at high enough
malonate concentrations, inhibition at other sites may occur. Although 30
mM malonate does not inhibit NADH oxidation in beet mitochondria
(Wiskich et al., 1960), some inhibition has been observed in mosquito parti-
cles (10-15% at 1-10 mM) (Gonda et al, 1957) and in Tetrahymena NADH
oxidase (35% at 6.2 mM and 70% at 18 mM) (Eichel, 1959). Such inhi-
bition could, of course, be on the NADH dehydrogenase rather than on
INHIBITION OF SUCCINATE DEHYDROGENASE
21
enzymes common to the succinate pathway. Malonate does not usually
interfere with ascorbate oxidation, but in the silkworm it was found both
spectroscopically and manometrically that malonate inhibits the oxida-
tions of succinate and ascorbate equally (Sanborn and Williams, 1950).
Observations such as these, coupled with those showing greater inhibition
of succinate oxidase compared to the dehydrogenase, make it necessary
to exert some caution in assuming a completely specific action in all cases.
Competitive Nature of the Inhibition
It has been often stated that competitive inhibition was first demonstrat-
ed by Quastel and Wooldridge (1928) for the inhibition of E. coli succinate
dehydrogenase by malonate. However, inhibitions were not calculated
and the data presented do not lend themselves to quantitative interpreta-
tion. Indeed, when their data are plotted on a l/w, — 1/(S) graph (Fig. 1-2) a
straight line is not obtained. Since no comparable control studies were done
in the absence of malonate, the fact alone that increasing the succinate con-
centration increases the rate in the presence of malonate does not prove
competition. These points are brought out not to criticize pioneering work
but to illustrate that conclusions about the type of inhibition cannot be
made so readily as many imagine.
Fig. 1-2. A double reciprocal plot for the
inhibition of E. coli succinate dehydrogenase
by malonate at 1.43 mM. Succinate concen-
trations are in vaM. (Data from Quastel and
Wooldridge, 1928).
22
1. MALONATE
Competition between succinate and malonate has been claimed to occur in
the following: rat liver homogenates (Potter and DuBois, 1943), oyster
muscle homogenates (Humphrey, 1947), oyster egg homogenates (Cleland,
1949), carrot root (Hanly et al, 1952), yeast (Krebs et al, 1952), pig heart
particulates (Thorn, 1953), cockroach muscle homogenates (Harvey and
Beck, 1953), the trypanosome Crithidia (Hunter, 1960), and the soluble
succinate dehydrogenase from heart (Keilin and King, 1960). In all cases,
the inhibition produced by a certain malonate concentration is reduced by
increasing the succinate concentration, but a quantitative analysis of the
data has been seldom carried out. Most of these studies were made with
the usual assay methods, but competition has recently been shown by meas-
uring the reductions in the electron spin resonance signals produced by
malonate at different succinate concentrations (Commoner and Hollocher,
1960).
Fig. 1-3. A single-curve plot for the inhibition of
cockroach muscle succinate oxidase by malonate at
0.33 mM. Ki = 0.105 mM and KJK^ = 275. (Data
from Harvey and Beck, 1953).
Single-curve plots (type F, see Chapter 1-5) were made for two of the
inhibitions mentioned above (Figs. 1-3 and 1-4). In the case of Crithidia,
Hunter showed competition with a 1/'U-1/(S) plot and the single-curve plot
confirms this, K^ having the value of 0.22 mM and K^^JK^ calculated from
the slope a value of 53. The results with cockroach muscle likewise fall
roughly on a straight line, giving K, as 0.11 mM and KJK^ as 275, values
differing somewhat from those calculated by Harvey and Beck using a
INHIBITION OF SUCCINATE DEHYDROGENASE
23
different plotting procedure {K^ = 0.13 mM, and KJK^ = 200). In most
studies insufficient data are available for plotting. It must be emphasized
that a change in inhibition observed at two or three succinate concentra-
tions is not adequate to prove a purely competitive inhibition. It seems
to be rarely considered that an inhibition may not be either completely
competitive or completely noncompetitive. The conditions for partially
competitive inhibition were given (Eqs. 1-3-14 and 1-3-15) and the types
of plot to be expected discussed (Chapter 1-5). An interesting example
of this is provided by the work of Honda and Muenster (1961) on the inhi-
bition of succinate oxidation in lupine mitochondria. Here the osmolarity
of the preparation and assay media was varied with sucrose, and it was
found that the interaction constant, a, defined in Eqs. 1-3-5 and 1-3-6,
Fig. 1-4. A single-curve plot for the inhibition of
Crithidia succinate dehydrogenase by malonate at
1 mM. Ki = 0.22 mM, and KJE^ = .53. (Data from
Hunter, 1960).
varies quite markedly from values indicating nearly completely competitive
inhibition to those showing noncompetitive inhibition. This work will be
discussed in greater detail in a later section (see page 46), but it suffices
to show that partial competitive inhibition by malonate is possible and that
the type of inhibition may vary with the experimental conditions.
The most elegant treatment of malonate inhibition is by Thorn (1953)
at the St. Thomas's Hospital Medical School in London, using succinate
oxidase preparations from pig heart muscle. The activity was measured by
the reduction of ferricyanide in the presence of cyanide to block the cyto-
24
1. MALONATE
chrome pathway. The usual Ijv - 1/(S) plots (Fig. 1-5) show apparently
completely competitive inhibition, and values of the substrate and inhibi-
tor constants, to be discussed in the next section, were calculated. Using
average values of K„^ and K^ obtained by Thorn, the inhibition curves in
(Malonate) = 0 0089mM
No Malonate
Fig. 1-5. Double reciprocal plots for the inhibition
of pig heart succinate dehydrogenase by malonate,
showing pure competitive inhibition. Succinate con-
centrations are in mM and v in spectrophotometric
units. (Data from Thorn, 1953).
Fig. 1-6 were plotted. These curves show the expected reduction in inhi-
bition as the succinate concentration is increased at constant values of
malonate concentration. It may be noted that overcoming the inhibition
is much more difficult at high inhibitor concentrations and this must be
taken into account in experiments designed to show a competitive type of
action. K„i was used rather than K,. because under the usual experimental
conditions it is this constant that determines the behavior.
A word must now be said about malonate reversibility. One of the cri-
teria of competitive inhibition is that the inhibitor should leave the active
site readily when its concentration is reduced, or that it should be displaced
rapidly when more substrate is added. These points have seldom been
INHIBITION OF SUCCINATE DEHYDROGENASE
25
tested with malonate. However, it was demonstrated quite early that the
inhibition of rabbit muscle succinate dehydrogenase is reversible by washing;
the preparation was incubated for 30 min. with 100 milf malonate anaerobi-
cally and then washed 3 times on a filter — the reduction times were 15
min for the control, 61 min with the malonate, and 17 min for the washed
preparation (Hopkins et al., 1938). A trypanosomal succinate dehydroge-
nase, however, showed no reversal of the inhibition, even when succinate
was added to 50 times the malonate concentration, which is particularly
surprising since in the living cells a good reversal was observed (Agosin and
von Brand, 1955). The concentrations of malonate used here were not
unduly high and so these data are unexplainable.
Fig. 1-6. Curves showing the calculated reductions
of the malonate inhibition of succinate dehydrogen-
ase by various concentrations of succinate, using
K„ = 0.366 mM, and K^ = 0.0076 mM, as given
by Thorn (1953).
Constants of the Inhibition
It is not surprising that the values of the inhibitor constant, K^, for
malonate inhibition are quite variable in the literature, because not only
do the experimental conditions affect this constant markedly but the suc-
cinate dehydrogenase varies in its properties from species to species. From
the values in the accompanying tabulation and from reasonable estimates
based on assumed or determined substrate constants for the data in Table
1-6, it is seen that the K^'s for most preparations vary between 0.005 mM
26
1. MALONATE
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INHIBITION OF SUCCINATE DEHYDROGENASE
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INHIBITION OF SUCCINATE DEHYDROGENASE 29
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INHIBITION OF SUCCINATE DEHYDROGENASE
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INHIBITION OF SUCCINATE DEHYDROGENASE
33
Preparation
Ki {mM)
Reference
Pig heart
0.0076
Rat heart
0.01
Yeast
0.0105
Ehizobium japonicum
0.017
Claviceps purpurea
0.03
Corynebacterium diphtheriae
0.037
Beef heart
0.041
Beef heart
0.045
Mytilus edulis
0.06
Beef heart
0.13
Micrococcus lactilyticus
0.23
Phaseolus vulgaris
0.24
Lupinus albus
0.91
Thorn (1953)
ITyhn and Matsumoto (1964)
Ryan and King (1962a)
Cheniae and Evans (1959)
McDonald et al. (1963)
Strauss and Jann (1956)
Kearney (1957)
Keihn and King (1960)
Ryan and King (1962 a)
Lee et al. (1964)
Warringa and Giuditta (1958)
Hiatt (1961)
Honda and Muenster (1961)
and 0.05 vaM, and mammalian tissues generally yield quite low constants.
The interest in the K, lies in its relationship to the binding energy of mal-
onate to the active center, so that variations in the K^, unless due to exper-
imental conditions, may be attributed to differences in the topography
of the enzyme surface. The values of £",„ for different preparations are not
particularly significant, except for characterizing a certain preparation
under specified conditions, because .K",,, does not usually equal K^.. Inasmuch
as the ratio of K^ to K j is of some significance in interpreting interactions
with the active center, it will be worthwhile to discuss the only instance in
which this has been determined.
The ratio, K„,!K„ has been given by various investigators as ranging
from 10 to 60. Thorn (1953) pointed out that K,„ for succinate dehydroge-
nase is often quite different than K, (Slater and Bonner, 1952). K,„ thus
equals (A-j + A'2)/A:j, and A-j is dependent on the experimental conditions.
Thorn obtained values of KJK^ from 3 to 60 by varying the electron-
acceptor dye and the reaction rate. The faster the rate, the greater the de-
viation of ^,„ from K^. Thus extrapolation to zero rate from a series of
experiments at different methylene blue concentrations enabled Thorn to
determine the true ratio of succinate and malonate affinities; KJK, turned
out to be 3. Since K, is approximately 0.0076 milf . K, = 0.023 mM. a value
which checks well with directly determined values of the rate constants
(A'i = 3.35x 10^, and A-_i = 0.99). Similar low ratios would probably be found
in most succinate dehydrogenase preparations. The difference in binding
between succinate and malonate is thus not so great as previously believed.
It is of great practical importance to realize that the degree of inhi-
bition of succinate oxidase by malonate varies with the rate of succinate
oxidation and the electron acceptors present. This may explain some of the
34 1. MALONATE
differences in Table 1-6. If one is to compare the inhibitory potencies of
malonate on succinate oxidases from different tissues or species, equivalent
rates of succinate oxidation should be used and similar methods of deter-
mining the rate should be employed.
Inhibition of Succinate Dehydrogenase by Other Dicarboxylates
Before discussing the more intimate nature of the inhibition and the
possible ways by which malonate is bound to the active center, it will be
useful to consider the inhibitory potencies of other dicarboxylate ions.
The configuration of an active center may often be approached by comparing
the relative affinities of analogous compounds for the enzyme. At the end of
this chapter a more complete discussion of inhibitors related to malonate
will be given; for the present we shall be interested only in the inhibition
of succinate dehydrogenase. Inhibitions and inhibitor constants are summa-
rized in Tables 1-7 and 1-8.
(a) U nsubstituted dicarboxylate ions. It was stated by Quastel and Whet-
ham (1925) that oxalate, glutarate, and adipate do not interfere with
succinate oxidation, in contrast to malonate, and, although in later work
they found some inhibition, this has generally been confirmed. Accurate
comparisons must be made using inhibitor constants, and these are not
available, but there is no doubt that in the series ~00C — (CHa)^ — C00~
the inhibition reaches a sharp maximum at w = 1. One must assume that
malonate best fits the intercationic distance on the active center of suc-
cinate dehydrogenase, and this includes succinate itself since K^ is generally
larger than K, for malonate.
(6) Unsaturated dicarboxylate ions. It is interesting to compare the two
isomers, fumarate and maleate, since they differ in the intercarboxylate
distance (Table 1-1). For mammalian succinate dehydrogenase it would
appear that fumarate is bound much less tightly than either succinate or
malonate (Table 1-8), but this does not necessarily hold for the bacterial
enzymes, providing evidence that the active center configurations may be
quite different in different dehydrogenases. One might expect a rather low
affinity for fumarate because of the fairly long intercarboxylate distance,
but it is surprising that maleate is such a poor inhibitor inasmuch as its inter-
carboxylate distance in only 0.38 A greater than in malonate. A complicating
factor is the ability of maleate to react with SH groups, and actually the
inhibitions observed by Hopkins et al. (1938) and Morgan and Friedmann
(1938 b) were obtained only after prolonged incubation. The possible
significance of these observations will be discussed in the next section.
On the other hand, acetylene-dicarboxylate, with a distance of 5.16 A
between carboxylate groups and a restriction to linearity, does inhibit and
this is completely competitive (Thomson, 1959).
INHIBITION OF SUCCINATE DEHYDROGENASE
35
P5
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1. MALONATE
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INHIBITION OF SUCCINATE DEHYDROGENASE 37
^ lO
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38
1. MALONATE
Table 1-8
Inhibitor Constants for Dicarboxylate Ions on Succinate Dehydrogenase
Source of enzyme
Inliibitor
Ki (mM)
Reference
Corynebacterium
Malonate
0.037
Strauss and Jann
diphtheriae
Fumarate
1.8
(1956)
Micrococcus lactilyticus
Malonate
0.23
Warringa and Giix-
Fumarate
0.22
ditta (1958)
Claviceps purpurea
Malonate
0.03
McDonald et al.
(ergot)
Fumarate
0.93
(1963)
Yeast
Malonate
0.0105
Ryan and King
Fumarate
1.03
(1962 a)
Mytilus edulis (mussel)
Malonate
0.06
Ryan and King
Fumarate
0.15
(1962 a)
Rat kidney
Oxalacetate
0.0015
Pardee and Potter
(1948)
Acetylene-dicar-
0.81
Thomson (1959)
boxylate
Pig heart
Malonate
0.05
Hellerman et al.
Fumarate
3.5
(1960)
Oxalacetate
0.0016
Ethyloxalacetate
0.04
Diethyloxalacetate
No inh.
Acetylene-dicar-
1.4
boxylate
ci5-Cyclohexane-
132
1,2-dicarboxylate
Beef heart
Malonate
0.041
Kearney (1957)
Fumarate
1.9
Itaconate
1.8
Dervartanian and
Veeger (1962)
(c) Oxalacetate. The marked inhibition exerted by this substance was first
noted by Das (1937 b) and this has been confirmed on the succinate dehydro-
genases from a variety of organisms. It is at present recognized as the most
potent succinate dehydrogenase inhibitor among the dicarboxylate ions.
The binding of the oxalacetate to the enzyme would appear to be at the
active center because the inhibition is competitive with succinate (Pardee
and Potter, 1948; Kearney and Singer, 1956; Hellerman et al., 1960), and
the presence of oxalacetate on the enzyme protects the active center from
various sulfhydryl reagents (Stoppani and Brignone, 1957). The reason for
INHIBITION OF SUCCINATE DEHYDROGENASE 39
this strong inhibition is still not clear but Hellerman and his group have
advanced some interesting speculations. Oxalacetate in aqueous solution
may exist around neutrality in three forms in equilibrium. It was suggested
that the enolate form may be the potent inhibitor, especially the trans-
'CX)C coo" ^ "OOC^ H
^c=c'^ -* »^ 'ooc-c-CH,— coo" -* ^ ,c=c^
-Q^ ^n 'o^ ^coo"
cJs -Enolate Keto tautomer trans -Enolate
tautomer tautomer
enolate tautomer where the enolate group and the carboxylate group are
on the same side. Evidence for this was provided by showing that the
oxalacetate monoethyl ester is inhibitory (about as potent as malonate)
EtOCO^ H
^C = C^
~o^ ^coo'
^raws- Enolate
/ tautomer of
monoethyl ester
(Table 1-8). The diethyl ester in inactive. Actually there is a difference in
binding energy of about 2.1 kcal/mole between oxalacetate and either
malonate or the monoethyl ester. This extra energy might be due to the
doubly negative charge on one end of the molecule. It could well be that
both the cis- and ^ra ws-enolate tautomers are bound, the charge distri-
butions being almost equivalent if one takes the center of negative charge
as lying between the carboxylate and enolate groups. It is unfortunate that
the per cent of the oxalacetate in the enolate forms at pH 7.6 and 30° in
aqueous solution is not known. Another possibility is that the keto tautomer
combines with the enzyme, the extra binding energy arising from an inter-
action between the keto group and the enzyme; a hydrogen bond could ac-
count for the 2.8 kcal/mole difference between the binding energies of ox-
alacetate and succinate. One difficulty in assuming the enolate form as the
active inhibitor is the fact that maleate inhibits very poorly and fumarate
not a great deal better. The difference in binding energy between oxalace-
tate and fumarate for the pig heart succinate dehydrogenase is close to
4.8 kcal/mole, and it would be difficult to account for this large difference
simply on the basis of additional ionic interactions. A final possibility must
be entertained, although no evidence for it exists, namely, the complexing
of the enolate group with one of the nonheme iron atoms near the cationic
groups of the enzyme surface, since this could provide the extra energy
for the binding of oxalacetate. It should be noted that although oxalacetate
strongly inhibits succinate oxidation in rapidly respiring liver mitochondria,
both coupled and uncoupled, it stimulates when the mitochondria are in a
40 1. MALONATE
state of respiratory control (Kunz, 1963). Malonate does not stimulate
under these conditions and thus the effects of oxalacetate on intact mito-
chondrial succinate oxidation differ in some manner from those of malonate.
This, however, is probably explained by other reactions of oxalacetate, e.g.,
the oxidation of NADH to NAD, which would release the mitochondria
from respiratory control.
{d) Substituted malonates and succinates. Franke (1944 a) found that alkyl-
malonates are inactive on heart succinate dehydrogenase, confirming the
weak inhibitions reported by Thunberg (1933). Alkylsuccinates likewise are
inactive until the length of the alkyl chain is greater than eight carbon
atoms. These higher alkylsuccinates may exhibit some degree of competitive
inhibition but there is also inhibition of the oxidation of p-phenylenediamine
and, hence, of the cytochrome system. Therefore, it is clear that additions
of even small alkyl groups to malonate and succinate depress or abolish
the normal inhibitory activity. Hydroxymalonate (tartronate) also is es-
sentially inactive. These results indicate the importance of that part of the
molecule between the two carboxylate groups and would seem to argue
against a simple interaction of the molecules with cationic groups on a
flat surface, the substituted groups protruding outward. Of course, it is
necessary that the — CHgCHo — grouping of succinate interact with the
enzyme in order for dehydrogenation to take place. The failure of short-
chain alkylmalonates to inhibit appreciably must be attributed to some
manner of steric interference by the alkyl groups.
(e) Cyclic dicarhoxylate ions. Cyclobutane and cyclopentane dicarboxylates
are weak inhibitors of succinate dehydrogenase. Since the intercarboxylate
distance in cyclobutane- 1,1-dicarboxy late and CT's-cyclopentane-l,2-dicar-
boxylate are not too far from that in malonate (Table 1-1), the poor inhibi-
tion may be due to the bulkiness of the rings interfering sterically, as do the
alkyl groiips discussed in the previous section. It is strange, however, that
there is no inhibitory difference between the cis and trans isomers of cyclo-
pentane-1,2-dicarboxylate, since the intercarboxylate distances differ by
1.39 A. Thus the inhibition may not be by the same mechanism as for mal-
onate; indeed, cw-cyclohexane-l,2-dicarboxylate inhibits succinate dehy-
drogenase noncompetitively (Hellerman et al., 1960).
Nature of the Active Center and the Binding of Malonate
The evidence indicates the presence at the active center of two cationic
groups and a nearby SH group. The cationic groups, perhaps 3-4 A apart,
are suggested by the very weak inhibitions exerted by monocarboxylates
(Quastel and Wooldridge, 1928; Dietrich et al., 1952) and the complete
lack of a competitive inhibition by compounds in which the negative
charges on the carboxylate groups are eliminated. Malondialdehyde (Holt-
INHIBITION OF SUCCINATE DEHYDKOGENASE 41
kamp and Hill, 1951), malondiamide (Fawaz and Fawaz, 1954), and var-
ious derivatives of succinate, in which one or both of the carboxylate
groups have been replaced with nonionic groups (Dietrich et al., 1952),
/COO
/OH
^CHO
CONH,
H,C
H-C
H^C^
H,C^
1
1
^CHO
CONHj
"^^-Br
"^*^^CN
Malondialdehyde Malondiamide /3-Bromopropionate /3- Hydroxy propiononitrile
are all lacking in inhibitory activity. An SH group close to the cationic
attachment points is proved by the high sensitivity of the dehydrogenase to
substances reacting with SH groups and the protection that malonate af-
fords against such substances. The latter is actually better evidence for the
proximity of the SH group because sulfhydryl reagents can alter protein
structure and exert effects for some distance over the enzyme, whereas the
blockade of these substances by a small molecule such as malonate would
be almost certain proof. Hopkins et al. (1938) showed that malonate, can
protect succinate dehydrogenase from oxidized glutathione (GSSG), which
is a potent inhibitor. Incubation of the enzyme with GSSG increased the
methylene blue reduction time from 10 min to 3 hr; malonate at concentra-
tions from 0.2 to 100 mM gave almost complete protection. That the
SH groups are protected by malonate was also shown by titration of these
groups with iodine. Potter and DuBois (1943) reported protection against
quinone inhibition and Barron and Singer (1945) against arsenicals. It is
interesting that malonate also protects against oxygen poisoning of suc-
cinate oxidase, which was interpreted to mean that the SH groups of the
enzyme are involved in this inactivation (Dickens, 1946 b). Oxalacetate
has also been shown to be protective against mercurials and arsenicals
(Stoppani and Brignone, 1957), providing evidence that it binds to the
same site as succinate and malonate. The degree of protection depends on
several factors, including the concentrations of the inhibitors and the rates
at which they react with the enzyme; the less protection seen against the
mercurials is attributed to their comparatively rapid action whereas the
Inhibitor
Oxalacetate
Inhibitor
concentration
{mM)
concentration
(mM)
0/
/o
Protection
jj-MB
0.5
1.3
21.8
p-Chloromercuriphenol
0.76
3.4
24.0
HgCl,
0.38
3.4
53.8
Methylarsenoxide
2.0
3.4
67.2
Oxophenarsine
0.36
1.3
81.2
42 1. MALONATE
arsenicals require at least 30 min to reach their maximal inhibition. Such
protection on a competitive basis was called interference inhibition by
Ackermann and Potter (1949). Once the sulfhydryl reagents have reacted
with the enzyme, malonate will not reverse the inhibition, but only slows
down the rate at which the substance acts on the enzyme. (I find it difficult
to understand how, in some cases, such low concentrations of malonate
afford protection against irreversible inhibitors. For example. Potter and
DuBois reported that 0.33 mM malonate protects quite well against
p-quinone, and yet this concentration of malonate inhibits only around
20%, showing most of the enzyme uncombined with malonate).
Succinate dehydrogenase also contains nonheme iron and flavin dinucleo-
tide but the locations of these components relative to the succinate-binding
site are not known. Since the iron and the flavin both participate in the
electron transfer, it is reasonable to assume that at least one of them is
close, or even part of the active center. Most formulations have pictured
the initial step as a transfer of hydrogen atoms from succinate to the fla-
vin; if this is so, the topography of the active center must be rather
complex.
We shall now turn to the energetics of the binding of malonate in order
to determine if the ionic interactions generally assumed are reasonable.
An immediate difficulty is the variability in the values of K^ reported, even
for the same tissue; for example, 0.0076 mM (Thorn, 1953) and 0.05 milf
(Hellerman et al., 1960) for the enzyme from pig heart, and 0.041 mM for
beef heart (Kearney, 1957). A K^ of 0.0076 mM would indicate an over all
binding energy of 7.24 kcal/mole, or 3.62 kcal/mole for each carboxylate
interaction assuming only ion-ion contribution. This is a reasonably high
value for the interaction of C00~ and NH3+ groups and corresponds to
an intercharge separation of around 4.30 A (Fig. 1-6-16), which is not far
from contact of the groups. It is unlikely that other types of interaction
are important for malonate. The corresponding K, for succinate is 0.0028
mM, giving an interaction energy of 3.28 kcal/mole per carboxylate group
and a separation of 4.45 A. A difference of fit of 0.15 A would thus account
for the relative bindings of malonate and succinate. However, in the case
of succinate it is more likely that other energy terms are involved. There
is undoubtedly some interaction between the — CH2CH2 — region and the
enzyme, and it is probable that distortion of the succinate molecule occurs
upon binding. In any event, these rough estimates point to a fairly close
fit of malonate to the enzyme cationic groups for the pig heart enzyme.
The ability of malonate to bind to the active center of succinate dehydro-
genases from bacteria is apparently much less (Table 1-8).
The next question is: what is the most probable distance between the
two enzyme cationic binding sites? Information on this must be obtained
from the relative bindings of substances having negatively charged groups
INHIBITION OF SUCCINATE DEHYDROGENASE 43
different distances apart. Since malonate is usually bound more tightly
than succinate, and much more than oxalate, it is reasonable to assume an
intercationic distance approximating the intercarboxylate distance in mal-
onate. It is by no means necessary that the substrate in its free configu-
ration exactly fits the enzyme site. Pauling (1946, 1948) has suggested,
" an active region of the surface of the enzyme... is closely complementary
in structure not to the substrate molecule itself, in its normal configuration,
but rather to the substrate molecule in a strained configuration, correspond-
ing to the activated complex for the reaction catalyzed by the enzyme."
Now, the fact that malonate fits the active site well does not mean that the
enzyme cationic groups are the same distance apart as the carboxylate
groups (3.28 A). The calculations above indicate a distance of 4.3 A
between carboxylate and cationic groups and thus, depending on the
geometry of the binding, the cationic groups could be much farther apart
than 3.28 A. Extreme situations are shown in Fig. 1-7, where the intercat-
FiG. 1-7. Representations for the ex-
treme situa/tions in the interaction of
malonate with the two cationic groups
on the surface of succinate dehydro-
genase. In both cases (A and B) the
interaction distances and the energies
between the ionic groups are the same.
ionic distance may vary from 3.28 to 13.1 A, approximately the same energy
of binding being expected in either case. Situation A is not very likely
because it is improbable that protein cationic groups would occur so close,
and, furthermore, in this case oxalate might be expected to bind quite well.
Also, situation B would provide more opportunity for succinate to be dehy-
drogenated at the enzyme surface. Of course, the enzyme surface at the
44 1. MALONATE
active center may not be smoothly curved as shown, and we shall soon
examine evidence that it is not.
The problem of the interactions of di-ionic substances with receptor
groups has been recently treated by Schueler (1960, p. 448) and on the basis
of statistical calculations he has concluded, "The most dramatic alteration
in activity should occur upon approaching that agent in the series which
possesses a length distribution just capable of overlapping the negative-
charge spacing in the receptor, and this should be followed by a relatively
slow rate of loss in activity with respect to increasing length" (he is assum-
ing a positively charged drug). In view of what has just been said above,
there is some doubt if predictions like this can be made with confidence.
The large distances between the interacting charged groups make it very
difficult to assign receptor configuration and much will depend on the over
all configuration of the protein surface. Other factors, such as distortion of
long-chain molecules, interactions of regions between the end charged
groups the surface, and possible steric repulsions, must be considered.
Inasmuch as little accurate information on the intercationic distance can
be obtained from Kj and K^. values alone, let us now turn to more profitable
considerations of the topography of the active center. There is evidence
from several lines that the active center is not a flat or slightly convex
surface. In the first place, alkylsuccinates are bound to the enzyme very
poorly; methylsuccinate is oxidized at 23% the rate for succinate, and
ethylsuccinate at 18% the rate for succinate, while higher members are
neither oxidized nor are they inhibitory (Franke, 1944 a). In the second
place, as we have already seen, alkylmalonates are very poor inhibitors.
Indeed, even the introduction of a hydroxyl group (tartronate) reduces the
inhibition markedly. These observations indicate a rather close fit for
malonate and succinate at the active center, additional groups giving rise
to steric repulsion, as frequently reported for antigen-antibody reactions.
In the third place, fumarate is bound fairly well while maleate is not, indi-
cating again some steric repulsion since the intercarboxylate distances
alone would certainly allow predictions that maleate would be bound more
tightly. All molecules that bind appreciably to succinate dehydrogenase
seem to be simple linear substances, or substances capable of assuming a
linear configuration. All of this evidence points to a slit or tubular structure
for the active center, such that compounds with added groups or rigid non-
linear molecules cannot enter. Such a situation is pictured in Fig. 1-8 in
two dimensions. This is not to be construed as an attempt to represent
the actual configuration but merely to show the steric barriers impeding
attachment of larger or nonlinear molecules. Glutarate and adipate could
not fit well, not because of unsatisfactory intercarboxylate distances, but
because of the bulkiness of the longer hydrocarbon chains. Fumarate is
able to bind because its configuration is much like that of succinate in the
INHIBITION OF SUCCINATE DEHYDROGENASE
45
extended form shown in the figure, whereas maleate might not fit because
of its nonlinear structure. Acetylene-dicarboxylate would be expected to in-
hibit to some extent because of its linearity. Such a model would also ex-
plain why small alkyl groups added to succinate do not completely abolish
the binding. It may also be mentioned that this type of configuration would
allow the flavin and iron components of the dehydrogenase to be in posi-
tions close to the — CH2CH2 — group and thus able to participate in the
removal of the hydrogen atoms.
Succinate
Fig. 1-8. Representations of the binding
of malonate and succinate at the active
site of succinate dehydrogenase, indicat-
ing the steric barriers possibly surround-
ing the region of the two cationic sites.
The actual situation must be visualized
in three dimensions.
Activation of Succinate Dehydrogenase by Malonate
Preparations of beef heart succinate oxidase obtained using borate buf-
fer are not fully active but may be activated by the addition of phosphate.
This interesting discovery by Kearney (1957, 1958) may have important
bearings on the understanding of the active center of this enzyme, especially
as she later found that succinate, fumarate, and malonate also activate,
and indeed are much more potent than phosphate (see accompanying
tabulation). Once the enzyme has been activated, the activator can be
removed without loss of the activity; in fact, malonate must be dialyzed
away if the full activity of the enzyme is to be measured. The activation
constants are quite different from the Michaelis or inhibitor constants for
these substances. It would appear that malonate binds more tightly to the
less active form of the enzyme. Kearney favors the view that these activa-
46 1. MALONATE
tors convert a less active form of the enzyme to a more active form, and
that this transformation may involve a localized change in the protein
Activator
(mJf) (mM)
Malonate
0.0072
0.025
Succinate
0.12
0.52
Fumarate
5.6
0.80
Phosphate
100
—
structure because of the high energy of activation. Whatever the explanation
it is important to remember that malonate can exert an activating effect,
as weU as inhibiting, in media low in phosphate.
Effects of Various Factors on the Inhibition of Succinate Dehydrogenase
Very few illuminating studies on the modification of malonate inhibition
are available. The effects of temperature were mentioned in Volume I, where
the following thermodynamic parameters for the inhibition of beef heart
succinate dehydrogenase at 38° were calculated: AF = — 6.26 kcal/mole,
AH = — 5.48 kcal/mole, and AS = 2.6 cal/mole/degree. The K^^ for malonate
was found to be 0.025 mM at 20o-23o and 0.041 mM at 38o (Kearney, 1957).
The effects of osmolarity on the inhibition are surprisingly large (Honda and
Muenster, 1961). Lupine mitochondria were prepared in media of different
sucrose concentrations and assayed in media of two osmolarities (Table
1-9). These results were obtamed on mitochondria and it is possible that
the effects are not directly on succinate oxidase but on the permeability
or structural properties of the mitochondria. In this connection it has been
pointed out by Singer and Lusty (1960) that iV-methylphenazine measures
the full activity of succinate dehydrogenase in mitochondrial fragments,
but in intact mitochondria it measures only a fraction of the activity.
Various ways of damaging the mitochondria lead to increased succinate
dehydrogenase activity (such as increase in Ca++ concentration). This was
interpreted in terms of permeability barriers to A^-methylphenazine (and
also FMNHj), limiting the rate in intact mitochondria. It could also be
explained on the basis of structural changes in the enzyme complexes. In
any case, these observations demonstrate the importance of the mitochon-
drial state in the functioning of succinate oxidase, and it would not be
surprising if malonate inhibition were similarly sensitive. The inhibition
of succinate oxidation by malonate in rat heart mitochondria in KCl me-
dium is not altered by either halving or doubling the KCl concentration in
the assay medium (Montgomery and Webb, 1956 b). However, the effects
INHIBITION or SUCCINATE DEHYDROGENASE
47
Table 1-9
Effect of Osmolarity in the Properties of Succinate Oxidase in Lupine
Mitochondria "
Preparation
Assay
K
K-
osmolarity
osmolarity
(M)
(mM)
(mM)
KJKi
a
0.15
0.22
5.10
0.91
5.6
2
0.60
12.34
0.64
19.3
1
0.40
0.22
2.87
0.19
15.1
8
0.60
5.47
0.16
34.2
191
0.60
0.22
1.20
0.05
24.4
36
0.60
5.47
0.11
49.7
268
" Kj„ is the Michaelis constant for succinate, iT, the inhibitor constant for malonate,
and a is the interaction constant defined in Eqs. 1-3-5 and 1-3-6, indicating the type
of inhibition. The osmolarity is given in terms of sucrose concentration. (From Honda
and Muenster, 1961.)
of Ca'^^ concentration on both succinate oxidation and malonate inhibition
are complex (Fig. 1-9) and difficult to explain. The decrease in the rate
of oxidation beyond Ca+"^ concentrations around 5 m.M might be attributed
to a complexing of the succinate, but for the same reason, namely, the
complexing of malonate, which has a higher affinity for Ca+''" than does
succinate, the inhibition would be expected to decrease. Whether the
0.001 0 01
Fig. 1-9. Effects of Ca++ on the rate of succinate oxidation and
the inhibition by malonate in rat heart mitochondria. Succinate is
5 mM and malonate is 1 mM. (From Montgomery and Webb, 1956 b).
48 1. MALONATE
modest stimulation of succinate oxidation by low concentrations of Ca^^
is a permeability or structure-opening effect on the mitochondria, or due
to a more direct effect on the enzyme, is not known; certainly the oxidations
of other cycle substrates and pyruvate are strongly depressed by Ca++.
The effects of Mg++ concentratioti from 6.2 to 12.4 mM on trypanosomal
succinate dehydrogenase inhibition by malonate have been reported as
negligible (Agosin and von Brand, 1955). In connection with the relation-
ship between mitochondrial structure and malonate inhibition, it is interest-
ing to examine the effect of ATP, inasmuch as ATP protects or stabilizes
mitochondria after isolation from cells. ATP at 1 mM has very little effect
on the rate of succinate oxidation in rat heart mitochondria (in five exper-
iments a mean depression of 3%), but in the presence of succinate (5
mM) and malonate (5 mM) it stimulated the rate 67%, thus antagonizing
the inhibition by malonate (Montgomery and Webb, 1956 b). Similar re-
sults were seen with acetylene-dicarboxylate inhibition. If the effect of
ATP is to reduce the permeability to these inhibitors, it is surprising that
interference with succinate penetration does not also occur. Thus some
other explanation may have to be sought.
Inhibition of Fumarate Reduction
If succinate, fumarate, and malonate bind at the same site on succinate
dehydrogenase, the reverse reaction — the hydrogenation of fumarate to
succinate — should be inhibited by malonate; that is, malonate should
compete with fumarate as well as with succinate. The relative potencies
of the inhibitions on the two reactions would depend on the Michaelis
constants for succinate and fumarate, so that the inhibitions would not
necessarily be identical. Malonate was, indeed, found to inhibit the reduction
of fumarate by horse muscle succinate dehydrogenase, but less potently
than the oxidation of succinate (18 mM malonate required to inhibit 50%
in the former case and 3.6 mM in the latter) (Das, 1937 b). A more detailed
analysis was made by Forssman (1941) in Lund, using pig heart succinate
dehydrogenase and leucomethylene blue as a hydrogen donor, in this case
the inhibition by malonate being essentially equivalent for both forward
and backward reactions. More recent studies of beef heart succinate dehy-
drogenase, however, have given different values for K,: 0.025 mM for suc-
cinate oxidation and 0.12 mM for fumarate reduction (Singer et al., 1956 a).
This difference is unexpected and the suggestion was made that the binding
of malonate may be effected by the state of oxidation of the electron trans-
port components adjacent to the binding site. The affinities of fumarate
for the enzyme were also found to be different for each reaction. It may also
be of significance that iV-methylphenazine was the acceptor in the oxida-
tion of succinate, and FMNH, or leucodiethylsafranin the donor in the re-
duction of fumarate.
INHIBITION OF SUCCINATE DEHYDKOGENASE 49
The succinate dehydrogenase of Micrococcus lactilyticus behaves quite
differently and is poorly inhibited by malonate, a ratio of (malonate)/(fu-
marate) = 20 being required for 29% inhibition (Peck et al, 1957). It might
be thought that this enzyme is not succinate dehydrogenase, but another
enzyme that could be called " fumarate reductase," especially as fumarate
is reduced at a faster rate than succinate is oxidized, in contrast to the
mammalian enzymes. However, it has been conclusively demonstrated
it is not a separate enzyme and that the failure of malonate to inhibit
is to be attributed to a very high affinity for fumarate coupled with a rela-
tively low affinity for malonate (Warringa et al., 1958). The configuration
of the active center of the bacterial enzyme must differ from that of the
mammalian enzymes. This may also explain the "fumarate reductases"
obtained from yeast (Fischer and Eysenbach, 1937; Kovac, 1960) which are
rather insensitive to malonate.
Variations of Malonate Inhibition of Succinate Dehydrogenases
from Different Tissues and Species
The comparative biochemistry of enzyme inhibition is in its infancy and
accurate comparison of results is usually impossible due to the different
conditions under w^hich the inhibitions w^ere studied. Examination of Table
1-6 with a view to establishing phylogenetic relationships is made difficult
by the different types of preparation and assay procedure used. A correla-
tion graph, made by plotting inhibitions against (I)/(S) ratios, shows a very
marked scatter of the points. For example, at an (I)/(S) ratio of 0.1. the
inhibitions range from 10% to 100%. This variation cannot all be due to
the differences in technique. All of those cases in which the malonate inhibi-
tion is significantly below the mean turn out to be in the bacteria, inverte-
brates, or plants. However, this is not a strict correlation because some of the
potent inhibitions have been found in such organisms {e.g., Azotobacter, E.
colt, and Trypanosoma). The possibility of a relationship between succinate
dehydrogenase type in the bacteria and the oxygen requirements for growth
has been proposed. The enzyme from the obligate anaerobe Micrococcus
lactilyticus has low affinities for succinate and malonate, as discussed in the
previous section, whereas the enzyme from the facultative anaerobe,
Propionibacterium pentosaceum is intermediate in properties between Mi-
crococcus and the aerobic mammalian tissues (Singer and Lara, 1958). Cer-
tainly many invertebrate and plant tissues can withstand anaerobiosis
better than mammalian tissues, but at the present state of our knowledge
such a correlation is dangerous to make.
Comparisons of the malonate inhibitions of succinate dehydrogenases
from different tissues have been reported in a few cases. The epithelium
and muscle of guinea pig seminal vesicle were separated and the inhibitions
by malonate at four different concentrations were similar (Levey and Szego,
50 1. MALONATE
1955). However, the inhibitions of epithelial dehydrogenase appear to be
about 5% higher than for the muscle enzyme, although this may not be
statistically significant. Malonate was found to inhibit Hepatoma 134
tumor succinate dehydrogenase more than the enzyme from normal mouse
liver (Fishgold, 1957) over a range of five malonate concentrations; for
example, 0.21 mM malonate inhibited the liver enzyme 42% and the he-
patoma enzyme 76% at a succinate concentration of 1 vaM. Killer and
sensitive stocks of paramecia may have succinate dehydrogenases with
different sensitivities to malonate, but it is difficult to draw conclusions
from the data published (Simonsen and van Wagtendonk, 1956). The Og
uptake of homogenates of the two strains was increased to different de-
grees by 50 mM succinate and malonate inhibited both quite well (see
accompanying tabulation). The authors concluded that the enzyme from
the killer strain, is inhibited more, based on absolute reduction, but actually
Increase in Oj uptake from succinate
Sensitive strain Killer strain
Control 1.4 14.0
With malonate 80 mM 0.2 3.8
% Inhibition 86 73
the enzyme from the sensitive strain seems to be inhibited as w^ell. Our
work with succinate dehydrogenase from various rat tissues (Table 1-6)
indicates no significant difference in susceptibility to malonate.
INHIBITION OF SUCCINATE OXIDATION
IN CELLULAR PREPARATIONS
Attention will now be turned to the inhibition of the succinate oxidase
system when it is located in the normal cellular structure and succinate is
added exogenously to the preparations. When succinate is added to most
cell suspensions, minces, or slices, there is an increase in the O2 uptake,
and this response is inhibited to varying degrees by malonate (Table 1-10).
It is particularly important in cellular preparations to take account of the
endogenous respiration and the effect of malonate on it (see Chapter 1-9).
In many studies this has not been done and this is one factor that makes
it difficult to compare accurately the malonate inhibitions m vitro and in
vivo. Since the endogenous respiration is generally inhibited less than
succinate oxidation by malonate, failure to correct for endogenous respi-
ration usually leads to low values for the inhibition. This is illustrated in
INHIBITION OF SUCCINATE OXIDATION 51
the four examples given in Table 1-11. The importance of the endogenous
correction is seen to vary with the effect of malonate on the endogenous
respiration. When malonate inhibits the endogenous O2 uptake poorly,
as in rat liver, or actually stimulates the O2 uptake (due to its metabolism),
as in Euglena, the true inhibition of succinate oxidation is much higher than
would be calculated simply from the data on succinate and succinate +
malonate.
Comparison of the inhibitions in Tables 1-6 and 1-10 for the same species
or tissues, although this is qualitative only, shows that in several instances
the inhibitory potency of malonate seems to be significantly less in cellular
preparations. This is true for E. coli, Rhodospirilliim, Crithidia, Zygor-
rhynchus, pigeon muscle, and rat liver. Moses (1955) points out that in
Zygorrhynchus the inhibition of succinate oxidation in cell suspensions is
very weak, even at the low pH of 3.4, but when the cells are treated with
liquid nitrogen to destroy their structure, malonate inhibits normally.
The oxidation of succinate by cell susj^ensions of Bacterium succinicum is
not inhibited at all by 5 mM malonate whereas such oxidation in cell-free
extracts is inhibited completely (Takahashi and Nomura, 1952). There are
also several reports in which malonate was found to be ineffective but the
extracted succinate oxidase system was not directly tested; however, in
these cases one would certainly expect the enzyme to be sensitive to mal-
onate. For example, malonate (2 mM) does not inhibit the oxidation of
succinate by barley roots (Honda, 1957), or at 5-20 mM in dried cells of
CJdorella (Millbank, 1957), while in beech roots malonate (28.6 mM) actually
stimulates the rate of succinate oxidation (Harley and Ap Rees, 1959).
However, in some cases comparable inhibitions have been observed in vitro
and in vivo. Danforth (1953) showed in Euglena that malonate is quite ef-
fective in intact cells if the pH is low enough (around 4.5) and, although it
is difficult to compare the results with those obtained from homogenates
because of different concentrations of succinate and malonate, it would
appear that malonate is equally inhibitory in the two preparations. Similar
effects of malonate were also observed in our work (Montgomery and Webb,
1956 b) on rat heart slices and mitochondrial suspensions.
The failure of malonate to inhibit the oxidation of added succinate in
cellular preparations well, or at all, has usually been attributed to permea-
bility factors. However, it is difficult to understand how permeability could
explain these results, inasmuch as the penetration of both succinate and mal-
onate would be controlled by the same factors, presumably. That is, if there
is some barrier to malonate reaching the succinate dehydrogenase, how can
succinate pass this barrier? It is true that the "pK,, for succinate is higher
than for malonate but almost identical values of ^K,, have been reported;
thus the distribution of the ionic forms around neutrality would be approxi-
mately the same (Table 1-3). If only the uncharged forms of these acids —
52
1. MALONATE
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INHIBITION OF SUCCINATE OXIDATION
53
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1. MALONATE
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INHIBITION OF SUCCINATE OXIDATION
55
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56 1. MALONATE
Table Ml
Effects of Endogenous Respikation on Calculations of Malonate Inhibition
OF Succinate Oxidation "
Pigeon
muscle
Rat kidney
Rat liver
Euglena
O2 Uptake
Endogenous
11.8
14.3
14.7
11.4
Malonate
3.0
4.5
12.4
13.9
Succinate
19.0
58.5
20.2
28.0
Succinate + malonate
6.0
15.8
14.1
19.7
% Inhibition
Endogenous respiration
74.6
68.6
15.6
Stim21.9
Total respiration in the
presence of succinate
68.4
73.0
30.2
29.6
Succinate oxidation
58.3
74.4
69.1
65.1
° The true inhibition of succinate oxidation is given in the bottom row, and is to
be compared to the figures in the row immediately above where the correction for the
endogenous ejRFect has not been made. The concentrations were: pigeon muscle — suc-
cinate 1 vaM and malonate 10 iwM (Stare and Baumann, 1939); rat kidney — suc-
cinate 20 n\M and malonate 20 mM (Fawaz and Fawaz, 1954); rat liver — succinate
10 n\M and malonate 20 mM (Edson 1936); Euglena — succinate 10 mM and malonate
10 ml/ (Danforth, 1953).
HOOC — R — COOH — penetrated, succinate would enter cells somewhat
better than malonate, but, at least at pH's above 7, it seems unlikely
that this is the situation. Also, the entrance of enough of the undissociated
caid to be effective would presumably decrease the intracellular pH signi-
ficantly (see Chaper 1-14). Besides, some of these experiments in which
malonate was inactive were done at low pH's (3.4 to 5.5). A question that
must be considered is whether succinate oxidation is always entirely intra-
cellular. It is possible that succinate oxidase occurs both in the mitochondria
and in the plasma membrane. In many cases, malonate has little or no effect
on tissue metabolism or function at concentrations capable of inhibiting
the oxidation of added succinate completely. This is well seen in rat ven-
tricle slices (Webb et al, 1949) where the succinate may be oxidized at
the cell surface, and it is interesting that in this tissue the potency of mal-
onate in vivo and in vitro is the same. However, in most instances, the
succinate oxidase seems to be protected in some manner in intact cells.
There are several possible factors that could modify the malonate inhi-
INHIBITION OF SUCCINATE OXIDATION 57
bition of succinate dehydrogenase when the enzyme is isolated from the
cells. It will be well to mention some of these in order to emphasize that
there are usually many ways, other than by permeability, by which unex-
pected phenomena may be explained.
(a) The enzyme environment within the cell is different from the artificial
media used with isolated enzymes (see Chapter 1-9). Many substances may
be able to alter malonate inhibition; we have noted the effects of Ca+"'"
and ATP. The concentrations of such substances may be different in cell
and medium. The intracellular pH is also not that of most media used in
enzyme study and may be easily changed by the addition of external
substrate or inhibitor.
(b) The addition of succinate to cells may influence the endogenous respiration;
that is, the change in Og uptake upon adding succinate may not represent
accurately the rate of succinate oxidation. In other words, the correction
for endogenous respiration may be in error. Also, the addition of succinate
or malonate can secondarily alter the complex balance of the metabolic
systems. For example, oxalacetate is a very potent inhibitor of succinate
dehydrogenase and its concentration in the cell may be a controlling factor
in the operation of the cycle. Since oxalacetate can be formed from succin-
ate, and its formation inhibited by malonate, secondary changes in succinate
dehydrogenase activity may occur that are easily interpreted as due to
the direct effects of malonate.
(c) The concentration of succinate in the cell may already be appreciable
and the addition of more may reduce the inhibition by malonate because
of the competitive nature of the inhibition.
(d) The rate of succinate oxidation may be limited by the rate at which it
can enter into the cells or tissus; that is, the succinate oxidase is so active
that the succinate is oxidized as rapidly as it enters. In such a case, mal-
onate would be quite ineffective until the enzyme is inhibited sufficiently
to make it limiting.
(e) The ratio (malonate) I (succinate) may be different in the cell than in the
medium. This could be due to differences in the permeabilities of the cell
membrane towards these substances for, despite the fact that succinate and
malonate have similar properties, many cases of differential permeabilities
to more closely related ions are known. It could also be due to the somewhat
different p/C^ values, since the internal concentrations of the active ions are
determined by the ionization constants (see Eq. 1-14-146 for buffered
cells).
Whether these factors, or others, are responsible for the anomalous results
mentioned above is not known. It would be very useful to have data on the
58 1. MALONATE
rates of penetration of succinate and malonate into cells, obtained prefer-
ably with radioactive material and under the same conditions. Quantitative
studies on the malonate inhibition of intracellular dye reduction resulting
from succinate oxidation might also be informative in certain instances.
It has been shown that malonate inhibits the succinate-induced reduction of
neotetrazolium in adipose tissue cells (Fried and Antopol, 1957), but noth-
ing otherwise is known about the succinate dehydrogenase from this tissue.
Competitive Nature of the Inhibition in Cellular Systems
It has been shown in several tissues that the addition of succinate will
reverse the inhibition of respiration produced by malonate. Thus in Avena
coleoptile the inhibition by 50 mM malonate is reduced from 57.4% to
25.8% upon adding succinate (Bonner, 1948), and in spinach leaves from
75.4% to 20.5% (Bonner and Wildman, 1946). In chick embryonic carti-
lage, the depression of respiration by 10 mM malonate is reversed by 100
mM succinate but not by 10 mM (Boyd and Neuman, 1954). Such results
have occasionally been stated to prove the competitive nature of the inhibi-
tion but this reasoning is not completely valid. The mere increase in O2 up-
take seen on addition of succinate to malonate-inhibited tissues is alone
not evidence for competition. The effects of succinate on uninhibited tissue
must also be tested and it must be shown that the actual inhibition is de-
creased. A decrease in the inhibition brought about by increasing succinate
concentrations has indeed been reported in two tissues, pigeon breast muscle
(Krebs and Johnson, 1948) and the trypanosome Crithidia (Hunter, 1960)
and in the latter a true competitive inhibition was demonstrated by 1/v — 1 /(S)
plots. The data are given in Table 1-10. It is probable that the inhibition
of succinate oxidation in cellular systems by malonate would frequently
not obey strictly competitive kinetics, due to the various complexities that
arise, as discussed in the previous section, even though the primary inhi-
bition on the succinate dehydrogenase were competitive. Some of the prob-
lems involved in the determination of the type of inhibition in cells have
been discussed in Chapter 1-9.
INHIBITIONS OF ENZYMES
OTHER THAN SUCCINATE DEHYDROGENASE
It is very important to establish the degree of specificity that may be
achieved in the use of malonate under various conditions. To this end
we shall first discuss the direct evidence for the inhibition of enzymes
other than succinate dehydrogenase, and then proceed to the effects on the
operation of the tricarboxylic acid cycle, the accumulation of succinate and
other intermediates, and finally the antagonism of the malonate inhibition
ENZYMES OTHER THAN SUCCINATE DEHYDROGENASE 59
by fiimarate. We shall then be in a position to evaluate the specificity of
malonate. There are other reasons, of course, for taking up these subjects;
for example, malonate is frequently used to block the cycle in living tissue
and in this connection it is essential to understand how malonate can alter
cycle activity under various conditions.
Some effects of malonate on miscellaneous enzymes are presented in
Table 1-12. There are three major difficulties in the establishment of the
over all spectrum of action of malonate. First, there are many quite impor-
tant enzymes whose response to malonate has never been investigated
directly. Second, inspection of the table will show that in few cases has
more than one concentration been used, and often the single concentration
reported is either too high or too low to be of much value. Third, the same
enzyme from different species often shows widely varying susceptibility
to malonate (e.g. NADH oxidase, /5-glucuronidase, lactate dehydrogenase,
oxalacetate decarboxylase, and others in the table), making it clear that
there are many different spectra of malonate inhibition and that statements
on specificity must be qualifield by naming the source of the enzymes in
question.
Ideally, the results on the inhibition of an enzyme by malonate should
be given for several concentrations, preferably convering the range from that
concentrations just sufficient to produce some inhibitions, through that
causing approximately 50% inhibition to higher inhibitions (unless these
latter concentrations are unreasonably high). Another way of looking at the
problem is to consider that range of malonate concentrations most likely
to give useful information when tested on enzymes. This will depend
on the source of the enzymes. For example, in mammalian tissues it usu-
ally requires malonate concentrations between 2 and 5 mM to inhibit suc-
cinate dehydrogenase around 90% in the presence of 5-10 mM succinate.
In the interests of establishing the degree of specificity, it would thus be
most important to test the effects of malonate at concentrations around
5 mM on enzymes from such sources. When the organism studied possesses a
succinate dehydrogenase less sensitive to malonate, correspondingly higher
concentrations must be applied to the other enzymes.
Instances of Competitive Inhibition
Until it is time to discuss the matter of specificity, there is little to say
about these inhibitions since the results in the table speak for themselves.
It is evident that several enzymes other than succinate dehydrogenase
are readily inhibited. One would not be surprised if enzymes attacking the
dicarboxylate anions, where the carboxylate groups are separated by two
carbon atoms, were inhibitable by malonate to some extent, since it is
likely that these enzymes also possess cationic groups appropriately spaced.
Actually, fumarase, malate dehydrogenase, the malic enzyme, oxalacetate
60
1. MALONATE
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ENZYMES OTHER THAN SUCCINATE DEHYDROGENASE 61
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ENZYMES OTHER THAN SUCCINATE DEHYDROGENASE
65
decarboxylase, and the condensing enzyme are inhibited by malonate, al-
though usually not as potently as is succinate dehydrogenase. Enzymes
catalyzing reactions of the dicarboxylates in which the charges are farther
apart (e.g. a-ketoglutarate) would be expected to be less susceptible. It is
likely that these inhibitions are mosth^ competitive but sufficient data to
establish this are generally lacking.
The inhibition oifumarase by malonate has been shown to be competitive
in the thorough study of Massey (1953 b) but the affinity of the enzyme for
malonate is not very high {K^ = 40 n\M). This implies either a very dif-
ferent intercationic distance in this enzyme from that in succinate dehydro-
genase or a different configuration of the enzyme surface surrounding these
cationic groups, probably the latter. Both directions of the reaction cata-
lyzed by the malic enzyme are inhibited by malonate, which competes
with either malate (Stickland, 1959 b) or p\Tuvate (Stickland, 1959 a) (see
accompanying tabulation). The inhibition of this enzyme might well alter
Malate
% Inhibition by malonate at:
Pyruvate
% Inhibition by malonate at:
(mif)
2 mM
5 mM
20 mM
{mM)
1 mM
10 mil/
0.1
71
88
100
1
43
62
0.3
48
74
91
10
37
70
1
19
48
84
50
10
46
the operation of the tricarboxylic acid cycle under certain conditions. The
situation is different for lactate dehydrogenase, since malonate is competitive
with respect to lactate but noncompetitive with respect to pyruvate (Ot-
tolenghi and Denstedt, 1958), leading to the suggestion that these two sub-
strates react with different sites on the enzyme. The X/s are 6.4 m.M for
the oxidation of lactate and 27 mM for the reduction of pyruvate. Oxalate
and tartronate are much better inhibitors of this enzyme. The D-a-hydroxy
acid dehydrogenase of yeast, which oxidizes lactate, malate, a-hydroxybuty-
rate, and glycerate, is competitively inhibited by malonate with a K^ of
0.9 mil/ (Boeri et al., 1960). In this case, oxalate is a very potent inhibitor
{Ki = 0.0025 mM) while tartronate is of similar potency to malonate.
Finally, in reducing systems in which succinate can serve as an electron
donor, malonate maj^ inhibit competitively. This is the case for the particu-
late nitrate reductase of soybean root nodules, the K^ for malonate being
0.017 milf (Cheniae and Evans, 1959). The nitrate reductase is not inhibited
by malonate directly but the results on the over aU system might make it
appear to be the case. In all cases where malonate inhibits competitively,
the susceptibility of the reaction in complex systems will depend on the
concentration of the substrate, and thus may be quite high in living systems
where the concentrations of intermediates are frequently low.
66 1. MALONATE
Inhibition Due to Chelation with Metal Ion Cofactors
Many enzymes are dependent on dissociable metal ions for their activity
and the operation of most of the important metabolic systems thus requires
the presence of these cofactors. The list of enzymes requiring Mg++ is a
long one and includes the oxidases and decarboxylases for the keto acids,
most of the enzymes involved in phosphate metabolism (e.g., the kinases,
the transphosphorylases, the phosphatases, and the acyl — CoA synthetases),
some dehydrogenases e.g., phosphoglucose dehydrogenase, phosphogluco-
nate dehydrogenase, and isocitrate dehydrogenase), some peptidases,
phosphoglucomutase, and enolase. Several enzymes require Zn++, such as
lactate dehydrogenase, glutamate dehydrogenase, alcohol dehydrogenases,
carboxy peptidase, and carbonic anhydrase. Since malonate is able to chelate
effectively with these metal ions, inhibition may result from the reduction
of metal ion concentration in the medium or the removal of the metal
ions from the enzyme. The ability of malonate to inhibit by this mechanism
will depend on the affinity of the enzyme for the metal ion. The binding
of Zn++ to enzymes is usually rather strong and it is difficult for malonate
to deplete the enzyme of this metal, but Mg++ is more loosely bound in most
cases and the activity of enzymes dependent on it is generally related to
the Mg+'*" concentration in the medium. The effect of malonate on Mg++-
dependent enzymes will thus depend on the concentrations of Mg++ and
malonate, and on the relationship between enzyme activity and Mg+*^
concentration. In the use of malonate, especially at higher concentrations,
it is imperative to consider the possibility of such effects. It is likely that
some of the inhibitions in Table 1-12 are due to metal ion depletion.
Inhibition by reaction with an activator was discussed briefly in Chapter
1-3 and it was seen that in the general case no simple expression for the
inhibition is possible. Nevertheless, it should be reasonably easy to deter-
mine if the inhibition is purely the result of activator depletion, since the
concentration of free activator can be calculated from the dissociation
constant of the activator-inhibitor complex by an equation similar to
Eq. 1-3-72:
(Mg) = ^ V [(I,) - (Mg,) + K-\- + 4(Mg)if - -^ [(I,) - (Mg,) + K] (1-3)
If the effect on the enzyme is only to reduce the Mg++ concentration, the
addition of malonate should bring the activity to that value corresponding
to the reduced free Mg++. Another possibility is that the Mg-malonate
complex is the active inhibitor, in which case the kinetics should be investi-
gated with the calculated concentrations of this complex at different con-
centrations of Mg++ and malonate.
The most thorougly studied instance of the possible relationship of Mg++
to inhibition by malonate is the work on the utilization of oxalacetate
ENZYMES OTHER THAN SUCCINATE DEHYDROGENASE
67
in rat tissue homogenates by Pardee and Potter (1949). The formation
of citrate here probably involves decarboxylation of some of the oxal-
acetate to pyruvate, with subsequent condensation to enter the cycle. Al-
though a single enzyme was not studied, it is possible that the utilization
of oxalacetate was limited by the decarboxylase. This reaction is activated
by low concentrations of Mg++ and inhibited by higher concentrations
(upper curve in Fig. 1-10). It is clear that the inhibition by malonate de-
( Mg* • ).
30
mM
Fig. 1-10. Utilization of oxalacetate by a rat
kidney homogenate at diflferent concentrations
of Mg++, with and without malonate. (From
Pardee and Potter, 1949).
creases as the Mg"*"*" concentration is raised (lower curve). This was interpret-
ed to mean that the inhibition is mainly due to depletion of Mg++. Some
arguments may be brought against this interpretation. Table 1-13 gives
my calculations of the concentrations of free Mg^^, free malonate, and the
complex at the different levels of total Mg++ used in the experiment shown
in Fig. 1-10. It is seen that the malonate at 10 mM does indeed reduce the
free Mg++, but in the higher range of Mg++ concentrations this should in-
crease the activity rather than decrease it, because in this range Mg"'"+ is
somewhat inhibitory. Pardee and Potter did not consider the reduction in
free malonate concentration, which is very marked as shown in Table 1-13.
This could well acount for the decrease in the inhibition from 47% at zero
Mg++ concentration to 12% at 30 roM Mg++. One might speculate that the
reduction in rate at high Mg++ concentrations could be due to the complexing
of oxalacetate so that it is unable to react with the enzyme. The question
of the mechanism of the malonate inhibition is thus not settled. It was
claimed that the inhibition is not typically competitive because increase
in the oxalacetate concentration actually increases the inhibition somewhat,
68 1. MALONATE
Table 1-13
Effects of Total Mg++ Concentration on the Concentrations of Free Mg++,
Free Malonate, and Ms-Malonate Complex "
(Mg++,)
(Mg++)
(Mg
-malonate)
(Malonate)
(milf)
(mif)
(mif)
(mM)
0
0
0
10
3
1.59
1.41
8.59
10
6.13
3.87
6.13
20
14.09
5.91
4.09
30
22.98
7.02
2.98
" Conditions as in Fig. 1-10 from the work of Pardee and Potter (1949). The total
malonate concentration is 10 mM in all cases. The values were calculated from Eq.
1-3 using 9.77 X 10^^ M for the dissociation constant of the Mg-malonate complex.
although no figures were given so the magnitude of the effect is unknown.
An increase of oxalacetate might reduce the amount of Mg++ bound to
malonate and thereby increase the free malonate concentration. It would
appear unlikely that the Mg-malonate complex is inhibitory since its con-
centration increases with Mg++ concentration (Table 1-13), whereas the
inhibition decreases. However, it is evident that malonate inhibits this
enzyme system in some manner directly, not only from the above consid-
erations but also because of the marked inhibition observed in the absence
of Mg++.
Dialkylfluorophosphatase is activated by Mn++ and its inhibition by
malonate was attributed by Mounter and Chanutin (1953) to the chelation
of the activator. Since no concentrations of either Mn++ or malonate were
given, it is impossible to evaluate the results. However, their data show
that malonate inhibits just as well, if not better, in the absence of Mn++
(21% with Mn++ added and 28% without Mn++ at 15 min). These results
might better be interpreted as due to removal of free malonate by the
Mn++. If Mn-malonate is incubated with the Mn-free enzyme, activity
slowly appears, indicating that the Mn++ is transferred from the malonate
to the enzyme, which is not surprising since the affinity of the enzyme for
the Mn++ is much greater, as shown by the dissociation constants {pK for
enzyme-Mn++ complex is 7.7). It would be very surprising under these
circumstances if malonate were able to reduce the Mn++ sufficiently to
inhibit the enzyme.
There are several other claims for this mechanism of malonate inhibi-
tion but in all cases there is either inadequate evidence or no evidence at
EFFECTS ON TRICAKBOXYLIC ACID CYCLE 69
all. The examples discussed above indicate the impossibility of establishing
such a mechanism without considering the changes in malonate concentra-
tion or treating the data quantitatively. Despite this lack of positive evi-
dence, there is certainly no doubt but that this type of inhibition can occur
and may be sometimes very important. In the oxidation of pyruvate by
rat heart mitochondria, the Mg++ concentration must fall below 1 mM
before there is any significant decrease in the rate (Montgomery and Webb,
1956 b). We usually used 5 mM Mg++ in the medium so that it would have
required at least 40 mM malonate to produce a detectable inhibition by this
mechanism. Malonate at 50 mM did indeed inhibit around 50% but this
must certainly be due to other actions to a large extent. These experiments
were done with the a-ketoglutarate oxidase blocked by parapyruvate so
that any inhibition of succinate oxidation would not be involved.
EFFECTS OF MALONATE ON THE OPERATION OF THE
TRICARBOXYLIC ACID CYCLE
Malonate is usually assumed to produce its major effects on cellular meta-
bolism and function by disturbing the operation of the cycle* and reducing
the rate of formation of ATP. Malonate has often been used to establish if
the cycle is operative in a tissue or if a particular functional activity is
dependent on the cycle. It is thus important to examine critically the
nature of the cycle block and the effects it may have on the over-all oxida-
tive metabolism. There are two aspects that are especially relevant to this
question. There is the problem of the specificity of action of malonate on
succinate dehydrogenase and this will be considered later. In the present
section we shall assume that the only inhibition is on the oxidation of suc-
cinate and discuss the problems relative to the interpretation of such
a block. Before treating the actual results obtained with malonate, the
nature of the cycle and its responses to inhibition will be outlined.
Some General Principles of Cycle Block
The primary function of the cycle is to incorporate and oxidize acetyl-CoA,
whether this arises from pyruvate, acetate, fatty acids, or elsewhere, and
thus it is particularly important to discuss the effects of malonate block on
this. The situation is relatively clear in suspensions of isolated mitochondria,
in which the concentrations of the cycle intermediates are low and the endo-
genous respiration is generally negligible. Pyruvate, or other substances
giving rise to acetyl-CoA, may be oxidized through the cycle only if some
* In this chapter the term "cycle" will always refer to the tricarboxylic acid cycle
for convenience and other cycles will be designated by their special names.
70 1. MALONATE
cycle intermediate (sparker) is provided to furnish oxalacetate to condense
with the acetyl-CoA. A very small amount of such a sparker may suffice to
initiate the entry of the acetyl-CoA into the cycle, and the cycle will then
perpetuate itself through the continuous formation of oxalacetate, in which
case pyruvate will be completely oxidized to COg and water. Such a system
should be quite sensitive to malonate, because an inhibition of the oxidation
of succinate will reduce the amount of oxalacetate formed and consequently
the amount of acetyl-CoA entering the cycle. On the other hand, if an
approximately molar equivalent of fumarate, malate, or oxalacetate is
initially present with the pyruvate, there wiU be an adequate concentration
of oxalacetate to incorporate acetyl-CoA at a rapid rate, and the process will
not depend on a regeneration of oxalacetate. This system will not be very
sensitive to malonate, because a block of succinate oxidase will not apprec-
iably reduce the amount of oxalacetate present. The first important princi-
ple is, therefore, that the degree of cycle inhibition by malonate will depend
on the source of oxalacetate.
When the cycle is operating in a steady state, the concentrations of
intermediates are low, and oxalacetate is formed just as rapidly as pyruvate
is incorporated, the cycle rate being limited by the entry of acetyl-CoA,
This may be the normal state of the cycle (Krebs and Lowenstain, 1960) but
probably in cells, and certainly in isolated preparations, there are times when
the cycle is not in a steady state. There is an initial rise in citrate concentra-
tion during the oxidation of pyruvate by heart mitochondria in the presence
of malate (Montgomery and Webb, 1956 a), indicating that the tricarboxy-
lates cannot be handled as rapidly as pyruvate can enter the cycle when the
supply of oxalacetate is not limiting. The rate of oxygen uptake is initially
very high, falls to a new level during the first 40 min, maintains this level
for 2-3 hr, and then suddenly fails when the pyruvate is completely utilized.
The first phase occurs when oxalacetate is readily available, and corresponds
to the accumulation of citrate; the rate of oxygen uptake during this period
is not an accurate measure of the rate of operation of the entire cycle —
block of succinate oxidation may have very little effect on the oxygen up-
take because relatively little of the respiration arises from this region of the
cycle. The second steady-state phase should be more sensitive to malonate
because the oxidations of succinate and malate now contribute 2 of the
total of 5 oxygen atoms taken up per molecule of pyruvate. The second
principle is thus that malonate inhibition will sometimes depend on the time
interval during which the oxygen uptake is measured, particularly whether
it is the initial rate or the total oxygen consumed.
Succinate usually accumulates during malonate inhibition (see page 90)
and this will progressively reduce the degree of inhibition due to the com-
petitive nature of the inhibition. Eventually a new steady state may be
reached during which the succinate concentration remains constant. This
EFFECTS ON TRICARBOXYLIC ACID CYCLE 71
will always tend to lessen the effect of malonate and under certain circum-
stances it might effectively overcome the inhibition. The third principle
is that the degree of malonate inhibition will depend on the level of suc-
cinate accumulation in the system studied.
Oxalacetate can often be formed by reactions outside the cycle. Pyruvate
and phosphoenolpyruvate can be carboxylated to oxalacetate in the presence
of oxalacetate decarboxylases or oxalacetokinase (Bandurski, 1955), and
transamination between a-ketoglutarate and aspartate may also give rise
to oxalacetate. In such cases the inhibition of pyruvate oxidation in the
cycle by malonate will be reduced because the incorporation of pjTuvate
will not be dependent only on the regeneration of oxalacetate (Holland and
Humphrey, 1953). Such reactions may occur in isolated mitochondria, as
well as in cells, since in heart mitochondria, where pyruvate alone is not
oxidized at all, the presence of bicarbonate or COg allows a substantial rate
of pyruvate oxidation (Montgomery and Webb, 1956 a), presumably
through the carboxylation of some of the pyruvate to oxalacetate. The
fourth principle is that the degree of malonate inhibition will depend on
noncycle sources of oxalacetate.
Alternate metabolic pathways involving cycle substrates or intermediates
may occur in some tissues. There are many opportunities for the metabolism
of pyruvate, in addition to its oxidation through the cycle, and in the pres-
ence of malonate these pathways may become important. This is particu-
larly true in microorganisms but the ability to decarboxylate pyruvate to
acetate is common to most species and tissues. Thus, in the presence of
high concentrations (50 mJf ) of malonate, pyruvate is quantitatively trans-
formed into acetate by rabbit heart mitochondria (Fuld and Paul, 1952).
An alternate pathway for succinate that would circumvent a malonate
block is the cleavage of succinate (in the presence of NADH, CoA, and
ATP) to 2 acetyl-CoA molecules. This succinate-cleaving enzyme was dis-
covered in Tetrahymena (Seaman and Naschke, 1955) but it is also active
in several rat tissues and in certain bacteria. This reaction will not, of course,
restore cycle activity but it can lead to the formation of acetate or other
products from acetyl-CoA, as well as reduce the concentration of succinate.
Finally, the recently delineated glyoxylate cycle (Kornberg and Krebs,
1957) could bypass that region of the cycle containing succinate oxidase,
malate being formed from isocitrate through the condensation of glyoxylate
and acetyl-CoA, the over-all process being the formation of succinate from
2 PjTuvate + 3/2 O2 -> succinate + 2 CO2 + HgO
pyruvate. This shunt would allow a greater utilization of pyruvate and a
greater oxygen uptake in the presence of malonate than would be the case
with the tricarboxylic acid cycle alone. The glyoxylate cycle has been
found in many microorganisms and there is some evidence for its occurrence
72 1. MALONATE
in certain plants, but its role in animal tissues is as yet unknown (Krebs and
Lowenstein, 1960). As a result of these considerations, the fifth principle
of cycle block is that the degree of inhibition by malonate will depend on
the activity of various alternate pathways and shunts; in addition, it will
depend on what is measured, e.g., oxygen uptake, CO2 production, or
pyruvate disappearance.
There is no doubt, therefore, tha the operation of the cycle and any
ancillary pathways will vary with the experimental or physiological condi-
tions, and that one must expect marked differences in the behavior of the
cycle in different species or tissues. In addition to the factors discussed a-
bove, there are several other reasons for variability in response to malonate;
the different susceptibilities of succinate dehydrogenase to inhibition (see
page 49), the failure of malonate to penetrate readily into cells, and the pos-
sibility that malonate can inhibit other enzymes. The reliability of malonate
as an indicator of cycle activity in a tissue must be evaluated in the light
of these considerations. Certainly the lack of an expected response to mal-
onate cannot be immediately interpreted as indicating the absence of the
cycle, and the production of a significant effect by malonate should be
substantiated by other more direct evidence before the operation of the
cycle is established.
Inhibition of Cycle Substrate Oxidation by Malonate
A summary of some of the effects of malonate on cycle oxidations is
given in Table 1-14. In many cases the concentration of malonate is too
high to act specifically on succinate dehydrogenase, and the results are to
some extent meaningless. It is very difficult to make any generalizations
but malonate concentrations above 10 mM in subcellular preparations must
be looked upon as probably not completely specific, whereas in cellular
systems it is impossible to evaluate the specificity because the intracellular
concentration is not known. In attempting to interpret the inhibitions
observed, it is often necessary to know the pathway of metabolism of the
substrate and how much oxygen is normally taken up per molecule utilized.
For example, when a-ketoglutarate is added to a mitochondrial suspension,
it may be oxidized to fumarate (or malate) taking up 2 atoms of oxygen,
or to oxalacetate taking up 3 atoms of oxygen, or completely taking up 8
atoms of oxygen. If succinate oxidation is completely blocked by malonate,
only 1 atom of oxygen will be taken up. Thus the maximal inhibitions in
these three cases are 50%, 67%, and 87.5%, respectively. Similar reasoning
applies to each substrate and in many cases the exact fate of the substrate
is not known so that it is difficult to estimate what effect might be expected
from malonate. It is also important in this connection, as pointed out in
the previous section, to distinguish between inhibition over an initial short
period of oxidation and inhibition of the total oxygen uptake.
EFFECTS ON TRICARBOXYLIC ACID CYCLE 73
(a) Inhibition of pyruvate oxidation. The oxidation or disappearance of
pyruvate in cellular preparations is usually not depressed very much by mal-
onate at concentrations less than 10 miH , whereas in mitochondrial prepara-
tions the expected degree of inhibition is usually observed. This may be
partly explained by poor penetration into the cells and partly by the alter-
nate pathways that may reduce the importance of the cycle. One type of
correction that can be applied for a more accurate determination of cycle
inhibition by malonate is that used by Speck et al., (1946). In malarial
parasitized erythrocytes, pyruvate is oxidized without the appearance
of acetate, but in the presence of malonate, some acetate in formed. Cor-
rection was made for that pyruvate that went to acetate, since this fraction
of the pyruvate utilization is not dependent on the cycle. The inhibition of
over all pyruvate utilization was 12% but corrected for the acetate it was
31%. In the free parasites, the over all inhibition was 33% and the cor-
rected inhibition 76%. Of course, pyruvate here or in other cells may be
metabolized in other ways, so that the correction for acetate alone may
not give the true cj^cle inhibition, but at least is provides a better value.
Malonate should inhibit the oxidation of pyruvate more strongly when
there is a low concentration initially of oxalacetate or a substance forming
oxalacetate (see page 70). This was shown in homogenates of rat tissues
by Pardee and Potter (1949). In each case the inhibition by 4 mM malonate
Tissue
Substrates
% Inhibition
Heart
Pyruvate
91
Pyruvate + oxalacetate
56
Kidney
Pyruvate
93
Pyruvate -|- oxalacetate
55
Brain
PjTuvate
74
Pyruvate + oxalacetate
47
Liver
Pyruvate
28
Pyruvate + oxalacetate
15
is less when oxalacetate, is present. Since the oxygen uptake was deter-
mined from 10 to 30 min after the start of the experiments and inasmuch as
the concentrations of pyruvate and oxalacetate were 3.5 mM, it is unexpect-
ed that so much inhibition would be exerted when the mixture is present.
This might be due to the decarboxylation of sufficient oxalacetate so that
it was less effective than anticipated, or even at this low concentration
malonate may have been inhibiting some reaction other than the oxidation
of succinate. In rat heart mitochondria we found that 5 mM malonate
inhibits the oxidation of 5 mM pyruvate only about 10% in the presence of
74
1. MALONATE
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EFFECTS ON TRICARBOXYLIC ACID CYCLE 83
5 mM malate. However, when the malate concentration is between 0.1 and
1 mM, the inhibition is close to 50% (Montgomery and Webb, 1956 b).
(6) Inhibition of a-ketoglutarate oxidation. The possible inhibition of a-ke-
toglutarate oxidase by malonate is important not only because of the bearing
it has on the effects of malonate on the operation of the cycle, but also
because malonate has been frequently used to block succinate oxidation
in order to study in particulate systems the oxidation of a-ketoglutarate
uncomplicated by further oxidations. This technique was first proposed by
Ochoa (1944), who showed that high concentrations (25-50 mM) of malon-
ate would allow a-ketoglutarate to be oxidized to succinate in enzyme pre-
parations from cat heart. In four experiments with 50 mM malonate, 0.86
mole of succinate was formed for every mole of a-ketoglutarate utilized,
indicating that even here an appreciable fraction of the succinate formed
is further oxidized. No data were given as to whether these concentrations
of malonate inhibit the utilization of a-ketoglutarate. Slater and Holton
(1954) used 10 mM malonate to study the oxidation of a-ketoglutarate
in heart mitochondria, and it was shown that this concentration does not re-
duce the utilization of a-ketoglutarate, although 20-40 mM does inhibit it.
Malonate was also used to investigate the formation of a-ketoglutarate from
citrate in Micrococcus sodonensis (Perry and Evans, 1960), but the rationale
for this is obscure since malonate by inhibition of succinate oxidation would
not depress the disappearance of a-ketoglutarate. However, at the concen-
tration of malonate used (75 mM), it is quite possible that the a-ketoglu-
tarate oxidase was inhibited.
The oxygen uptake with a-ketoglutarate as the substrate in particulate
preparations from several sources has been shown to be reduced by malonate
as expected if the succinate formed is partially protected from oxidation
(see Table 1-14). Malonate concentrations around 10 mM inhibit 40-60%
in most cases. No definite information on the possible inhibition of the
a-ketoglutarate oxidase can be obtained from such studies.
If the oxidation of a-ketoglutarate stops at fumarate, the system is a
two-step linear chain (neglecting the other reactions involved in the forma-
tion of succinate). An atom of oxygen is taken up in each step:
1/2 Oa 1/2 O.
a-Ketoglutarate -> succinate ->■ fumarate
(1) (2)
SO that a complete and specific inhibition of reaction (2) would result in a
maximal inhibition of 50% with respect to the oxygen uptake. However, in
case the first reaction is much faster than the second, malonate would not
inhibit the initial rate of the reaction, even though it inhibited the final total
oxygen uptake. Therefore, the inhibition may theoretically vary from 0 to
84 1. MALONATE
50%, depending on the relative rates of the reactions and the period during
which the oxygen uptake is measured. If the fumarate is further oxidized,
more oxygen will be consumed and the inhibition by malonate may be
greater than 50%. Furthermore, it is essentially impossible to inhibit the
succinate dehydrogenase completely, especially as succinate will accumulate
and progressively overcome the inhibition.
The use of malonate to study a-ketoglutarate oxidation in mitochondria
involves the assumption that malonate does not significantly affect the a-
ketoglutarate oxidase directly. Unfortunately, no investigations of the in-
hibition of a-ketoglutarate dehydrogenase by malonate have been report-
ed, and thus it is difficult to compare the sensitivities of the two dehydro-
genases. Several studies have determined the effects of malonate on the
disappearance of a-ketoglutarate (a-KG) during periods when the oxygen
uptake is depressed, and it is rather strange that some effect, either positive
or negative, has always been reported. The pertinent data have been sum-
marized in the accompanying tabulation, and it is seen that an inhibition
Preparation
Malonate
(mM)
0
0
Inhibition of
O2 uptake
a-KG utilization
Streptomyces coelicolor
10
41
Stim 9
Pea seedling mitochondria
10
66
21
Blowfly sarcosomes
10
10
Stim 25
Heart mitochondria
10
33
Stim 8
Rat brain homogenate
3.3
59
29
of a-ketoglutarate disappearance is observed in some cases and a stimulation
in others. The oxidation of a-ketoglutarate depends on Mg++ and it is
possible that the differences are related to the degree of Mg++ requirement
and the concentrations of Mg++ used in the assay media. Price (1953)
found that even 1 mM malonate would inhibit a-ketoglutarate utilization
in pea seedling mitochondria and that 30 raM had a very marked effect
(60% inhibition). This is not a competitive type of inhibition because
increasing the a-ketoglutarate concentration does not lower the inhibition,
and even increases it somewhat. The possibility of Mg++ reduction was
explored and it was found that the calculated drop in the Mg++ concen-
tration could not have been responsible for the inhibition. Furthermore, it
was not possible to reverse the inhibition by increasing the Mg^^ concen-
tration from 1 mM to 4 mM. An inhibition by a Mg-malonate complex
was considered to be the most likely explanation, but yet no inhibition
was seen when the concentration of Mg++ in the medium was reduced to
zero, although the enzyme was still partially complexed with Mg+"'". What-
EFFECTS ON TRICARBOXYLIC ACID CYCLE 85
ever the explanation, it is obvious that in such mitochondria malonate
could not be used to isolate a-ketoglutarate oxidation.
Another quantitative study on the effects of malonate on a-ketoglutarate
oxidation was made by Grafflin et al. (1952), who were attempting to find
a good assay system for a-ketoglutarate oxidase in rabbit kidney homogen-
ates. They concluded that the use of malonate is unsatisfactory and aban-
doned this procedure. The difficulty lies particularly in the inability,
except at high malonate concentrations (around 30 mM), to inhibit com-
pletely the oxidation of succinate, as determined from the total oxygen
uptake compared with the theoretical value for a one-step oxidation of
a-ketoglutarate. Although no evidence on the effect of malonate on the
a-ketoglutarate oxidase was presented, it would be surprising if concentra-
tions of malonate above 20 mM had no effect. Lewis and -Slater (1954)
also remarked that even in the presence of 10 milf malonate, the oxygen
uptake from the oxidation of a-ketoglutarate greatly exceeds the disap-
pearance of a-ketoglutarate, .lO/.Ja-KG ratios generally being above 2,
in blowfly sarcosomes. Also, the oxygen uptake over 35-45 min is depressed
only 10% at this concentration of malonate.
It is difficult to understand why the oxidation of succinate is so resistant
to malonate under these circumstances. Taking the values of Kj that
have been found in mammalian tissue studies, malonate at 10 mM should
inhibit well over 95% even at succinate concentrations that might occur
experimentally. For example, in beef heart preparations, where K^ is about
0.04 WlM, 10 vnM malonate would inhibit over 99% at a succinate concen-
tration of 2 mM. It may be that in the intact system the oxidation of endo-
genously formed succinate from a-ketoglutarate via succinyl-CoA is ki-
neticaUy different than the oxidation of exogenous succinate, or that local
concentrations of succinate can reach much higher levels than predicted
on the basis of over all analyses. One must conclude at least at the present
time that the specific inhibition of succinate oxidation, even in these rela-
tively simple systems, is generally impossible.
This problem has been approached recently by Jones and Gutfreund
(1964), who experimentally determined the steady-state concentrations of
succinate in guinea pig liver mitochondria during the oxidation of a-keto-
glutarate. The O2 uptake of uninhibited mitochondria is 40-45% due to
a-ketoglutarate oxidation, 40-45% due to succinate oxidation, and 10-20%
due to other oxidations. The effects of malonate on the oxidation of exoge-
nous succinate were compared with the effects on the oxidation of succin-
ate-C^* formed from a-ketoglutarate-C^^. The rate of utilization of a-keto-
glutarate is not altered up to 8 mM malonate. The steady-state succinate
concentration by total analysis is 0.04 mM, and this is not affected by
malonate until its concentration is higher than 0.04 mM; half the succinate
formed from a-ketoglutarate accumulates with 0.7 mM malonate, and
86 1. MALONATE
80% accumulates at 8 mM malonate. The Og uptake due to the oxidation
of succinate formed from a-ketoglutarate is reduced 50% by 0.2 mM mal-
onate. The data indicate that endogenously formed succinate is at a much
higher concentration than that determined by total analysis. The inhibition
studies indicate the steady-state succinate concentration in the mitochon-
dria to be 4.6-13 mM and the isotopic studies suggest concentrations ex-
ceeding 4 mM. Such high succinate concentrations would protect the suc-
cinate dehydrogenase and result in less malonate inhibition than expected.
Since there appear to be no permeability barriers in the mitochondria to
succinate, the authors suggested that there is a spatial relation between
the succinate dehydrogenase and the enzyme forming the succinate. If
this is true, it raises the interesting possibility that certain enzymes are
specific not only for their substrates but also for other enzymes with
which they interact to form functional metabolic complexes.
(c) Effects on tricarboxylate utilization. Malonate has variable affects on
the oxidation of citrate and isocitrate, often inhibiting rather well, but
sometimes having little effect or actually stimulating (Table 1-14). The
relationship between malonate inhibition and isocitrate oxidation is prob-
ably complex in most instances. There seems to be minimal direct inhibi-
tion of isocitrate dehydrogenase, although data are lacking. Hiilsmann
(1961) showed that malonate at 11 mM reduces quite potently the utiliza-
tion of isocitrate by rabbit heart mitochondria when acetate or /?-hydroxy-
butyrate is the additional substrate (these accelerating the utilization of iso-
citrate). The stimulation of isocitrate utilization by these substrates was
claimed to be due to the formation of acetoacetyl-CoA, an intermediate in
fatty acid synthesis, and this alters the oxidation-reduction states of NAD-
NADH and NADP-NADPH through the transhydrogenase reaction,
isocitrate oxidation rate being dependent on the concentration of NADP.
Kasamaki et ol. (1963) showed that malonate inhibits citrate and isocitrate
oxidations in Proteus vulgaris much less when NADP is added. Chappell
(1964 a) suggested that the effects of malonate on tricarboxylate utilization
in rat liver mitochondria are due to the block in the formation of malate
from succinate, malate being necessary for the oxidation of isocitrate, since
it provides a means for reoxidation of NADPH mediated through the
transhydrogenase and malate dehydrogenase. We see here a possibly im-
portant control in the operation of the cycle and how malonate can exert
effects indirectly on steps distant from the succinate oxidation reaction,
{d) Effect of cycle substrate concentration on the inhibition. Some claims
have been made that increases in the citrate concentration will tend to over-
come inhibition of the oxygen uptake by malonate, and sometimes this
has been attributed to a competitive effect, it being assumed that the higher
concentration of substrate will give rise to a higher succinate level. Thus
EFFECTS ON TRICARBOXYLIC ACID CYCLE 87
Laties (1953) found that increasing the citrate from 1 raM to 10 mM
would reduce the inhibition by 10 mM malonate from around 52% to zero in
cauliflower homogenates, and Pierpoint (1959) reported that a 10-fold rise
in citrate concentration brings about a decrease from 62 to 45% in the
inhibition by 21 mM malonate in tobacco leaf mitochondria. Laties pointed
out that this could not be due to a competitive effect because of an increased
production of succinate, since at the malonate concentration used the
oxidation of 10 mM succinate is completely inhibited. However, if what
was suggested in the previous section regarding the difficulty in the inhi-
bition of the oxidation of endogenously produced succinate is valid, data
on the inhibition of exogenous succinate may not be applicable. Laties felt
that the explanation might lie in the interference between electron-trans-
port systems, so that when citrate concentration is high, the contribution
made by succinate oxidation to the total respiration is less. Clear-cut effects
of concentration on inhibition have not been observed with a-ketoglutarate
(Grafflin et al., 1952; Pierpoint, 1959), pyruvate (Smyth, 1940), acetate
(Jowett and Quastel, 1935 c), or malate (Pierpoint, 1959).
In experiments of this type, it is well to remember that the pattern of
oxygen uptake may change with the concentration of the substrate. The rela-
xO yO
S -> succinate ->• P
tive amounts of oxygen taken up before and after the succinate step may be
altered by substrate concentration due to factors previously discussed in
this chapter. Thus the ratio x/y in the above equation will vary and hence
the effect of an inhibitor of succinate oxidation. If the inhibition on succinate
oxidation is i and the ratio xjy = r, the over all inhibition on the total oxy-
gen uptake will be ijil + r), so that anything that changes r will change the
inhibition. If high concentrations of the substrate produce an accumulation
of some intermediate (as citrate rises when pyruvate enters the cycle rapidly),
r will increase, at least over the initial period, and the inhibition will be
less at lower substrate concentrations.
(e) Stimulation of cycle substrate utilization by malonate. In a number of
cases there is clearly a stimulation of the utilization of pyruvate, acetate,
or citrate by malonate. Sometimes this is recognized in an increased oxygen
uptake but occasionally the disappearance of the substrate is accelerated
while the oxygen uptake is inhibited. Often this effect is very marked. In
the mycelia of Ashbya gossypii, oxygen uptake due to addition of acetate is
stimulated 48% by 4 m.M malonate, whereas 40 m.M malonate inhibits
53% (Mickelson and Schuler, 1953). The increase in the respiration from
pyruvate in bull sperm by 10 mM malonate is almost as great (Lardy and
Phillips, 1945). Table 1-14 cites a number of other instances.
88 1. MALONATE
The mechanisms for such stimulations must vary with the particular sub-
strate used, but it may be useful to suggest some possible ways in which
malonate could produce this apparently anomalous effect. (1) If the prepara-
tion has an active oxalacetate decarboxylase, this may reduce the concentra-
tion of oxalacetate for condensation with acetyl-CoA. Malonate is able to
inhibit this enzyme in some cases (Table 1-12) (Pardee and Potter, 1949),
in which case oxalacetate may be protected so that cycle entry of acetyl-
CoA is facilitated. (2) If ADP concentration is low and ATP concentration
high normally in the preparation, malonate by inhibiting certain phases of
the cycle might increase ADP concentration and thus stimulate electron
transport in other oxidative processes by providing more phosphate ac-
ceptor. (3) If there is competition between the different oxidative reactions
in the cycle for some common coenzyme or cofactor (e.g. NAD, NADP, or
Co A), inhibition of some oxidations by malonate might allow these factors
to be used more readily by other systems. Such competition between the
pyruvate and a-ketoglutarate systems has been suggested in heart mito-
chondria (Montgomery and Webb, 1956 a). (4) If malonate is metabolized by
the preparation, it is possible that a product would accelerate the utiliza-
tion of some cycle intermediate. In some organisms malonate can form acetyl-
CoA and acetate, as well as other products. (5) In intact cells, malonate
might increase the permeability of the cell membrane so that certain sub-
strates, such as pyruvate, citrate, or a-ketoglutarate, could enter more
readily. (6) By chelation with inhibitory metal ions that may occur in the
preparation, malonate might accelerate the rates of certain reactions. All of
these mechanisms are purely hypothetical, since in no case has the actual
mechanism been established.
Intracellular Concentrations of Cycle Intermediates
Interpretation of intracellular inhibition by malonate and other inhi-
bitors acting on the cycle should ideally involve in many cases a knowledge
of the concentrations of certains intermediates. Data collected for different
types of cells are shown in Table 1-15. These figures were calculated on
the basis of intracellular water contents. However, it is likely that these
substances are not distributed homogeneously throughout the cell water.
There is evidence that some intermediates may occur within the mito-
chondria at different concentrations than in the surrounding medium. Thus
the mitochondria/medium ratios for sheep kidney mitochondria under
certain conditions were found to be: pyruvate 0.84, fumarate 7.42, a-keto-
glutarate 1.0, citrate 0.83, and oxalacetate 0.13 (Bartley and Davies,
1954). Furthermore, the values given in the table are all abnormal since
truly normal tissues were not used. The rats were fasted for 24 hr while
the suspensions of E. coli and yeast were metabolizing acetate rather than
a more normal substrate. In normal rat tissues the concentrations may well
EFFECTS ON TRICARBOXYLIC ACID CYCLE
89
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90 1. MALONATE
be higher, and in yeast metabolizing sugar the values may be lower. One
value for oxalacetate in normal rat liver is available (0.036 mikf) and in
this case fasting for 24 hr did not alter this appreciably (Kalnitsky and
Tapley, 1958).
These values do not represent a thermodynamic equilibrium based on dif-
ferences in free energy, but rather a dynamic or kinetic equilibrium, depend-
ing mainly on the relative rates of the cycle reactions and competing process-
es. The higher concentration of fumarate compared to malate in rat tissues
illustrates this because in a thermodynamic equilibrium there would be
about one-fourth the concentration of malate. It is interesting that the values
differ so widely from tissue to tissue and certainly this must be one factor in
determining the different responses to malonate or other competitive inhi-
bitors. The levels of succinate are generally low, except in yeast, but as
pointed out above the concentrations within the mitochondria or at the re-
gion of the active center of succinate dehydrogenase may well be higher.
The effects of malonate on the concentrations of these intermediates in cells
will be taken up in the following section.
Analyses of plant tissues have not been presented here because there is
some doubt as to the significance of the figures. Most plants contain large
amounts of organic acids, including the cycle intermediates. Beevers (1952)
postulated that the cj^cle in plants is less readily blocked than in animal
tissues because of these high concentrations of succinate and other cycle
intermediates. This could well be an important factor, but actually the
concentrations of these acids in the plant cytoplasm are not known in
most cases, total analyses including the vacuolar fluid, which is often of
greater volume than the cytoplasm and contains most of the organic acids.
There is another way by which these plant acids could protect against
malonate. The presence of large amounts of fumarate or malate, or of any
substance capable of forming oxalacetate, would allow pyruvate to be incor-
porated into the cycle even in the state of complete block of succinate oxi-
dation. Examples of the overcoming of malonate inhibition by fumarate
and malate will be presented shortly.
ACCUMULATION OF SUCCINATE DURING MALONATE
INHIBITION
An effective inhibition of succinate oxidation should lead to a rise in
the concentration of succinate under conditions in which succinate can still
be formed. Such accumulation of succinate has been frequently observed
and some of the more quantitative results are summarized in Table 1-16.
In addition to the examples in the table, accumulation of succinate has
been reported in the following species and tissues: Shigella (Yee et al.,
1958), Nocardia (Cartwright and Cain, 1959), Aspergillus (Shimi and Nour
ACCUMULATION OF SUCCINATE 91
El Dein, 1962), tobacco leaves (Vickery, 1959; Vickery and Palmer, 1957),
potato slices (Romberger and Norton, 1961), avocado mitochondria (Avron
and Biale, 1957), pea leaf particulates (Smillie, 1956), barley roots (Laties,
1949 b), Colpidium (Seaman, 1949), Trypanosoma (Bowman et al., 1963),
carp liver mitochondria (Gumbmann and Tappel, 1962 b), rat heart ho-
mogenates (Lehninger, 1946 b), rat liver slices (Elliott and Greig, 1937),
human heart slices (Burdette, 1952), ascites carcinoma cells (Dajani et al.,
1961), and in many tissues of rats and rabbits (Busch and Potter, 1952 a;
Forssman, 1941). In the experiments leading to the results in Table 1-16, the
preparations were incubated for one to several hours with malonate and the
succinate analyzed at the and of the incubation, so that the rates of succinate
formation at any time are difficult to evaluate, and may well have been
greater initially. The over all succinate concentrations may be estimated
from the volumes in which the expriments were run and in most cases the
final succinate concentrations range between 0.5 and 2.5 mM.
Several points are brought out by the results in Table 1-16. It is seen
that succinate can be formed from essentially all the cycle substrates and
intermediates in the presence of malonate. Rapid rates are found when
oxalacetate or some substance forming oxalacetate is added with pyruvate,
as would be expected, because in the absence of a source of oxalacetate, the
malonate would reduce the incorporation of pyruvate into the cycle and
hence the rate of formation of succinate. It may be noted in some cases
that, in the absence of added substrates or malonate, some succinate ac-
cumulates (yeast, Avena coleoptile, spinach leaves, and dog heart), which
implies that under the experimental conditions succinate is formed more
rapidly than it can be oxidized. This is somewhat surprising because it is
usually assumed that the activity of succinate oxidase is quite high in
most tissues. The possibility of the accumulation of sufficient oxalacetate
to inhibit succinate dehydrogenase when little acetyl-CoA is available
cannot be ignored. This phenomenon is also evident in the analyses for
succinate given in Table 1-15. The interesting effects of malonate concen-
tration are seen in two investigations. In spinach leaves, the maximal suc-
cinate accumulation occurs at malonate concentrations around or below
50 milf; at higher concentrations, the malonate is apparently acting on
other enzymes in the cycle and reducing the rate of formation of succinate.
Likewise, in brain minces, the high concentration of 200 mM malonate is
seen to depress succinate accumulation.
Quantitative conversion of cycle substrates to succinate in the presence
of malonate is generally not observed. In fact, in most cases in which the
disappearance of substrate was determined simultaneously with the for-
mation of succinate, only a small fraction appeared as succinate. For exam-
ple, Speck et al. (1946) found in malarial parasitized erythrocytes that
only 22% of the pyruvate utilized in the presence of 20 mM malonate was
92
1. MALONATE
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ACCUMULATION OF SUCCINATE
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96 1. MALONATE
recoverable as succinate. The highest conversion efficiency was observed by
Krebs and Eggleston (1940) in pigeon muscle brei, where 75-85% of the
pyruvate utilized in the presence of fumarate and 12.5 mM malonate went
to succinate. Complete conversion to succinate would not, of course, be ex-
pected unless one could inhibit succinate oxidation completely and specifi-
cally, which in most instances cannot be done.
Factors Determining Succinate Accumulation
The effects of inhibition on the concentrations of intermediates in multi-
enzyme systems have been treated in Chapter 1-7. Some of the most impor-
tant factors involved in malonate inhibition will be summarized.
(I) Degree of inhibition of succinate oxidase: hence, the concentration
of malonate, the affinity of the enzyme for malonate, and the ability
of malonate to penetrate if the preparations are cellular.
(II) Rate of formation of snccinate: this will depend primarily on the availa-
bility of cycle substrates and their concentrations.
(III) Action of malonate on enzymes other than succinate dehydrogenase:
such actions may slow down the formation of succinate, as discussed
above.
(IV) Other pathways of succinate metabolism: several reactions of succinyl-
CoA or succinate are known and these would tend to prevent accumula-
tion.
(V) Diffusion of succinate from cells: when only cells or tissues are analyzed
for succinate, intracellular accumulation will be reduced by the loss of
succinate into the medium or the blood.
(VI) Time after addition of malonate: although this has never been studied,
it is probable that the succinate concentration will follow characteristic
time courses in each case, in some cases perhaps decreasing after a peak
level has been reached.
Another factor, about which nothing is known, is the possible effect of
rising succinate concentration on the reactions forming succinate. The
oxidation of a-ketoglutarate forms succinyl-CoA, and succinate can arise in
at least three different ways from succinyl-CoA.
P-enzymes
Q -Ketoglutarate — >- sue cinyl - Co A *- sue einate
transaeylases
ACCUMULATION OF SUCCINATE 97
What effects succinate concentration might have on these reactions are
unknown, but some inhibition during succinate accumulation is possible,
although a simple backing-up of the a-ketoglutarate -^ succinate reaction
would be thermodynamically unlikely.
The relative potential rates of a-ketogluti^rate oxidase and succinate
oxidase under normal intramitochondrial conditions are not known, but the
oxidation of succinate is certainly one of the most rapid reactions seen in
mitochondrial suspensions. It may be that there is no accumulation of suc-
cinate in the cycle under physiological conditions, and that the small
amounts of succinate found in tissues do not truly indicate the situation in
the regions of succinate oxidase. Succinate oxidase is, perhaps, the one
enzyme that has never been considered as normally limiting the cycle rate.
If the maximal rate of succinate oxidation is much higher than the rate at
which succinate can be formed, it would require a fairly high inhibition of
the oxidase before succinate accumulates markedly. For example, if the rate
of succinate formation is one-tenth the rate at which it can be oxidized,
90% inhibition of the succinate oxidase would make the rates equivalent,
and the succinate concentration would not rise very much (probably not
more than 0.02 0.05 n\M). Under any likely conditions, calculations from
Eqs. 1-7-8 and 1-7-9 make it clear that succinate oxidation must be inhibited
fairly strongly to produce a significant succinate accumulation. The com-
m.on assumption that succinate must accumulate rather quantitatively
when sufficient malonate has been added to inhibit succinate oxidase
75-90% is thus unjustified. If alternate pathways for succinyl-CoA or suc-
cinate exist, the accumulation of succinate would be even less evident.
Several instances of failure of succinate to accumulate during malonate
inhibition have been reported. Hanly et al. (1952) found that in only two
of six experiments with carrot root slices did succinate accumulate, and
in these the rise was insignificant. One might suspect a lack of penetration,
but 15-50 mM malonate was used at pH 4; under these conditions, malonate
depressed respiration strongly. Weil-Malherbe (1937) found no succinate
accumulation in guinea pig brain slices with malonate 4-40 mM, respiration
being markedly reduced at the higher concentrations. A depression of suc-
cinate level in Streptomyces due to 10 vaM malonate was noted by Coch-
rane (1952), with either malate or citrate as substrate. Since the incubation
was 16 hr and the pH 5.1-5.4, it is possible that a nonspecific acid damage
from malonic acid penetration was responsible for the cycle depression, or
it might be that in this organism malonate is not specific at 10 mM, or,
as Cochrane suggested, succinate may not be formed via the cycle. The fail-
ure of succinate to accumulate, even when the respiration is suppressed by
malonate, is difficult to explain except on the basis of actions other than
on succinate dehydrogenase.
98 1. MALONATE
Succinate Accumulation in the Whole Animal
Some of the most interesting and suggestive experiments on succinate
accumulation resulting from malonate inhibition have been performed with
whole animals and, although such work is often difficult to interpret in a
quantitative fashion, the results have demonstrated that malonate can
partially block the succinate oxidase in various tissues of the living animal.
Such inhibition has obvious implications for developments in the study of
drug actions and chemotherapy. The first work of this type was done by
Krebs et al. (1938), who determined the urinary excretion of succinate, cit-
rate, and a-ketoglutarate following injections of cycle intermediates and mal-
onate into rats and rabbits. Some of their results are shown in Table 1-17.
The effects on citrate and a-ketoglutarate will be discussed later (see page
104). Although malonate increases the succinate excretion some 5-fold,
only 2.9% of the injected fumarate is recovered as succinate compared to
0.6% in the controls. The injection of 10 millimoles of a substance into a
rabbit will lead to a maximal extracellular concentration of approximately
30 TdM, so that reasonably high concentrations of malonate were probably
achieved. The effect of the malonate had mainly disappeared after 24 hr
due to the excretion and destruction of the malonate. An almost 10-fold
increase in the urinary succinate was seen in the more recent experiments
of Thomas and Stalder (1958), in which 3.7 millimoles/kg of sodium mal-
onate were fed to rats, the succinate over a 40-hr period rising from a
control value of 2.35 mg to 28.0 mg.
The blood concentration of succinate is increased in rabbits following
the injection of malonate (Forssman, 1941). Intravenous injection of 2.8
millimoles/kg of malonate leads to a slow rise in the blood succinate to
around 0.20 mM at 3 hr, while injections of 3.5-5.1 millimoles/kg give
levels as high as 0.77 mM. A lethal dose of 8.25 millimoles/kg produces
death in 35 min and at the time of death the succinate concentration is
1.1 mM. The lower doses produce no obvious effects on the animals. The
normal values for blood succinate are about 0.025 milf.
The succinate found in the urine and blood in these studies originated
mainly in the tissues of the animals. Are malonate inhibition and succinate
accumulation especially related to a particular tissue, or do all the tissues
contribute to the metabolic disturbance? Can differences in the metabolic
patterns of the various tissues be demonstrated by their responses to the
administration of malonate? How do tumors compare with normal tissues in
their susceptibility to malonate? It was to answer such questions as these
that Busch and Potter (1952 a, b) at the McArdle Memorial Laboratory
at Wisconsin undertook their excellent series of studies on the accumulation
of succinate in various tissues of rats following injections of malonate.
Analyses for malonate and succinate were made by anion exchange chrom-
atography (Busch et al., 1952) at various times after the subcutaneous
ACCUMULATION OF SUCCINATE
99
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1. MALONATE
injection of 12 millimoles/kg of malonate (Fig. 1-11). Maximal concentra-
tions of malonate are reached in most tissues 1-2 hr after the injection and
the time course of the succinate levels is well correlated with that of malo-
nate. The tissue succinate concentration is linearly related to the malonate
concentration, except in the Flexner-Jobling tumor (Busch and Potter,
Fig. 1-11. Tissue levels of malonate and succinate
in rats injected with sodium malonate (12 milli-
moles/kg). (From Potter et al., 1952).
1952 a) (Fig. 1-12). The slopes of the lines give some measure of the degree
of malonate effect in the particular tissue, but several factors are involved
so that they are not quantitative indications of succinate oxidase inhibition.
The urinary excretion of malonate, succinate, and citrate is shown in Fig.
1-13. The malonate and succinate levels in several normal tissues and tu-
mors following 24 millimoles/kg malonate subcutaneously are given in
Table 1-18. If the figures in the table are multiplied by approximately
0.7, one will obtain the millimolar concentrations in the cell water, except
for kidney and blood, the former tissue having extracellular fluid high in
succinate and malonate.
The ratios of (succinate)/(malonate) in the tissues give essentially the
slopes of the lines in plots such as Fig. 1-12, except for tumors where a
ACCUMULATION OF SUCCINATE
101
Liver
/
Tumor
y^ Kidney
/
^^
Q\oo^^^^^^^
0 10 20
Malonote (millimoles/ kg) -
Fig. 1-12. Relationships between malonate and suc-
cinate concentrations in the tissues of rats injected
with malonate. (From Busch and Potter, 1952 a).
Time(hr)-
FiG. 1-13. Urinary excretions of malonate, succinate, and citrate
following the injection of 12 millimoles/kg sodium malonate into
rats. (From Busch and Potter, 1952 a).
102
1. MALONATE
Table 1-18
Tissue Succinate and Malonate Concentrations Following Injections of
Malonate in Rats "
Tissue
Succinate
Malonate
Ratio
Control
After
malonate
Control
After
malonate
succinate
malonate
Spleen
0
12.8
0
12.8
1.00
Liver
2.2
12.0
2.4
14.2
0.85
Brain
0
2.0
0
2.8
0.71
Thymus
0
9.0
0
21.0
0.43
Kidney-
0.6
16.4
0.3
44.0
0.37
Lung
0
5.6
0
16.0
0.35
Muscle
1.0
3.0
2.0
8.8
0.34
Heart
0
5.0
0
18.4
0.27
Blood
0
2.6
0
24.6
0.11
Tumors
Flexner
-Jobling
carcinoma
0
9.2
0
17.6
0.52
Walker
256 carcinoma
0
8.0
0
5.2
1.54
Jensen
sarcoma
0
8.8
0
8.0
1.10
Hepatoma
0
6.0
0
12.0
0.50
Papilloma
0
4.4
0
4.4
1.00
Average
0
7.3
0
9.4
0.78
" The figures are in /<equivalents/g wet weight of tissue. Malonate was injected
subcutaneously at a dosage of 12 millimoles/kg, and after 1 hr a similar amount was
again injected; 1 hr following the last injection the animals were sacrificed. (From
Busch and Potter, 1952 b.)
linear relationship may not be followed. These ratios indicate the amount
of succinate formed per unit concentration of malonate and do not relate
directly to the degree of inhibition of succinate oxidase but more to the
ability of the tissue to form succinate in the presence of the inhibition.
As discussed previously, for the same degree of block, succinate will be form-
ed at greatly different rates in different tissues, depending on the succin-
ate-forming substrates available and the activity of the pathways leading
to succinate. The concentrations of malonate in the tissues vary greatly
and this must be a reflection of the differing permeabilities of the tissues to
malonate. It may be noted that the concentration in the brain is quite low,
ACCUMULATION OF SUCCINATE 103
a phenomenon seen with most ionic substances, and this must account for
the poor accumulation of succinate in this organ and the relative lack of
effect of malonate on central nervous system function.
The low concentration of succinate in the blood is interesting since it
implies that succinate does not leave the tissues readily. The conclusion
that must be reached is that the rate at which succinate diffuses out of the
tissues into the blood is slower than the rate of renal excretion of the
succinate. Substances that are not resorbed by the renal tubules are excret-
ed rapidly and their concentrations in the blood can be maintained at a
low level despite a continuous influx. Nevertheless, it shows that succinate
leaves the tissue cells rather slowly under physiological conditions. The slow
penetration of malonate into the tissues is suggested by the fact that the
peak levels in the blood occur around 30 min after administration whereas
the peak levels in liver and tumor occur 30 min later (Fig. 1-11), the kid-
ney concentration paralleling the blood levels because of the excretory
function of this organ.
The degree of succinate accumulation does not necessarily reflect the
cycle activity in a tissue. For one reason, succinate can often be formed
from other pathways. In certain tissues the amino acid content rises mark-
edly (e.g. + 215% in thymus and + 160% in spleen) during malonate
inhibition, while in others, especially the Flexner-Jobling carcinoma, the
amino acids decrease as the succinate increases. It is likely in the latter
tissues that some of the succinate is derived from amino acids, probably
mainly glutamate. Thus the cycle activity in this tumor may be quite low
and the normal accumulation of succinate due to other sources for the suc-
cinate. The other tumors do not show such marked decreases in amino acid
content and this was attributed to their greater necrosis. It may be recall-
ed that the incorporation of acetate by Flexner-Jobling tumor is slower
than in most tissues and very little labeled succinate is formed from la-
beled acetate (Busch and Potter, 1953), indicating a low degree of cycle
activity. We have seen that this tumor also differs from normal tissues in
the nonlinearity of the plot of tissue succinate against tissue malonate
(Fig. 1-12). At low levels of malonate the ratio (succinate)/(malonate) is
near 3 but at high concentrations diminishes to 0.5. This means that as
the malonate concentration rises, the ability of the tumor to accumulate
succinate decreases. This is the type of curve expected if malonate at higher
concentrations is inhibiting the reactions forming succinate. If the supply
of cycle substrates in this tumor is low, a relatively small block of the cycle
might reduce the formation of succinate through the cycle to zero. The
slope approaches that of the blood, and it is possible that above the inflection
point the slow rise in succinate may be due only to the rise in the blood.
This type of investigation could well be applied to other inhibitors
and certain chemotherapeutic agents. First, one is able to correlate tissue
104 1. MALONATE
concentrations of inhibitor with the metabolic disturbance produced.
Second, the inhibition occurs under physiological conditions, rather than
in slices or minces or other preparations in which the cell metabolism may
be very abnormal. Last, one is able to compare the different tissues with
respect to their metabolic patterns and perhaps determine some of the
reasons for the selective actions of inhibitors or drugs. These methods of
investigation, called "m vivo metabolic blocking techniques" by Busch
and Potter, if applied properly, would help to provide a more rational
basis for development in chemotherapy and the selective depression of
tumor growth.
ACCUMULATION OF CYCLE SUBSTRATES
OTHER THAN SUCCINATE
Specific inhibition of succinate oxidase would be expected to lead to the
accumulation of succinate but of no other cycle intermediates, because
the free energy differences between them are of such magnitude that no
backing-up from succinate would be anticipated. When other members of
the cycle are found to accumulate in the presence of malonate, it is generally
considered to be evidence that either the action of malonate is not specific
or that secondary reactions are proceeding. Malonate has been shown many
times to cause an accumulation of certain cycle intermediates, especially
citrate and a-ketoglutarate, in cell suspensions, slices, and whole animals.
The nature of these effects will first be summarized and then some possible
mechanisms will be considered.
Accumulation of Citrate
The administration of malonate to dogs (Orten and Smith, 1937), rabbits
(Krebs et al., 1938),- and rats (Busch and Potter, 1952 a) leads to an increased
urinary excretion of citrate (Table 1-17 and Fig. 1-13). There is also a rise
in plasma citrate following injections of malonate in rabbits (Forssman,
1941) and dogs (Stoppani, 1946). Tissue citrate also rises in mice injected
with 10 millimoles/kg malonate: kidney (16 to 20), heart (40 to 70), liver
(5 to 10), and diaphragm (70 to 225) (values in milligrams per kilogram
wet weight) (Chari-Bitron, 1961). Brain, however, shows no increase in cit-
rate, perhaps due to the poor penetration of malonate. Some accumula-
tion of citrate has also been observed in suspensions of Ashbya gossypii
mycelia metabolizing acetate and oxalacetate (Mickelson and Schuler,
1953), Schizophyllum commune mycelia metabolizing pyruvate and malate
(J. G. H. Wessels, 1959), and Ehrlich ascites tumor cells metabolizing fu-
marate (Kvamme, 1958 c) in the presence of malonate. Thus this phenom-
enon is widespread, occurring in different types of organism and under a
variety of conditions.
SUBSTRATES OTHER THAN SUCCINATE 105
On the other hand, 4-10.5 millimoles/kg of malonate injected intra-
venously into rabbits does not increase plasma citrate appreciably (Forss-
man and Lindsten, 1946), and a number of reports have indicated a depres-
sion of citrate formation by malonate. Eat brain and liver homogenates oxi-
dizing oxalacetate, or pyruvate and oxalacetate, form less citrate in the
presence of 10 mM and 30 milf malonate, respectively, this being attributed
to an inhibition of oxalacetate decarboxylase (Pardee and Potter, 1949).
The formation of citrate from acetate in yeast is inhibited 73% by 17 mill
malonate, while simultaneously succinate accumulates markedly (Barron
and Ghiretti. 1953). The incorporation of C^^ from glucose into citrate in
potato tuber slices is also depressed 71% by 50 mM malonate (Table
1-19) (Romberger and Norton, 1961). Although there is an increase in
citrate in excised tobacco leaves during culture with malonate, this increase
is generally less than in the controls, so that this probably represents an
inhibition of citrate formation (Table 1-20) (Vickery, 1959; Vickery and
Palmer, 1957). Finally, malonate inhibits the formation of citrate from a
variety of substrates in kidney and testis breis, the effects being surprisingly
large for the reasonable concentrations of malonate used (Table 1-21)
(Hallman, 1940). It is, therefore, evident that citrate levels may be affected
by malonate in a variety of ways, depending on the malonate concentration,
the type of preparation, and the conditions of the experiment. It is not dif-
ficult to explain the falls in citrate level brought about by malonate, since
this could arise either by a depression of succinate oxidation (reducing the
rate of entry of acetyl-CoA into the cycle) or inhibitions of other reactions
(such as the condensation of oxalacetate and acetyl-CoA), especially at the
high malonate concentrations often used. It is, on the other hand, difficult
to interpret the accumulation of citrate and to this end we must direct
our efforts, although only suggestions can be offered because of the lack
of sufficient data.
Citrate is being formed and metabolized continuously and thus, generally
speaking, a rise in the citrate level implies an inhibition of citrate utili-
zation or an acceleration of its formation, or both. Although there is little
evidence for a direct affect of malonate on isocitrate utilization, we have
noted that an inhibition of succinate oxidation can interfere with isocitrate
oxidation by depletion of NADP mediated by a fall in malate concentration
(Jones and Gutfreund, 1964). This could certainly contribute to the accu-
mulation of the tricarboxylates in some instances.
The inhibitions of citrate oxidation in Table 1-14 can be mostly explained
on the basis of a block at the succinate oxidase step. On the other hand,
there is certainly no evidence that the formation of citrate via the cycle
can be stimulated by malonate, most data pointing instead to a depression
if there is any effect. It would appear that effects of malonate on the cycle
alone are not sufficient to explain an accumulation of citrate. Other path-
106
1. MALONATE
Table 1-19
Distribution of Radioactivity Following 3-Hr Incubation of Potato Tuber
Slices with Glucose-u-C^* "■
Component
Radioactivity (cpm)
Control
Malonate
% Change
Sucrose
344,000
306,000
- 11
Glucose
31,900
26,800
- 16
Fructose
5,950
5,830
- 2
Oligosaccharides
8,050
6,000
- 25
Weak acids
Citrate
4,350
1,280
- 71
Isocitrate
235
757
+222
Succinate
925
6,300
+582
Fumarate
190
125
- 34
Malate
8,900
1,740
- 80
Glycolate
2,750
1,830
- 33
Total
67,300
41,500
- 38
Acidic amino acids
56,100
16,700
- 71
Neutral amino acids
32,100
56,200
+ 75
Basic amino acids
950
750
- 21
Phosphorylated compounds
28,000
10,000
- 64
Lipids
111
367
+230
Respiratory COj
22,600
17,300
- 23
" Fresh slices of potato tubers incubated at 28° and pH 5 for 3 hrs with uniformly
labeled glucose, in the absence of and presence of 50 xnM malonate. (From Romberger
and Norton, 1961.)
ways for the formation of citrate are known, e.g., the citrase reaction from
acetate and oxalacetate, or through isocitrate by the isocitrase reaction
from succinate and glyoxylate. However, the free energy changes for these
reactions are such that citrate would not accumulate in significant amounts
even though the substrates for its formation accumulated. Also the results in
Table 1-22 show that the effect is not specific for malonate, but is seen with
succinate, malate, fumarate, and other organic anions. The marked effect
of glutarate, which is a poor inhibitor of succinate oxidase, suggests that
the citrate accumulation may not be related to the block of this enzyme.
It may be noted that part of the augmented citrate excretion may be attri-
SUBSTRATES OTHER THAN SUCCINATE
107
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1. MALONATE
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SUBSTRATES OTHER THAN SUCCINATE
109
Table 1-22
Urinary Excretion of Citrate Following Administration of the Sodium Salts
OF Various Weak Acids
Substance
Urinary citrate " after dose '' of :
4.35
26
26
39
None
NaCl
NaHCOg
Malonate
Succinate
Pyruvate
a-Ketoglutarate
Fumarate
Malate
Oxalacetate
Glutarate
Adipate
Maleate
Acetate
Citrate
Aconitate
0.47
16
0.50
—
0.90
44
14.76
583
14.80
418
336
17.10
275
18.50
287
—
216
—
1150
—
200
61.30
210
1.53
—
15.2
—
413
2.4
9-25
54.2
29.5
10.2
12.3
25.8
13.1
6.7
44.3
3.7
11.4
26.3
10-17
72.4
39.4
42.3
36.6
20.1
45.7
16.2
65.8
17.6
45.0
34.5
52.5
42.0
" The second column shows the 24-hr urinary citrate (mg/kg) in dogs. From Orten
and Smith, (1937.)
The third column shows the urinary concentration of citrate (mg%) in rats.
(From Simola and Kosunen, 1938.)
The fourth and fifth columns show the 24-hr urinary citrate (mg) in rats. (From
Krusius, 1940.)
* Dose in miUimoles per kilogram.
buted to alkalosis. Crawford (1963) confirmed that the injection of 10 mil-
limoles/kg of malonate in rats causes a marked rise in urinary citrate
(1.7 -> 47 /^moles/kg/hr), but found that succinate, malate, and sodium
bicarbonate also have this effect. Serum citrate simultaneously rises very
moderately. It was concluded that all the effects are due to an alkalosis
induced by the administration of sodium. However, this cannot account for
all of the actions of malonate, nor could it be responsible for the citrate ac-
cumulation in cell suspensions and isolated preparations.
Another possibility is that the citrate arises from the substances that
are administered. This was favored by Orten and Smith (1937), but it was
110 1. MALONATE
difficult then with incomplete knowledge of the cycle and related pathways
to understand how such conversions could take place; it still is. That is,
if there is no significant impairment of the utilization of citrate by mal-
onate, it is difficult to conceive of a pathway for the formation of citrate from
malonate that would be so rapid as to lead to a large rise in the citrate level.
A possibility that seems not to have been considered is that the substance
determined as citrate may not have been citrate but a related tricarboxylic
acid or some other compound giving a positive test. Although Hallman
(1940) examined the specificity of the determination, there are many
substances that have not been tested. For example, it is easy to formulate
reactions in which malonyl-CoA could react with various carbonyl sub-
stances, such as glyoxylate or pyruvate, to form tricarboxylate anions which
might be oxidized to pentabromacetone in the citrate test and be mistaken
for citrate. Certain dicarboxylates. such as itaconate, also are determined in
this test. Such substances may not be readily metabolized and hence would
accumulate much more readily than citrate. Although this posibility may
seem far-fetched, it would be well to make certain that it actually is citrate
that is accumulating during the action of malonate. It would be necessary
to convert only a small fraction of the administered malonate to such a
compound, since a dose of 26 millimoles/kg (see column 3 of Table 1-22)
would theoretically give rise to almost 1 g of a substance with a molecular
weight near that of citrate, whereas actually only around one-twentieth
of this was determined as citrate. Of course, such estimations depend on the
degree of sensitivity of the test to the compound. If there is any validity in
this suggestion, it may be that depressions of citrate levels may occur in
those preparations or tissues where such reactions of malonate or its meta-
bolic products do not occur, i.e., where the response to malonate is the one
expected on the basis of its inhibition of the functioning of the cycle.
Accumulation of a-Ketoglutarate
Very large increases in urinary a-ketoglutarate following the administra-
tion of malonate to rabbits and rats were reported by Krebs et al. (1938)
(Table 1-17) and this has been confirmed by El Hawary (1955), who found
a 3.7-fold increase in serum a-ketogluterate 30 min after the intraperitoneal
injection of 20 millimoles/kg malonate. As in the case of citrate, Krusius
(1940) found that a-ketoglutarate excretion is increased not only by mal-
onate but by many organic anions: malonate (46.4), maleate (42.0), malate
(17.7), succinate (17.7), /5-hydroxybutyrate (15.7), acetate (14.3), pyruvate
(12.1), fumarate (9.3), and sodium bicarbonate (0.6-7.8) (the figures give
urinary excretion in milligrams/day). He concluded that essentially all the
substances that increase citrate also raise the a-ketoglutarate excretion.
However, glutarate is a notable exception, for it potently augments citrate
formation but has no effect on a-ketoglutarate excretion. This would seem
SUBSTRATES OTHER THAN SUCCINATE 111
to disprove the theory that glutarate is active because it undergoes /?-oxi-
dative decarboxylation to malonate. Little has been done with isolated
preparations, but in three cases accumulation of a-ketogkitarate has been
observed in the presence of malonate: in locust sarcosomes from malate
(Rees, 1954), in suspensions of ascites cells from fumarate (Kvamme,
1958 c), and in ascites cells and rat heart mitochondria from glutamate and
malate (Borst, 1962), in all instances the malonate concentration being
rather high (20-50 mM).
The mechanism of such accumulation is poorly understood. We have seen
that in some tissues the a-ketoglutarate oxidase can be inhibited by malo-
nate, especially at concentrations above 10 mM, so that the results can be
partially explained in this way (at least for the ascites cells and locust
particulate preparations). The moderate increases in a-ketoglutarate excre-
tion brought about by the cycle intermediates and substrates might well be
due to a greater rate of formation with unchanged utilization rate. However,
the possibility of a formation of c-ketoglutarate from glutamate cannot be
ignored. It will be recalled that in potato slices the succinate formed in the
presence of malonate comes partly from such a source (Romberger and
Norton, 1961). Permeability effects causing a leakage of a-ketoglutarate
and other anions from the tissues must also be considered. El Hawary (1955)
found that several inhibitors (arsenite, maleate, iodoacetate, alloxan, and
fluoride) increase the serum a-ketoglutarate levels in rats, and simulta-
neously raise pyruvate levels. It may well be that any severe metabolic
disturbance causes a release of cycle substrates from the tissues and an
increased excretion, as well as secondary changes such as hyperglycemia.
Effects on the Levels of Other Cycle Substrates
The tissue concentrations of all the cycle substrates are probably altered
by malonate and it is sufficient here to mention the results with potato tuber
slices (Table 1-19) and tobacco leaves (Table 1-20). The reduction in the
incorporation of C^** from glucose into malate in the former and the marked
falls in malate level in the latter would be anticipated from a block of suc-
cinate oxidation. Fumarate and oxalacetate levels probably are changed
similarly. In this connection, one wishes that more information were avail-
able on the factors that control the tissue pools of cycle intermediates, since
it is evident that these substances do not occur only in the mitochondria
in kinetic equilibria depending on the relative rates of the cycle reactions,
but must also be present in cellular compartments. The transfer of the
substances between these compartments and the active cycle regions must
depend on processes that could be affected by inhibitors. Such compart-
ments are well known in plant cells but it is probable that similar situations
are applicable to animal cells.
112 1. MALONATE
Sequential Inhibition with Malonate and Fluoroacetate on Citrate Levels
Before leaving the subject of accumulation of cycle intermediates, a
few words must be said on the effects of malonate on the increases in citrate
brought about by fluoroacetate. This was discussed in Volume I (page 502)
and the results obtained by Potter (1951) presented (Fig. 1-10-5). The prin-
ciple of the experiments is simply that fluoroacetate blocks the utilization
of citrate so that the tissue citrate levels rise at different rates and to dif-
ferent degrees. If malonate is administered to the animals prior to the
fluoroacetate, the accumulation of citrate may be modified. These studies
thus provide information on the effects of malonate on the rates of forma-
tion of citrate and supplement the results discussed above.
The tissues differ greatly in their response to malonate in the presence of
fluoroacetate. In thymus, for example, malonate blocks the formation of
citrate completely and no citrate accumulation at all occurs. The kidney
behaves similarly but some citrate begins to accumulate an hour after
the fluoroacetate is injected. In spleen and brain the inhibition of citrate
formation is around 50%. Heart responds quite differently from the other
tissues. Here malonate actually increases the citrate formation somewhat.
Potter believed these results to indicate that in heart there are pathways
other than the cycle for the synthesis of oxalacetate, and suggested the con-
version of ketone bodies (malonate induces ketonemia) to citrate by a
pathway involving oxalacetate, the ketone bodies arising from fatty acids.
Liver was not studied but if similar reactions occur in this tissue, they
could,. at least in part, account for the increases in serum citrate and urinary
excretion of citrate. In any event, these experiments illustrate very well
the inherent differences between tissues with respect to their metabolic
pathways. The stimulation of citrate formation in the heart in the presence
of fluoroacetate was confirmed by Fawaz and Fawaz (1954), whereas in the
kidney only a depression was observed. The use of fluoroacetate to block
the utilization of citrate is a useful technique by which to study the effects
of malonate uncomplicated by possible effects on the rate of disappearance
of citrate.
ANTAGONISM OF MALONATE INHIBITION
WITH FUMARATE
The overcoming of an inhibition by the addition of an intermediate nor-
mally arising distal to the site of the block is often excellent evidence
for the locus of action of the inhibitor and for the specificity of the inhi-
bition. For this reason, fumarate has frequently been used in malonate-
inhibited preparations and a reversal of the inhibition taken as proof for
the specific action of malonate on succinate oxidase. The first acceptable
ANTAGONISM WITH FUMARATE 113
data for fumarate reversal were reported by Quastel and Wheatley (1935),
who showed that the inhibition of acetoacetate utilization by rat liver in
the presence of malonate is mainly abolished when fumarate is added,
and they used this as evidence that malonate does not act directly on the en-
zyme involved in the breakdown of acetoacetate. The use of fumarate im-
mediately became popular and many studies since 1935 have included its
addition for the purpose of demonstrating that the observed effect of mal-
onate is indeed due to its block of succinate oxidation.
Most of the tests for fumarate reversal, it must be admitted, have not
been done properly and the results have been evaluated uncritically, so that
little of value has been demonstrated. There are several points that must
be considered in the planning and interpretation of such experiments.
(a) An increase in the oxygen uptake upon addition of fumarate to a
malonate-inhibited preparation is not, by itself, very meaningful. Let us take
a typical experiment similar to many reported in the literature. The nor-
mal Qq of a tissue preparation is 10 and this is decreased to 4.5 by mal-
onate. When malonate and fumarate are added together, the Qq is 8.7.
It has been generally stated in such cases that fumarate is capable of
antagonizing the malonate inhibition. Actually, an increase in the respira-
tion could have been brought about by the addition of any substrate that
is oxidized by the preparation, including substrates completely unrelated
to the cycle. One can say that fumarate has overcome the malonate inhibi-
tion but the results are of no particular significance with respect to malonate
specificity or the site of the inhibition. What one has shown is that fumarate
can be oxidized in the presence of malonate and this need not imply that
the operation of the cycle has been even partially restored. If fumarate
had been added to the uninhibited preparation and a Qq of 14.2 found, the
following might have been concluded: malonate inhibits the endogenous
respiration 55% and the respiration in the presence of fumarate either 13%
(with respect to the endogenous control) or 39% (with respect to the rate
with fumarate alone present), in both cases the inhibition being less than that
for the endogenous respiration. Actually, it has been shown that the oxi-
dation of fumarate is unaffected by malonate, since the same absolute
rise in the Qq^ is obtained from fumarate in the uninhibited and inhibited
preparations. If the oxygen uptake resulting from the addition of fumarate
results mainly from the oxidation of malate, the results would have little
bearing on the mechanism or selectivity of malonate inhibition.
(6) The addition of fumarate would not in any case restore the complete
cycle, since oxidation of succinate would still be depressed and less energy
would be available from this step. It is possible in some instances that the
energy from succinate oxidation, rather than the over-all energy production
from all oxidations, is important, and this would not be restored by fu-
marate.
114 1. MALONATE
(c) The accumulation of succinate due to malonate inhibition would, of
course, not be reversed by fumarate; instead, it is usually increased. If some
response to malonate is dependent on the rise in succinate concentration
(e.g., a direct effect of succinate on other enzymes or some cell function, or
the increased formation of some substances derived from the succinate),
this would not be antagonized by fumarate.
(d) The response to fumarate will often depend on what is being measured.
One example will be used here to illustrate this and others will be men-
tioned later. Malonate inhibits the oxidation of trilaurin and octanoate
in liver and kidney slices, and also reduces the amount of C^^Og formed from
labeled substrates (Geyer et al., 1950 a). It was found that fumarate is very
ineffective in counteracting the inhibition of C^^O, production, and this
might be attributed to a direct effect of malonate on fatty acid oxidations.
However, the results can be explained on the basis of an inhibition of suc-
cinate oxidase. Much of the C^* taken into the cycle from acetyl-CoA
would accumulate in succinate and fumarate would have no effect on this.
For the full release of CO2, the operation of the entire cycle is required, and
thus fumarate would increase the C^^Og formation only moderately in the
presence of malonate. If the oxygen uptake had been determined, fumarate
could well have shown a complete reversal of the malonate inhibition.
(e) Absence of a reversal by fumarate, or the failure to achieve a complete
reversal, can be due to a variety of causes. In an experiment, such as the one
shown in the following tabulation, done on Aplysia muscle slices:
Additions Qq^
Endogenous
0.33
Succinate
1.80
Malonate
0.33
Succinate -f malonate
0.96
Succinate + malonate + fumarate
0.90
although it was stated that fumarate was unable to relieve the malonate
inhibition (Ghiretti et al., 1959), it may simply be that fumarate would not
have been oxidized if added to the uninhibited tissue. Certainly the direct
inhibition of succinate oxidase would not be expected to be overcome by
fumarate, but in any case a control with fumarate alone must be run to
give any significance to the results. It is even possible for fumarate to in-
crease the inhibition produced by malonate. This was observed for the respi-
ration of Helix hepatopancreas (see accompanying tabulation) (Rees, 1953).
Here the endogenous respiration apparently is little dependent on the cycle,
whereas upon the addition of fumarate, through the formation of oxalace-
ANTAGONISM WITH FUMARATE 115
tate and its partial decarboxylation to pyruvate, the cycle presumably
becomes activated. The high inhibition of fumarate oxidation by malonate
cannot, of course, be attributed solely to an inhibition of succinate oxidase,
Additions Og uptake % Inhibition
Endogenous
30
Malonate
25
Fumarate
167
Malonate + fumarate
47
16.7
71.9
and an effect on oxalacetate decarboxylase or some other enzyme at the
relatively high malonate concentration (33 mM) must be assumed. It is
desired to point out that the effects with fumarate are unrelated to the
inhibition of the endogenous respiration observed with malonate and
throw no light on the mechanism of the inhibition.
Let us now turn to some experiments in which the addition of fumarate
provides results indicative of the mechanism of malonate inhibition. The
most significant studies usually have involved the determination of the
utilization or formation of a particular substance. In the original work
of Quastel and Wheatley (1935) mentioned above, the disappearance of
acetoacetate in rat liver slices was determined, and fumarate was found to
increase its utilization in the presence of malonate, whereas in the absence
of malonate it had essentially no effect. Krebs and Eggleston (1940) likewise
showed that fumarate would increase the utilization of pyruvate in pigeon
muscle in the presence of malonate, at high malonate concentrations the
amount of pyruvate utilized being equivalent to the fumarate added. The
results obtained by Stare et at. (1941) with pigeon muscle show definitely
that the block of pyruvate utilization produced by malonate is effectively
overcome by fumarate (see accompanying tabulation). Results such as
Additions PjTuvate utilized
Endogenous 16 . 3
Malonate 7 . 2
Fumarate 21.4
Malonate + fumarate 23.3
these demonstrate that fumarate can counteract the effect of malonate on
a specific process and, from the nature of the experiments, it is likely that
the cycle block is being overcome in a sense, that is, that oxalacetate is
being made available for condensation with acetyl-CoA. They also provide
116 1. MALONATE
some information on the site and specificity of the malonate inhibition.
Quite different results on acetate utilization by yeast were obtained by
Stoppani et at. (1958 b), who found that fumarate is completely unable to
reverse the inhibition by malonate (see accompanying tabulation). These
Additions Acetate utilized
Acetate 30
Acetate + malonate 4.4
Acetate + fumarate 27
Acetate + malonate + fumarate 3.8
data would nuply that the inhibition of acetate utilization by malonate is
not mediated through a block of succinate oxidation but by another jnecha-
nism. It is difficult to explain this by a failure of fumarate to penetrate
into the cells since malonate seems to enter readily. It was suggested that
malonate might interfere with acetate activation by depleting the system
of coenzyme A, due to the formation of relatively stable malonyl-CoA.
Whatever the mechanism, these results point to an action of malonate
other than on succinate oxidase.
An interesting illustration of the useful information that may be ob-
tained from the use of fumarate is given in the inhibition of urea formation
by malonate. The formation of both arginine and urea from citrulline and
glutamate in liver homogenates is potently inhibited by malonate (Cohen
and Hayano, 1946; Krebs and Eggleston, 1948). Fumarate is able to counter-
act this block completely. These results were difficult to understand ini-
tially, but it is now known that transamination must occur between gluta-
mate and oxalacetate to form aspartate, which reacts with the citrulline
to form arginosuccinate, from which arginine and urea are derived. The
effect of malonate is to reduce the supply of oxalacetate for transamination
and it is clear why fumarate will abolish this inhibition (Ratner, 1955).
Krebs and Eggleston (1948) observed the formation of 3-5 molecules of
urea for each molecule of fumarate added. This may be explained by the
fact that the formation of arginine from arginosuccinate involves the release
of fumarate, which can again go to oxalacetate.
The demonstration that fumarate will counteract the inhibitory action
of malonate on some tissue function is indicative of a primary block of the
succinate oxidase, but even complete reversal does not prove a specific ac-
tion of malonate. One example would be the inhibition of Br~ uptake in
barley roots by malonate (Machlis, 1944). Malonate (10 mM) inhibits the
uptake 57% but if fumarate is present the uptake is 35% above the control.
On the surface this would imply an effective reversal of the malonate inhi-
SPECIFICITY OF MALONATE INHIBITION IN THE CYCLE 117
bition, but actually fumarate alone increases the uptake to 71% above the
control. Thus malonate inhibits a significant amount even in the presence
of fumarate, pointing to an action other than on succinate oxidation.
Another example would be the malonate inhibition of cell division oiArhacia
eggs (Barnett, 1953). Cleavage is inhibited almost completely by 60 milf
malonate and an equimolar concentration of fumarate abolishes this. It is
likely that fumarate overcomes the cycle block but one cannot conclude
that the action of malonate is specific on succinate oxidase. Malonate, can
also inhibit to some extent steps in the utilization of fumarate, but due to
the high concentration of fumarate enough oxalacetate is formed to allow
cleavage. Indeed, succinate atGOmJf also abolishes the inhibition, indicating
that as a result of the competitive nature of the inhibition enough succinate
has broken through the block to restore cleavage. It must be remembered
that a complete reversal of a metabolic block is not always necessary for a
cell function to proceed normally.
SPECIFICITY OF MALONATE INHIBITION IN THE CYCLE
At this point we may summarize some of the conclusions with respect to
the specificity of action of malonate on the succinate dehydrogenase. Possi-
ble effects of malonate outside the cycle will be discussed in a later section.
We have seen that malonate can inhibit enzymes other than succinate dehy-
drogenase rather potently (Table 1-12), that the oxidations of certain cycle
substrates are suppressed more than predicted on the basis of a selective
action on succinate oxidation (Table 1-14). that the accumulation patterns
of cycle intermediates are sometimes distorted by malonate in ways implying
inhibition at more than one site, and that fumarate is seldom able to reverse
the actions of malonate completely. Of particular significance are the clear
demonstrations of the inhibition of reactions related to the entry of acetyl-
CoA into the cycle, particularly those of Pardee and Potter (1949) pointing
to an inhibition of the condensation reaction, those of Stoppani et al.
(1958 b) reporting a marked inhibition of acetate utilization apparently
unrelated to an inhibition of succinate oxidase, and our own results on rat
heart mitochondria where malonate inhibits p^Tuvate oxidation in the
presence of malate and with the a-ketoglutarate oxidase completely blocked.
Another susceptible site is a-ketoglutarate oxidation, especially in view
of the clear proof by Price (1953) that specific inhibition can not even be
obtained in the simple system,
a-Ketoglutarate -^ succinate ->■ fumarate
There is thus a large amount of evidence that malonate can at certain
concentrations in various conditions inhibit other reactions than succinate
oxidation.
118 1. MALONATE
It would be convenient if one could specify a malonate concentration,
or range of concentrations, which would most likely be specific, but there
are too many factors involved to do this with any confidence. Some of the
factors may be listed: (a) the species, tissue, or preparation used, (b) the
enzymes or metabolic pathways involved in what is measured, (c) the
conditions of the experiment, e.g., the pH or the Mg++ concentration, (d)
the degree to which succinate can accumulate and antagonize the inhibi-
tion, (e) the effective concentration of malonate within cells, and (f) the
possibility of nonenzyme effects on cell membranes or other structures.
Specificity for an inhibitor is not a constant to which can be given a value
for all cases, but a characteristic that must be evaluated for each experi-
ment. Aside from the direct actions of malonate on enzymes, there are the
problems of metal cation depletion, the possible effects of Na+ or K+ added
with the malonate, and the inactivation of the coenzyme A in some pre-
parations through the formation of malonyl-CoA.
Bearing in mind these difficulties, a few general remarks may be made.
It is probable that malonate usually does not inhibit any cycle enzyme more
strongly than succinate dehydrogenase, so that the major effect will be re-
lated to the inhibition at this site, but it will be recalled that in certain
species the succinate dehydrogenase is not very susceptible to malonate. It
is impossible to achieve a nearly complete block of succinate oxidation
without affecting other cycle reactions; if one wishes a specific effect, one
must be satisfied with a moderate inhibition of succinate oxidation. If a
single malonate concentration for general usefulness had to be chosen, 5 mM
might be taken provisionally. Although in some cases rather incomplete inhi-
bition will be obtained, this concentration will probably not inhibit other
cycle reactions significantly. This applies to noncellular preparations where
penetration is not a factor but otherwise higher external concentrations
may have to be used. In any case, good evidence for a specific action should
be obtained under the conditions of the investigation, and reliance should
not be based on generalities.
EFFECTS OF MALONATE
ON OXIDATIVE PHOSPHORYLATION
An uncoupling action on oxidative phosphorylation has been claimed for
malonate several times and it is important to determine if this is actually
so. This is quite difficult because pertinent or reliable data are generally
lacking. There are four important ways in which malonate could alter the
P : 0 ratio: (1) a direct effect on the coupling between oxidation and phos-
phorylation, (2) an alteration of the pattern of substrate oxidation, since
different substrates may have different P : 0 ratios, (3) a differential inhi-
bition of electron transport pathways for a single substrate but with differ-
EFFECTS ON OXIDATIVE PHOSPHORYLATION 119
ent P : 0 ratios, and (4) an effect on the hydrolysis of ATP or other high
energy substances. Only the first mechanism should be considered as true
uncoupling, although it is often difficult to determine the exact mechanism.
Included in the first mechanism would be the chelation of malonate with
Mg++ or Mn++, since these cations are beheved to be cofactors in phospho-
rylation.
Claims for an uncoupling action will be discussed first. Lehninger (1951)
stated that malonate uncouples oxidative phosphorylation associated with
the oxidation of /5-hydroxybutyrate by rat liver mitochondria but no data
were given. In a previous report (Lehninger, 1949) 7.5 mM malonate was
shown to inhibit oxygen uptake 48%, the formation of acetoacetate from
/?-hydroxybutyrate 64.7%, and phosphorylation 37.3% (as determined by
the incorporation of P''- into the ester fraction). Since phosphorylation is
inhibited less than oxidation, no uncoupling is evident, and indeed the P:0
ratio should increase. Berger and Harman (1954) claimed that malonate
inhibits phosphorylation associated with the one-step oxidation of a-
ketoglutarate and completely suppresses phosphorylation during the oxida-
tion of L-glutamate by muscle mitochondria. However, the malonate con-
centration is not given and the absence of data prevents evaluation of the
results. An inhibition of phosphorylation does not necessarily mean an
uncoupling action. Malonate at 30 milf drops the P:0 ratio from 1.6 to 0.5
in the oxidation of choline by rat liver mitochondria (Rothschild et al.,
1954). Although no control with malonate alone was reported, it would
appear that this is the most valid instance of uncoupling by malonate.
The malonate concentration was high and a reduction of Mg++ (total con-
centration was 5.7 mM) must be considered. Finally, the phosphorylation
associated with succinate oxidation in lupine mitochondria was shown to
be strongly depressed by 10 mM malonate (Conn and Young, 1957), but
it may be observed that the oxygen uptake was inhibited even more (Ta-
ble 1-23), so that no uncoupling occurred.
All of the reports in which P:0 ratios were calculated are summarized
in Table 1-23 and in all cases, except for E. coli, Carcmvs maenas, and the
oxidation of choline, it is seen that the P:0 ratio is actually increased by
malonate. However, most of these only illustrate mechanism (2) above,
because in the oxidation of a-ketoglutarate a rise in the P:0 ratio would
be expected upon blocking succinate oxidase due to the fact that P:0 for
succinate oxidation is 2, whereas for the one-step oxidation of a-ketoglu-
tarate to succinate usually it is experimentally between 3 and 4. All one can
say from such data is that there is no evidence for an uncoupling action by
malonate. Copenhaver and Lardy (1952) used 3-20 mM malonate in all
their media in the study of the phosphorylation associated with a-keto-
glutarate oxidation and obtained high P:0 ratios, again providing evidence
against any uncoupling activity. It was shown by Slater and Holton in
120
1. MALONATE
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EFFECTS ON OXIDATIVE PHOSPHORYLATION 121
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>
^
cs
5^
x>
>i)
o^o^^d PM pm § 0^ Ph p5 m
122 1. MALONATE
rat heart (1954) that malonate increases the P:0 ratio with a-ketoglutarate
as the substrate as the malonate concentration is increased up to 20 mM;
at 40 mM the P:0 ratio decreases somewhat, so that at this high concen-
tration a small degree of uncoupling may occur. Azzone and Carafoli (1960)
found the same in pigeon muscle. Evidence against uncoupling by malonate
intracellularly is provided by the study on ascites carcinoma cells by
Greaser and Scholefield (1960). Comparison of the changes brought about
by malonate (20 mM) in the respiration and the sum of the concentrations
of ADP and ATP led to the estimation of a 15% increase in the P:0 ratio,
whereas the classic uncoupler, 2,4-dinitrophenol, depressed the P:0 ratio
70%. Malonate may actually stimulate the esterification of phosphate. In
addition to the two examples in Table 1-23, Jackson et al. (1962) reported
an 8% elevation in phosphate uptake by 0.1 mM malonate in barley root
mitochondria oxidizing succinate, and Rosa and Zalik (1963) found such a
stimulation in pea seedling mitochondria, which is maximal around 0.01
mM malonate, the oxygen uptake not being altered. Above this concentra-
tion, both phosphorylation and oxidation are depressed in a parallel fashion;
respiration is hence always depressed somewhat more than is phosphate
esterification, and no uncoupling is seen at any malonate concentration.
The transfer of phosphate from phosphorylated coenzyme Q to ADP to
form ATP in mitochondrial preparations is not altered by even 10 milf
malonate (Gruber et al., 1963), whereas dinitrophenol inhibits this readily.
It would therefore appear to be legitimate to conclude that malonate
does not exhibit uncoupling activity except possibly at high concentrations
(above 30 mM). The evidence for uncoupling at concentrations commonly
used is considered to be inadequate and outweighed by the mass of indirect
evidence that P:0 ratios are not reduced in the oxidations of citrate, a-
ketoglutarate, and succinate.
EFFECTS OF MALONATE ON GLUCOSE METABOLISM
The actions of malonate on carbohydrate, lipid, amino acid, porphyrin,
and other types of metabolism will now be considered, after which it will be
possible to evaluate the specificity of malonate more broadly. The effects
on glucose metabolism are important but difficult to analyze. In the first
place, the interrelationships between the cycle and the glycolytic pathways
are complex and secondary effects on glucose utilization must be expected.
In the second place, much of the work on the alteration of glucose utiliza-
tion by malonate has not been adequate for the determination of mecha-
nisms nor do the results even provide useful information in many cases, and
for this reason only certain reports will be discussed. There are three basic
ways by which malonate, might alter glucose oxidation. (1) Malonate will
usually cause a depression of the oxygen uptake related to glucose metabo-
EFFECTS OF MALONATE ON GLUCOSE METABOLISM 123
lism because in the total oxidation of glucose to CO, and water, five-sixths
of the oxygen is taken up in cycle reactions. This effect would be due to a
block of the cycle and would be roughly equivalent to the inhibition of
of pyruvate oxidation via the cycle. (2) Malonate may cause an increased
uptake and utilization of glucose as a result of inhibition of oxidative reac-
tions in the cycle, the mechanism being essentially that of the Pasteur reac-
tion. (3) Malonate may have direct actions on the cellular uptake of glucose
or on the glycolytic pathways. Evidence for each of these mechanisms will
be presented, and then a more detailed analysis of the possible shifts in
metabolic patterns brought about by malonate will be given.
Effects on Oxygen Uptake Associated with Glucose Oxidation
There are many reports on the effects of malonate on respiration with
glucose as the substrate, but in few have the data provided a correction for
an effect on the endogenous respiration. Indeed, an endogenous correction
to determine the oxygen uptake associated only with glucose oxidation
is particularly unreliable, even when the data are available. This is because
of the well-known effect of glucose in suppressing endogenous respiration and
mitochondrial oxidations (Crabtree effect). Thus the nonglucose respiration
may be significantly changed when glucose is added and, perhaps, completely
suppressed in some cases.
Let us assume that the oxygen uptake measured derives only from glucose
and that malonate acts only on the cycle. What inhibitions could one theo-
retically expect? The inhibition will depend, other than on the degree of
cycle block, on the final products of glucose oxidation before and after the
addition of malonate. The following equations give the molar oxygen up-
takes for the oxidation of glucose to varying degrees of completeness:
To CO2 and water
CeHi^Oe + 6 O2 ^ 6 CO, + 6 H,0
To pyruvate
CgHi^Oe + O2 -> 2 CH3COCOOH + 2 H2O
To acetate
CeHiaOe + 2 0, -> 2 CH3COOH -f- 2 CO, + 2 H^O
To succinate
CfiHi^Oe + 5/2 O2 -> HOOCCH2CH2COOH + 2 CO2 + 3 H2O
CsHiaOg + 2 HOOCCH2COCOOH + 4 0, -> 2 HOOCCH2CH2COOH + 6 CO2 + 4 H^O
To lactate
CsHi^Oe + 2 H -). 2 CH3CHOHCOOH
The second equation for the formation of succinate might express the situa-
tion in which there is a source of oxalacetate other than from pyruvate
124 1. MALONATE
carboxylation, whereas the first equation assumes no external oxalacetate
source. If the inhibition on succinate oxidase is complete and glucose is
oxidized completely in the control, we may calculate the inhibitions for the
situations where in the presence of malonate the glucose is transformed into
the various oxidation products: to succinate with external oxalacetate
source (33%), to succinate in a closed system (58%), to acetate (67%),
to pyruvate (83%), and to lactate (100%). Of course, experimentally it is
probably impossible to achieve a complete cycle block with malonate, so
that the inhibitions will usually be less than the theoretical. These calcula-
tions also assume that the rate of glucose disappearance is not altered by
the inhibition. The degree of inhibition, therefore, even under these simple
conditions, will vary a good deal, depending on the terminal product of
glucose metabolism and thus on the enzymes and pathways occurring in
the cells under consideration. Actually, it is likely that various substances
will accumulate during glucose metabolism, and not only those given above
but others into which these substances can be metabolized.
A strong inhibition of the oxygen uptake related to glucose metabolism
probably indicates the participation of the cycle in the oxidation, at least
when the malonate concentration is not unreasonably high. Thus, in the
instances shown in the tabulation and selected at random of inhibition of
Preparation
Malonate
(mif)
%
Inhibitior
1 Reference
Sarcina lutea
10
50
Dawes and Holms (1958)
Yeast
12.5
73
Krebs et al. (1952)
Bull sperm
10
57
Lardy and Phillips (1943 a)
Malarial parasitized
RBC
20
69
Speck et al. (1946)
Brain mince
17
100
Huszak (1940)
/iL+-Stimulated brain slices
10
62
Kimura and Niwa (1953)
glucose respiration, one would conclude that the role of the cycle is impor-
tant, but no information on the exact mechanism or on the fate of glucose
in the presence of malonate may be obtained. However, in the cases of brain
mince and yeast, fumarate is unable to overcome the inhibition, so that
some doubt is introduced even into this interpretation. The inhibition of
the glucose oxygen uptake in parasitized erythrocytes is very similar to
that for the oxygen uptake associated with pyruvate oxidation, as expected
if the cycle is the terminal pathway for glucose metabolism. On the other
hand, there have been many reports that malonate, even at high concentra-
tions, does not inhibit the glucose respiration at all, and in these instances
little can be concluded because of the possibility that malonate does not
penetrate and that an adaptation of the glucose utilization occurs. The
EFFECTS OF MALONATE ON GLUCOSE METABOLISM 125
endogenous respiration of Ehrlich ascites cells is inhibited around 35% by
30 vaM malonate, but when glucose is present there is very little effect of
malonate (Seelich and Letnansky, 1960). Put in another way, in the pres-
ence of malonate the addition of glucose increases the 0, uptake somewhat
instead of depressing it as it does in uninhibited cells. Another possible
reason for a failure of malonate to depress glucose respiration is the oc-
currence of oxidative pathways unassociated with the cycle. Caldariomyces
fumago has no hexokinase and the usual Embden-Meyerhof glycolytic
pathw^ay is absent; the oxidation of glucose occurs more directly through
glucose and gluconate oxidases (Ramachandran and Gottlieb, 1963). The
respiration in this organism is not affected by malonate at 10 mM.
Effects on Glucose Utilization
A depression of cycle oxidations by malonate would be expected to cause
an increased utilization of glucose, as do hypoxic conditions, in cells capable
of exhibiting a Pasteur reaction. This is a manifestation of one interrela-
tionship between the cycle and the glycolytic pathway, and is partially
mediated through changes in the concentrations of inorganic phosphate
and phosphate acceptors, such as ADP. For such a response to occur, the
utilization of glucose must have been initially limited by the coupled
phosphorylation at the 3-phosphoglyceraldehyde oxidation step. Another
mechanism for the influence of cycle activity on glucose utilization involves
the membrane transport of glucose and its phosphorylation. The inward
transport of glucose under certain conditions may limit glucose utilization,
and it has been shown that anoxia accelerates this transport in rat heart
(Morgan et al., 1961 a, b). It is quite possible that malonate by its depres-
sion of cycle activity, can alter these transport processes. A third mechanism
might involve the rate of oxidation of NADH. The addition of pyruvate to
ascites cells stimulates the formation of C^^02 from labeled glucose (Wenner
and Paigen, 1961) Initially the rate of pyruvate oxidation is limited by the
rate of NADH oxidation and the exogenous pyruvate acts as an electron
acceptor, 1 mole of lactate appearing for each mole of pyruvate oxidized
through the cycle. The accumulation of pyruvate as a result of the cycle
block by malonate could initiate this dismutation so that more glucose
would be utilized than otherwise. Indeed, lactate formation is often increased
during malonate inhibition. In all these ways, and perhaps others, a cycle
block might affect glucose uptake and utilization.
A stimulation of glycolysis by malonate was first observed by Kutscher
in Heidelberg (Kutscher and Sarreither, 1940; Kutscher and Hasenfuss,
1940). Malonate was injected into guinea pigs, the muscle removed later,
and the formation of lactate determined in a brei. In some cases, glucose,
succinate, or fumarate was also injected. Malonate accelerates lactate for-
mation and this is overcome by both succinate (see accompanying tabula-
126 1. MALONATE
tion) and fumarate. It was also shown that the rate of glycogen disappear-
ance in muscle brei is stimulated some 35% by malonate. These responses
were equated with the Pasteur reaction. Similar effects have been found in
Injection
Lactate formation
Control
2.95
Malonate
9.2
Glucose
5.1
Glucose + malonate
9.4
Succinate + malonate
-1.0
Glucose + succinate + malonate
-0.1
other tissues more recently. The rates of glucose utilization and of lactate
formation in brain slices are stimulated 13% and 44%, respectively, by
10 mM malonate (Takagaki et at., 1958), an effect much like that produced
by azide. Simultaneously less glucose is oxidized, the increased utilization
being diverted to lactate. Ehrlich ascites tumor cells exhibit a more rapid
rate of glycolysis when the oxygen tension is reduced and a similar response
is seen with malonate (Kvamme, 1957, 1958 d). When fumarate is added
to the malonate-blocked cells, glucose utilization and lactate formation are
suppressed. Strictly speaking, it is fumarate that gives rise to a Pasteur
reaction in the presence of malonate, just as the addition of oxygen does
in preparations previously anaerobic. Kvamme obtained data which led him
to conclude that this effect is mediated through changes in the concentra-
tions of inorganic phosphate and phosphate esters. The relationships
between glycolysis and mitochondrial oxidations are particularly well seen
in the reconstructed systems of Aisenberg et al. (1957). A supernatant frac-
tion from rat liver forms lactate from glucose and the addition of a mito-
chondrial suspension suppresses this markedly. Malonate partially prevents
this suppression; that is, added to the complete system, lactate formation is
increased. This stimulation of glycolysis occurs despite the fact that mal-
onate inhibits the glycolytic rate 15.7% in the supernatant fraction. This is
a good illustration of how malonate can produce different effects on glucose
metabolism, depending on the conditions and the factors controlling gly-
colysis. Stimulation of aerobic glycolysis by malonate, and a variety of
other inhibitors, is particularly well seen in thymocytes; malonate ac-
celerates lactate formation sigmoidally from 3 mM to 100 mM (Araki
and Myers, 1963).
Malonate may have no effect on glucose uptake, or may inhibit it, in
other tissues. Chick chorioallantoic membrane infected with influenza
virus exhibits a 47.5% reduction in the endogenous respiration in the pre-
sence of 6 mM malonate, and the final virus titer drops to zero, but the
EFFECTS OF MALONATE ON GLUCOSE METABOLISM 127
uptake of glucose is unaffected (Ackermann, 1951). An example of a marked
inhibition was reported for guinea pig brain slices, glucose utilization being
reduced by 10 mM malonate in both normal and K+-stimulated slices (see
accompanying tabulation) (Tsukada and Takagaki, 1955). It may be ob-
KCl
(70 mM)
Malonate
(10 mM)
Glucose
utilization
(//mole/g)
Lactate
formation
(//mole/g)
O2 uptake
(//mole/g)
35.9
34.9
54.1
—
+
4.71
28.0
61.5
+
—
112
99.0
104
+
+
63.7
117
44.9
served that the amount of lactate formed per glucose consumed is increased
by malonate, but some of the lactate must be derived from other sources
than glucose. It is difficult to understand the discrepancy between these
results and those obtained later on the same tissue (Takagaki et al., 1958),
where a slight stimulation of glucose utilization by 10 mM malonate was
reported. There are so many different results obtained relative to the utili-
zation of glucose that it often appears each organism or tissue exhibits a
characteristic pattern of response. Malonate at 40 mM has relatively little
effect on the uptake of glucose by brain and kidney slices, depressing it
slightly in the former and perhaps accelerating it in the latter, but reduces
the C^^Og formed from uniformly labeled glucose 79% and 52%, respective-
ly, the O2 uptake being suppressed comparably (Cremer, 1962). Since much
less of the glucose goes to amino acids and proteins in the presence of mal-
onate (Cremer, 1964), it is likely that here there is an accumulation of
certain cycle intermediates, such as succinate, and of lactate. Succinate is
normally formed and released into the medium by Trypanosoma cruzi.
It is formed aerobically through both the glycolytic pathway and the cycle,
and by CO2 fixation; some must be metabolized through the cycle since
malonate elevates the succinate level even further (Bowman et al., 1963).
The uptake of glucose is increased almost 20% by malonate but there is no
change in C^^Oo formation, most of the excess glucose probably appearing
as succinate, acetate, and related anions.
Direct Effects on the Glycolytic Pathways
There is no evidence that any enzyme of the Embden-Meyerhof glycolytic
pathway is significantly inhibited by malonate at concentrations below 20
mM (Table 1-12), although some of the enzymes have never been investigat-
ed. Several of the enzymes in this pathway require Mg++, or a related cation.
128 1. MALONATE
and under certain conditions malonate could inhibit through the chelation
of these ions. A possibly sensitive enzyme is lactate dehydrogenase, which
functions in anaerobic glycolysis, although insufficient quantitative work
has been done, and it is likely that the enzyme from different sources would
be inhibited to different degrees by malonate. It is unfortunate that so
little work on the effects of malonate on anaerobic glycolysis has been done,
since studies under aerobic conditions are always complicated by the sec-
ondary reactions discussed above. In a number of cases, an inhibition of
aerobic glycolysis has been observed, i.e., a decreased formation of lactate
in air or 95% oxygen, and these inhibitions are usually small. In bull sperm
(Lardy and Phillips, 1943 b), 10 mM malonate inhibits 6.7%; in beef
thyroid homogenates (Weiss, 1951), 33 mM malonate inhibits 8%; and in
rat liver supernate (Aisenbergef oi., 1957), 25 mM malonate inhibits 15.7%.
Although rises in lactate can be explained on an indirect basis, a decrease
in the rate of lactate formation must usually be attributed to some inhibi-
tion along the glycolytic pathway, since it is not very likely that malonate
would shift the fate of pyruvate from lactate to the cycle. Greater inhibi-
tions have been observed: in mouse brain, 40 mM malonate inhibits lactate
formation 57% and in mouse liver mitochondria 100%, this being taken as
evidence of some direct effect on glycolysis (du Buy and Hesselbach, 1956).
This concentration is, of course, rather high and could have depleted the
Mg++ from either the 3-phosphoglyceraldehyde dehydrogenase or enolase
systems, or could have inhibited lactate dehydrogenase. Since the substrate
in these cases was 3-phosphoglyceraldehyde, an action earlier in the pathway
is impossible. There are also miscellaneous reports which might be inter-
preted as indicating an inhibition of glycolytic pathways, for example the
results of Greville (1936) on rat brain, in which 20 mM malonate inhibits
glucose oxidation around 50% and pyruvate oxidation only around 15%.
Only one instance of a direct test on anaerobic glycolysis has come to my
attention, that of Covin (1961), who found that 5 mM malonate inhibits
lactate formation in rat ventricle slices, but the rate of lactate formation in
this tissue is so slow, that Covin expressed some doubts as to the reliability
of the measurements. One must conclude from the incomplete data, that
there is some evidence for a minor inhibition of glycolysis by malonate,
especially at the higher concentrations.
The problem of the direct inhibition of glycolysis by malonate has been
studied particularly well by Eva and George Fawaz at the American Univer-
sity of Beirut. They had observed that 30 mM malonate almost completely
blocks the cycle, leads to the accumulation of succinate, and yet does not
depress the dog heart significantly (Fawaz et ol., 1958). However, 60 mM
malonate causes rapid reduction in cardiac frequency and contractile
failure, although no more succinate accumulates than with 30 mM. This de-
pression of cardiac function must be related to an action other than on the
EFFECTS OF MALONATE ON GLUCOSE METABOLISM 129
cycle. The formation of lactate from glucose in extracts of rat muscle is
inhibited over 90% by 60 mM malonate; simultaneously there is a decrease
in inorganic phosphate, creatine-P, and readily hydrolyzable phosphate,
accompanied by an increase in nonhydrolyzable phosphate (Fawaz and
Fawaz, 1962). It was concluded that the acid-resistant phosphate fraction
occurring in the presence of malonate must be made up of glycolytic inter-
mediates, and it was then shown by analyses of the incubated extracts at
various times that there is accumulation of 3-P-glycerate and glycerol- 1-P
particularly, with smaller contributions from phosphoenolpyruvate and
2-P-glycerate (see accompanying tabulation). Furthermore, the addition
Incubation
Accumulation of intermediates (mg ^
P/100 g
tissue)
time (min)
P-enolpyruvate
2-P-glycerate
3-P-glycerate
Glycerol- 1-P
6
6.12
1.84
19
.00
22
.30
30
8.60
2.45
29
.10
30.
,80
120
10.90
3.18
45
.60
33
,40
of these intermediates to a malonate-treated extract resulted in the ap-
pearance of a major fraction as 3-P-glycerate, whereas in control incuba-
tions they break down to inorganic phosphate and pyruvate or lactate.
This might imply a block at the pyruvate kinase and some reversal of the
glycolytic pathway which cannot proceed beyond 3-P-glycerate due to the
lack of ATP. The addition of pyruvate to an inhibited extract leads mainly
to the formation of lactate. Pyruvate kinase from rabbit muscle is inhibited
86% by 60 mM malonate and this correlates quite well with the results
seen in the extracts. Malonate at 30 mM only partly inhibits glycolysis,
causes less accumulation of the phosphorylated intermediates, and inhibits
pyruvate kinase 67%. The relative lack of affect of 30 mM malonate on
cardiac function is probably due to a combination of two factors: the intra-
cellular malonate concentration is undoubtedly less than 30 mM, and it
is likely that glycolysis in the heart can be depressed to a certain degree
before failure occurs, the myocardium having other sources of energy
available.* Glycolysis in dog muscle extracts proceeds somewhat differently
than in rat muscle extracts (e.g. accumulation of hexose phosphates occurs)
and the response to malonate is consequently different (Fawaz et at.,
1963). High concentrations of malonate cause the accumulation of the same
* Since the heart is generally considered to obtain much of its energy from fatty
acid oxidation under certain circumstances, it would be important to know the effects
of these high concentrations of malonate on this pathway. However, with the cycle
inhibited, the generation of energy from the oxidation of fatty acids should be reduced.
130 1. MALONATE
intermediates in dog muscle as described above for rat muscle, but in ad-
dition there is an inhibition of the accumulation of fructose-l,6-diP, so
that some inhibition of a proximal step in glycolysis seems likely. Of the
four enzymes involved previous to fructose-l,6-diP, only phosphogluco-
mutase is sensitive to malonate, 40% inhibition being produced by 60 mM
malonate and 76% inhibition by 120 mM malonate. Thus a secondary site
for the inhibition of glycolysis is likely. Other mechanisms may be involved
in intact muscle cells.
Effects on the Distribution of C" from Labeled Glucose
If glucose is metabolized exclusively by the Embden-Meyerhof glycolytic
pathway and no initial decarboxylation of glucose or the hexose phosphates
occurs (as it would if the pentose-P pathway were operative), malonate
should depress the formation of C^'^Og from glucose-1-C^* and glucose-6-C^*
equally. However, if the pentose-P pathway is important, malonate should
decrease the formation of C^^Og from glucose-6-C^* relatively more than from
glucose-1-C^*, and hence increase the C-l/C-6 ratio. The difference between
the C^'^Oo formed from these precursors is often taken as a measure of the
activity of the pentose-P pathway; this may not be strictly true because
every hexosephosphate which is decarboxylated may not pass through the
pentose-P pathway completely, but it is certainly the best evidence for
the relative importance of these two pathways.
Experiments of this type were performed with slices of various tissues
from the rat (van Vals et al., 1956). The specific activities of the C^^Og
formed from C-1 and C-6-labeled glucose are essentially the same in the
controls, indicating the pentose-P pathway to be unimportant (see ac-
companying tabulation). Malonate increases the C-l/C-6 ratios, which was
C-l/C-6 ratio
Tissue
Control Malonate 30 mM
Heart
0.95
2.38
Brain
0.93
2.18
Kidnev
0.99
1.92
Diaphragm
1.02
1.32
taken as evidence for a malonate-induced appearance of the pentose-P
pathway, although it would account for only a small fraction of the glucose
oxidized. In the lung and various mouse tumors, in which the pentose-P
pathway is operative normally (C-l/C-6 rations between 1.4 and 5.6),
malonate further augments the importance of the pentose-P pathway as
EFFECTS OF MALOXATE ON GLUCOSE METABOLISM 131
determined by C-l/C-6 ratios, which are increased to vakies between 4
and 125. A similar situation has been encountered in sheep thyroid shoes
(the results of three experiments are averaged in the accompanying tabu-
lation) (Dumont, 1961). The formation of C^^Oofrom glucose-6-C^* is inhibit-
Ci*0, Control Malonate 100 mM
From glucose-1-C^^ (cpm)
10.7
9.72
From glucose-6-Ci* (cpm)
4.1
0.47
C-l/C-6
2.58
20.5
(C-1) — (C-6)
6.6
9.25
ed strongly, whereas that from ghicose-1-C^^ is scarcely affected. This, of
course, indicates an almost complete block of the cycle, which is not sur-
prising at this high malonate concentration, but it also suggests a greater
participation of the pentose-P pathway in the presence of malonate. These
results are perhaps more understandable in the light of the glycolytic
inhibitions by high malonate concentrations discussed in the previous
section. Quite different results were obtained in electrically stimulated rat
ventricle strips (see accompanying tabulation) in which the pentose-P
Ci^O, Control Malonate 5.6 mM
From glucose- 1-C^* (cpm)
0.318
0.271
From glucose-6-C''' (cpm)
0.285
0.264
C-l/C-6
1.12
1.03
(C-1) - (C-6)
0.033
0.007
pathway is presumably not important, malonate at this concentration
having little effect on glucose utilization (Rice and Berman, 1961). It is
possible that higher concentrations of malonate would produce changes
such as observed with other tissues. However, this concentration of mal-
onate is quite effective in modifying ventricular function.
An indirect mechanism for the acceleration of the pentose-P pathway by
malonate may involve the levels of NADP and ATP in the tissues. The oxida-
tive decarboxylation of glucose-6-P to initiate this pathway requires NADP,
the concentration of which may be changed due to the action of malonate on
the cycle. Also the phosphorylation of fructose-6-P in the Embden-Meyerhof
pathway requires ATP, the level of which may be reduced by high concen-
trations of malonate. However, little is known about the control of the
pentose-P pathway and further experiments are needed to elucidate its
132 1. MALONATE
role during malonate inhibition. Nothing is known of the possible direct
effects of malonate on the pentose-P pathway. The production of C^^Og
from ribose-l-C^* is inhibited strongly by malonate in heart homogenates
(Jolley et al., 1958) and it is believed that ribose is metabolized in this
pathway, but the inhibition may reflect an action on the cycle. It is inter-
esting that fluoroacetate does not inhibit C^^Og formation very potently,
except at very high concentrations, so that some direct effect on ribose meta-
bolism is possible. The results of the experiments discussed above would
argue against this. D-Xylose and D-ribose-5-P are oxidized through se-
doheptulose-7-P in extracts of Pseudomonas hydrophila and this is not af-
fected by malonate at 20 mikf (Stone and Hochster, 1956).
The only investigation of the effects of malonate on the general distri-
bution of C^^from labeled glucose is that of Romberger and Norton (1961) in
potato tuber slices incubated with uniformly labeled glucose for 3 hr (Table
1-19). The situation in this tissue is complex inasmuch as fresh slices do not
metabolize glucose appreciably, whereas 36-hr-old slices oxidize it quite
rapidly. In the aged tissue, CO2 production is inhibited 92% by 50 mM mal-
onate at pH 5, while the formation of CO2 in fresh tissue is stimulated 28%.
In the fresh tissue, they suggest that CO2 is formed mainly in the pentose-P
pathway and little through the Embden-Meyerhof sequence, but glycolysis
contributes more and more with time, so that the marked inhibition by
malonate in aged tissue is not surprising. The synthesis of sucrose accounts
for over half of the labeling from glucose-u-C^* and this is inhibited only
11% by malonate. After 3-hr incubation, however, one can deduce little
about the initial attack on glucose. It may be mentioned in this connection
that in Acetobacter xylmum, where the sole product of glucose assimilation
is cellulose, malonate at 10 mM does not inhibit the formation of cellulose.
Laties (1964) has investigated this problem in detail and found that dif-
ferent methods of aging result in metabolically different potato slices,
in that some exhibit a malonate-sensitive and some a malonate-resistant
respiration. In the malonate-sensitive slices the formation of C^^Og from
labeled glucose is almost completely abolished by malonate, whereas in the
malonate-resistant slices there is little effect by malonate on the production
of C^^Og. There is a similar correlation with respect to the effects of malonate
on glucose uptake. Since dinitrophenol does not interfere with glucose
uptake, one can eliminate depression of ATP formation by malonate as
responsible for the inhibition of the uptake in sensitive slices. Laties con-
sidered the possibility that increased citrate levels might inhibit phos-
phofructokinase, but the mechanism is not yet well understood.
Effects of Tissue Age on the Response of Glucose Metabolism to Malonate
We have seen above that aging of potato slices increases their sensitivity
to malonate with respect to glucose oxidation. It might be expected that
EFFECTS OF MALONATE ON GLUCOSE METABOLISM 133
the metabolic characteristics of tissues, would change with age and that
this would be reflected in difl"erent susceptiblies to inhibitors. The altered
response to inhibitors could provide some information on the nature of
metabolic aging. The results obtained on rat brain are, however, discordant.
Tyler (1942), using a minced preparation, found that malonate inhibition
of the respiration in the presence of glucose increases up to a rat age of 10
days, after which it remains at the adult level (see accompanying tabula-
tion). Although the control respiration rises, the malonate-resistant fraction
Rat age (days)
Control
Malonate 10
mi¥
"o Inhibition
1
656
547
16.9
2
582
507
12.9
6
814
590
27.6
8
712
559
34.3
10
1080
530
50.9
16
1500
810
46.0
31
1900
960
49.5
Adult
1723
864
50.0
of the respiration remains constant up to 10 days, i.e., the increased respi-
ration is all due to the development of activation of a system sensitive to
malonate, presumably the cycle. On the other hand, Muir et al. (1959)
reported that the glucose respiration of adult rat brain slices is less sensitive
to 10 inM malonate (30% inhibition) than the respiration of tissue from
young animals of 1-3 days (70%). In this case, the young brain respires
almost 2.5 times as rapidly as adult brain. The differences in these obser-
vations may be related to the preparations used (mince or slice). There are
several reasons why malonate sensitivity would change with age, for exam-
ple, an alteration of cycle activity, the development or loss of pathways
other than the cycle for the metabolism of acetyl-CoA, or a change in the
ability to demonstrate a Pasteur effect. The effect of age should also be
studied on the electrically stimulated or K+-stimulated respiration of the
brain, since it is more sensitive to malonate and the results might have
more physiological pertinence.
The respiratory rates and patterns of fungus spores are altered during
the initiation of germination and the subsequent development of the germ
tube (Gottlieb, 1964). During the incubation of the spores of Penicillium
oxalicum and Ustilago maydis and the progress of germination, the respira-
tion in the presence of glucose rises markedly and this is accompanied by an
increasing sensitivity to malonate (see accompanying tabulation) (Caltrider
and Gottlieb, 1963). It is somewhat difficult to determine if this implies
134
1. MALONATE
an increase in cycle activity since the concentration of malonate was 100
mM and more than the cycle might be inhibited.
Incubation (hr)
Respiration
0/
/o
Inhibition
0
1.3
0
6
6.0
16
9
11.1
22
12
23.5
32
Effects on Resting and Stimulated Glucose Metabolism in Brain
The effects of malonate on a tissue may depend on the activity of the
tissue as well as the age. Stimulation of a tissue such as brain alters the
metabolic pattern and this is exhibited in altered responses to inhibitors.
The results obtained by Heald (1953) on guinea pig cerebral cortex slices
are shown in Fig. 1-14. The resting respiration and aerobic glycolysis are
( Malonote !
10 100
mM
Fig. 1-14. Effects of malonate on guinea pig brain
slices with glucose as the substrate. Electrical stimula-
tion through grids. The respiration and aerobic glycol-
ysis in //moles/g wet weight/hr. (From Heald, 1953.)
EFFECTS OF MALONATE ON LIPID METABOLISM 135
little affected by malonate up to 10 mM, whereas the electrically stimulated
respiration is readily depressed (down to the resting level at 10 mM mal-
onate) and the stimulated lactate formation markedly increased. This means
that the glucose metabolism appearing upon stimulation is quite sensitive
to malonate and perhaps involves a greater participation of the cycle.
These results were confirmed by Kimura and Niwa (1953) in guinea pig
brain stimulated by K+, and a stimulation of lactate formation by 10 mM
malonate was observed by Tsukada and Takagaki (1955). An abolition of
the inhibition of respiration upon addition of fumarate occurs (Takagaki et
at., 1958). Rat brain slices behave similarly, the resting respiration in the
presence of glucose being unaffected by malonate up to 0.8 mM, while the
stimulated respiration is readily suppressed (Wallgren, 1960). A malonate
concentration as low as 0.2 mM inhibits the stimulated respiration 15%.
Pyruvate utilization and the associated oxygen uptake are also inhibited
more strongly in stimulated slices than in resting slices (Takagaki et al.,
1958). The C^^Og from labeled pyruvate is formed about twice as rapidly in
high K+ medium compared to the controls (Kini and Quastel, 1959), and
this is inhibited more strongly by malonate in the K+-stimulated slices,
while the stimulated respiration is depressed to the endogenous level.
These results taken together clearly indicate a dependency of malonate
inhibition on the metabolic activity of brain tissue, whether altered by
electrical stimulation or K+. A Pasteur effect is observed and it is possible
that the inhibition by malonate would have been greater if it had not
induced a greater utilization of glucose. The data do not necessarily imply
a specific activation of the cycle; a greater uptake or utilization of glucose
would impose a greater load on the cycle, and this might be inhibited more
readily. Whatever the explanation for these effects, such results have
important bearing on the actions of malonate on intact and functioning
nervous tissue.
EFFECTS OF MALONATE ON LIPID METABOLISM
The major pathway for fatty acid oxidation is a helical degradation into
acetyl-CoA, which normally enters the cycle by condensation with oxal-
acetate. Each turn of the helix, releasing one acetyl-CoA, takes up 2 atoms
of oxygen, and the complete oxidation of acetyl-CoA through the cycle
takes up 4 more atoms of oxygen. Thus, approximately two-thirds of the
oxygen uptake due to fatty acid oxidation occurs in the cycle,* and one
would expect malonate to depress this fraction in proportion to the cycle
* The term "cycle." as before, will indicate the tricarboxylate cycle only, and the
pathway of degradation of fatty acids to acetyl-CoA and other terminal products
will be designated the "helix" for convenience.
136 1. MALONATE
block it induces. The situation is vey similar to that of ghicose oxidation in
this respect. Generally speaking, malonate could act on either the cycle,
or the helix, or both. Despite the extensive work that has been done on the
effects of malonate on fatty acid oxidation, direct information on the actions
on the helix is lacking, since the five reactions involved in each turn of the
helix and the enzymes associated with these have not all been tested for
susceptibility to malonate, nor has the operation of the helix dissociated
from the cycle been studied. Our evidence on this point must be indirect.
Before considering this evidence, let us outline some of the possibilities
for mechanisms of helix inhibition. Just as in the oxidation of glucose, ATP
is required for the initiation of the helix reactions and Mg++ is a necessary
cofactor (e.g., for the fatty acid thiokinase), so that malonate might depress
the operation of the helix by depleting the system of either of these sub-
stances. The extent of such an inhibition will depend on the availablity of
ATP or the presence of systems generating it, and on the concentration of
Mg++. Possibly a more important factor is the requirement for coenzyme A.
Malonate could deplete conzyme A by at least two mechanisms. The for-
mation of a relatively stable malonyl-CoA would remove some of the
coenzyme A from participating in the helix. A block of the cycle would
impede the entrance of acetyl-CoA into the cycle and the regeneration of
coenzyme A will depend on the enzymes present for the metabolism of
acetyl-CoA. The usual pathways for acetyl-CoA are (1) a simple splitting
to form acetate, (2) a transfer of the coenzyme A to another acid, and (3)
a condensation of two acetyl-CoA's to form acetoacetyl-CoA and eventually
acetoacetate. As in the oxidation of pyruvate through the cycle, the fate
of acetyl-CoA will depend also on the presence of reactions forming oxal-
acetate by pathways other than the cycle. The rate of fatty acid oxidation
can thus be limited by the rate of regeneration of coenzyme A. These
considerations lead one to ])redict that the effects of malonate on fatty
acid oxidation would be variable and dependant on the metabolic charac-
teristics of the tissue studied and the conditions of the experiment. This
prediction is borne out.
There is a good deal of evidence that malonate in concentrations up to
20 raM does not directly inhibit the reactions of the helix. Although an
inhibition of the oxygen uptake or the CO2 production during fatty acid oxi-
dation is not indicative of the site of inhibition when the helix and the
cycle are operating together, the absence of inhibition implies a lack of
action on the helix. Malonate at 16.8 mM has no effect on the C^^Oa arising
from palmitate-1-C^* in soluble extracts of peanut cotyledons (Castelfranco
et al., 1955), nor does 1 roM malonate have an effect on the oxygen uptake
associated with palmitate oxidation in peanut microsomes (Humphreys
et al., 1954). The anaerobic dehydrogenation of C4-C18 fatty acids in liver
homogenates with methylene blue as an acceptor is not inhibited by
EFFECTS OF MALONATE ON LIPID METABOLISM 137
10 raM malonate (Blakley, 1952). The rate of oxidation of decanoate by
Serratia marcescens is also not affected by 10 mM malonate, although
20 m.M inhibits somewhat (Waltman and Rittenberg, 1954). Geyer and
Cunningham (1950) stated that their data indicated no inhibition directly of
octanoate oxidation in liver by 5 mM malonate (this work will be discussed
in greater detail later).
On the other hand, Lehninger and Kennedy (1948) reported that 10 mil/
malonate stronglj' inhibits octanoate oxidation in particulate suspensions,
from rat liver. Not only is the respiration from the oxidation inhibited
but the utilization of octanoate is almost completely suppressed. The ad-
dition of malate or oxalacetate reduces the inhibition only partially, the
utilization of octanoate still being inhibited around 70%. It may be noted
that the total Mg+"'" concentration in these experiments was 0.25 mM,
which is quite low, so that malonate could have inhibited by depletion of
this cofactor. An interesting point is that the strain of rats used is very
important, since 10 mM malonate inhibits octanoate oxidation 25% in
preparations from livers of Sprague-Dawley rats, but in preparations made
from a heterogeneous stock colony 2 mM malonate inhibits 50-75%.
Such differences in strain behavior may explain some of the discrepancies in
the reports on malonate inhibition. Weinhouse et al. (1949) found that
20 mM malonate almost completely blocks the oxidation of octanoate in rat
liver slices and that fumarate only partially overcomes this, suggesting to
them that malonate must have some action other than on succinate oxidase.
In several instances malonate has been found to inhibit the oxygen uptake
from fatty acid oxidation very potently. Butyrate respiration in peanut
mitochondria is inhibited 75% by 6 mM malonate and the formation of
C^^Og from butyrate-l-C^* is depressed even more strongly (Stumpf and
Barber, 1956). Malonate at 10 mM inhibits the oxidation of octanoate by
carp liver mitochondria 80% (Brown and Tappel, 1959). In suspensions of
particulates from desert locust thorax, butyrate oxidation is inhibited 70%
by malonate at concentrations as low as 1 mM and maximally 85% (Meyer
et al., 1960). The oxidation of octanoate- 1-C^* and myristate-1-C^* by sub-
cellular particles from the lateral line of the rainbow trout, as measured
by the C^^Oj released, is reduced 95% by 10 mM malonate (Bilinski and
Jonas, 1964). One of the most sensitive systems is found in the oxidation of
linolenate in liver mitochondria of vitamin E-deficient chicks, 0.25 mM
malonate inhibiting 40% (Kimura and Kummerow, 1963). These examples
must be interpreted as indicating either a direct or an indirect inhibition
of the helix by malonate. Finally, it was stated by Mudge (1951), on the basis
of unpublished experiments, that malonate inhibits fatty acid oxidation
more strongly than succinate dehydrogenase in kidney particulate prepa-
rations. The possibility of effects on the helix must therefore be entertained
on the basis of our present knowledge.
138
1. MALONATE
Effects on the Formation of Acetoacetate and Other Ketones
It was known before 1912 that acetate, oxalate, and maleate can either
be metabolized to acetoacetate or so alter metabolism that acetoacetate ac-
cumulates, and for this reason Momose (1914) in Berlin studied the effects
of malonate perfused through starved dog livers at a concentration of
approximately 13 mM. He found that acetone appears and detected a sub-
stance which he only later, after returning to Japan (Momose, 1925), proved
was acetoacetate. However, he postulated that malonate -^ acetate — >■
acetoacetate -^ acetone, which was not unreasonable considering the inhi-
bitory action of malonate was unknown. The appearance of acetone in the
urine of rabbits fed malonate or rats injected subcutaneously with malonate
was observed by Huszak (1935), and simultaneously Annau (1935) dem-
onstrated that malonate causes the formation of acetone in slices and
breis of rabbit kidney. Acetoacetate has been shown to accumulate in
tissues as a response to malonate (see accompanying tabulation). Since
acetoacetate and acetone are the most important ketonic substances ap-
pearing in the tissues, these results clearlj^ show that malonate is ketogenic.
Preparation
Substrate
Malonate
Reference
Whole rabbits (blood)
Endogenous
1.6g/kg
Handler (1945)
Whole rats (blood)
Endogenous
O.S
!g/kg
Mookerjea and Sadhu (1955)
Rat liver slices
Acetate
40
mM
Jowett and Quastel (1935 c)
Butyrate
40
mM
Jowett and Quastel (1935 c)
Guinea pig liver slices
Propionate
40
mM
Jowett and Quastel (1935 c)
Butyrate
40
mM
Jowett and Quastel (1935 c)
Rat liver slices
Endogenous
10
mM
Edson (1936)
Rat liver slices
Fatty acids
5
mM
Geyer and Cunningham (1950)
Rat liver suspension
Fatty acids
10
mM
Lehninger (1946 a)
Rat liver homogenates
Pyruvate
4
mM
Recknagel and Potter (1951)
Rabbit liver mitochondria
Fatty acids
15
mM
Cheldelin and Beinert (1952)
Jensen rat sarcoma mince
Glucose
6.7 mM
Boyland and Boyland (1936)
Peanut mitochondria
Butyrate
6
mM
Stumpf and Barber (1956)
Acetoacetate is an important substance in intermediary metabolism and
the pathways for its formation and utilization are often complex. The con-
centration of acetoacetate will depend on the relative rates of its formation
and utilization. The accumulation of acetoacetate in the presence of mal-
onate could result from either an acceleration of its formation or an inhibi-
tion of its utilization, or both. The earliest concept that malonate itself
gives rise to the acetoacetate was soon abandoned, and several investigators
assumed that malonate interferes with the disposal of acetoacetate, while
EFFECTS OF MALONATE ON LIPID METABOLISM
139
recently more emphasis has been placed on the diversion of carbohydrate
and fatty acid metabolism to acetoacetate by malonate. We may summarize
some of the major pathways of acetoacetate before discussing the mecha-
nisms for the action of malonate (Ac-CoA = acetyl-CoA, and AcAc-CoA =
acetoacetyl-CoA). Reaction (1) for the formation of Ac Ac-Co A from aceto-
Fatty acids (
Pyruvate ij
Butyrate
-»- Ac -Co A
AcAc — CoA
-»- Ac Ac — CoA
Even-numbered
fatty acids
•-Butyryl —CoA *- Ac Ac — CoA
Succinate + Ac Ac— CoA
/3-Hydroxybutyrate
Amino acids (phenylalanine,
tyrosine, leucine, and
isoleucine)
Acetone + COj
.AcAc-CoA (1)
I acetoacetate I ' *-AcAc— CoA (2)
Sterols
/3-Hydroxybutyrate
acetate is catalyzed by an activating enzyme in the presence of CoA and
ATP, while reaction (2) is catalyzed by a CoA transferase in the presence
of succinyl-CoA. All of these reactions do not occur in a single tissue and
the response to malonate depends in part on which reactions are possible
in any case.
A block of the cycle restricts the entrance of acetyl-CoA, derived from
pyruvate and fatty acids, into the cycle, unless there is an adequate synthe-
sis of oxalacetate from a noncycle source, which is seldom the case. If the
acetyl-CoA accumulates, coenzyme A soon becomes tied up and the oxida-
tion of pyruvate and fatty acids would cease. However, in most tissues 2
molecules of acetyl-CoA condense to form acetoacetate and coenzyme A is
regenerated; in other situations, hydrolysis to acetate may occur. Malonate
may thus divert acetyl-CoA from the cycle to acetoacetate. Quantitative
conversion to acetoacetate has been observed (Recknagel and Potter, 1951).
Another reaction possibly favoring acetoacetate formation during malonate
inhibition results from the accumulation of succinate, which can now react
more readily with acetoacetyl-CoA in a transfer of coenzyme A. The effec-
tiveness of such a mechanism depends on the continued formation of suc-
cinate and, hence, usually on a noncycle source of oxalacetate. Acetoacetyl-
CoA is also formed as the terminal product of the helical oxidation of even-
numbered fatty acids. These relationships are illustrated in Fig. 1-15
where a block of succinate oxidation induces accumulation of acetoacetate
by accelerating its formation through two mechanisms. The other pathways
for the formation of acetoacetate are probably less important in most tis-
sues and would not be accelerated by malonate. Accumulation of aceto-
acetate implies that its utilization must not be too rapid. Liver is notable in
this respect because it lacks enzymes to metabolize acetoacetate rapidly,
140 1. MALONATE
especially the activating system for the formation of acetoacetyl-CoA, and
possesses an active deacylase to split acetoacetyl-CoA. Therefore, acetoace-
tate accumulation is most readily observed in liver and most investi-
gations have been on this tissue. The urinary acetoacetate found after the
administration of malonate is probably derived mainly from liver. In heart,
on the other hand, the enzyme balance is such as to favor the rapid me-
tabolism of acetoacetate and it does not accumulate. Acetate rather than
acetoacetate accumulates in some cells, for example in heart mitochondrial
suspensions metabolizing pyruvate in the presence of 8.8 mM malonate
(Fuld and Paul, 1952).
(Malonate)
a -Ketoglutorate ^ Succmyl- Co A Succinate -X"^ Fumorcte — ^ —^Citrate
Fatty acids Pyruvate
Fig. l-l.'j. Diagram of some pathways involved in the effects
of malonate on the metabolism of acetoacetate.
One would predict that fumarate should counteract the ketogenic activi-
ty of malonate because, by supplying oxalacetate, acetyl-CoA will again be
able to enter the cycle. However, it may be noted that fumarate may lead
to an even greater accumulation of succinate and if the formation of aceto-
acetate by the transfer of coenzyme A from acetoacetyl-CoA to succinate is
important, fumarate will only augment the malonate effect. Administration
of fumarate with malonate to rabbits abolishes the appearance of acetone
that arises with malonate alone (Huszak, 1935). Addition of fumarate to
malonate-inhibited minces of rat sarcoma likewise prevents the accumula-
tion of acetone bodies (Boyland and Boyland, 1936). However, fumarate
has very little effect on the appearance of acetoacetate in rat liver slices
inhibited by malonate (Edson, 1936), and this might indicate a mechanism
for malonate action other than the inhibition of succinate oxidation, or the
importance of the coenzyme A transfer reaction.
It is now easy to see how malonate can reduce the oxygen uptake and
the CO2 production from fatty acid oxidation without necessarily decreasing
the utilization of the fatty acids. A fraction that would normally be com-
pletely oxidized is diverted into acetoacetate (or acetate, acetone, and other
products). One of the best indications that malonate does not inhibit the
helix directly is the fact that the C^Oo appearing in the end products from
labeled fatty acid is not reduced by malonate. To illustrate this it will be con-
venient to turn to the excellent studies of Geyer and his group at Harvard.
EFFECTS OF MALONATE ON LIPID METABOLISM 141
The basic procedure in these investigations was to incubate carboxyl-
labeled fatty acids with rat liver and kidney slices, and determine the dis-
tribution of C^* in acetoacetate and COg. Malonate at 5 mM depresses the
formation of C^^Og from octanoate-C^* 00^ around 60% and fumarate is
able to overcome this inhibition only partially (Geyer et al., 1950 a). Fu-
marate and other cycle intermediates increase the total COg formed but
have little effect on the C^^Oa- This was explained by the accumulation of
some of the C^* as succinate, this not being relieved by fumarate, and we
have previously cited this as an example of the importance of considering
what is measured in demonstrating a reversal by fumarate.
Where does the C^* go that does not appear as C^^Oo in inhibited slices?
They found that in the presence of malonate much of the C^* appears in
acetoacetate (Table 1-24) (Geyer and Cunningham, 1950). The ratio AcAc-
QujQuQ^ is near 1.21 in the controls and is increased to around 4.51 by
malonate, averaging the results from the five fatty acids used. It may
also be noted that malonate generally increases the total C" recovered, even
though succinate was not determined, showing that malonate does not in-
hibit the fatty acid oxidation directly. Later they determined both the car-
boxyl and carbonyl C^* in acetoacetate and the more complete results are
summarized in Table 1-25, where I have taken the liberty of averaging
the data for all the fatty acids used, inasmuch as the effects are always
in the same direction although differences between the different fatty acids
are evident. These results show clearly the diversion of fatty acid metabo-
lism into acetoacetate by malonate. Weinhouse et al. (1949) reported that in
rat liver slices 10 mM malonate inhibits COo formation and no acetoacetate
appears, which was so contradictory to the results obtained by Geyer that
the latter studied malonate in concentrations up to 20 mM, but found only
that even more acetoacetate accumulates. Also they tested three different
strains of rat and the results were the same. The reason for this discrepancy
could not be explained.
The differential labeling in the carboxyl and carbonyl groups of aceto-
acetate is difficult to explain. If acetoacetate arises by a condensation of
acetyl-CoA units, the labeling in these positions should be uniform. However,
the ratio is seldom unity as may be seen in the results summarized by Chai-
koff and Brown (1954). In the work of Geyer with rat liver slices, the ratio
CHgC^^ 0 — / — CHoC^* 00" is less than 1 in the controls and increases with
the length of the fatty acid chain. Malonate increases this ratio, that is, it
increases relatively the labeling in the carbonyl group. Chaikoff and Brown
have given a detailed analysis of the possible factors determining this ratio,
and the explanation is based on the existence of two types of 2-carbon
fragment formed from fatty acids, one designated the CH3CO — fragment
and the other the — CH2CO — fragment. These fragments are assumed
to arise from different portions of the fatty acid chain and only the — CHg
142
1. MALONATE
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EFFECTS OF MALONATE ON LIPID METABOLISM
143
Table 1-25
Effects of 5 mM Malonate on the Oxidation of Fatty Acids in Rat Liver
Slices "
Control
Malonate
Change
C^Og produced
10,694
4,536
- 6,158
AcAc-carbonyl-C* formed
3,572
8,488
+ 4,916
AcAc-carboxyl-Ci^ formed
7,268
13,438
+ 6,170
CHaCi^O— /— CH^Ci^OO-
0.47
0.63
Total AcAc-Ci* formed
10,840
21,926
+ 11,086
C'^Oa + AcAc-C" formed
21,534
26,462
+ 4,928
AcAc-CiVC'^Oii
0.83
4.55
" Slices incubated 1 hr with carboxyl-labeled fatty acids shown in Table 1-24 at
38° and pH 7.1. The data from the five fatty acids were averaged to indicate the
general effects of malonate. (From Geyer et al., 1950 b.)
CO— fragments are believed to enter the cycle. The CHgC^* o_/— CHaC^^
00" ratio in acetoacetate will depend on the rates of production and utili-
zation of these two fragments. As pointed out, these two types of 2-carbon
fragment may be only convenient designations for two reactive forms of
acetyl-CoA. Malonate is assumed to increase the formation of acetoacetate
by condensation of randomized fragments of the — CHaC^^O — type, so
that the ratio rises. The extra acetoacetate formed over the control when
malonate is present does indeed exhibit a ratio of unity for hexanoate and
octanoate oxidation (Geyer et al., 1950 b). If malonate does this by blocking
the cycle, a preferential accumulation of — CH2CO — fragments would occur,
and a greater proportion of the acetoacetate would be formed from them.
The effect of malonate on acetoacetate accumulation will depend on the
pathway of fatty acid oxidation in the uninhibited tissue and, hence, on the
experimental conditions. For example, Witter et al. (1950) found that 3 mM
malonate inhibits acetoacetate formation from hexanoate 4% and 10 mM
malonate inhibits 9% in suspensions of washed particles from rat liver.
However, hexanoate is quantitatively converted to acetoacetate in the con-
trols, presumably because no cycle intermediates, are present to form oxal-
acetate for condensation of the acetyl-CoA units. Under such circumstances
malonate would not be expected to increase acetoacetate and the small
inhibitions must be attributed to actions directly on the helix. The rela-
tionship between acetoacetate formation in malonate-inhibited systems
and the presence of cycle intermediates was illustrated and discussed by
Cheldelin and Beinert (1952).
144 1. MALONATE
We have seen that the accumulation of acetoacetate in the presence of
malonate can be attributed to an increased rate of formation of the aceto-
acetate. Is the accumulation due entirely to this or can malonate also inhibit
the utilization of acetoacetate in some tissues? The rise in the acetoacetate
in liver slices in the presence of malonate was believed by Jowett and
Quastel (1935 c) to be due to the inhibition of the decomposition of aceto-
acetate, since at that time the pathways for the formation of acetoacetate
were not understood. However, Quastel and Wheatley (1935) soon provided
evidence that malonate can interfere with the disappearance of acetoacetate
in rat liver and kidney slices. In kidney slices, malonate at 8 mM inhibits
around 42% and at 16 mM 64%, and in liver slices an inhibition of 74%
was observed with 40 mM malonate. Fumarate is able to counteract this
inhibition partially and it was concluded that acetoacetate oxidation must
be coupled with other oxidations inhibited by malonate. Very similar results
were reported by Edson and Leloir (1936); indeed, 20 mM malonate inhibits
disappearance of acetoacetate in rat kidney slices 93% and it was stated,
"Malonate is a powerful and relatively specific inhibitor of respiration and
of aerobic disappearance of acetoacetic acid in kidney." Both Handler
(1945) and Mookerjea and Sadhu (1955) in their work with whole animals,
favored the concept that malonate interfered with acetoacetate metabolism
accounting for the rises in blood acetoacetate. Inasmuch as several different
pathways are open to acetoacetate and these vary with the tissue used, it is
difficult to interpret accurately these results. In some tissues, acetoacetate
can be split into acetyl-CoA fragments that enter the cycle and here malo-
nate might inhibit by blocking the cycle and the formation of oxalacetate,
which is, of course, esentially the same mechanism adduced to explain the
increased formation of acetoacetate. There is evidence that malonate does
not inhibit the reduction of acetoacetate to /?-hydroxybutyrate (Edson and
Leloir, 1936), nor does it seem to interfere with the formation of sterols from
acetoacetate (Mookerjea and Sadhu, 1955). It is probably best in the
present state of our knowledge to attribute the accumulation of acetoacetate
in the presence of malonate primarily to a diversion of 2-carbon units away
from oxidation through the cycle, without eliminating the possibility that
malonate may interfere in other pathways for the utilization of acetoacetate.
Effects on Propionate Metabolism
Propionate arises terminally from the /^-oxidation of odd-numbered fatty
acids and in certain tissues, such as the liver, can be oxidized completely
through the cycle. However, the oxidation of propionate differs from that
of other fatty acids. The following sequence of reactions has been suggested:
CO,
+
. ATP ATP B,j
Propionate — > propionyl-CoA — > methylmalonyl-CoA — > succinyl-CoA
EFFECTS OF MALONATE ON LIPID METABOLISM 145
The over-all reaction is the carboxylation of propionate to succinate. Other
pathways occur in bacteria, e.g.
Propionate -> propionyl-CoA -> acrylyl-CoA -> lactyl-CoA ->■ pyruvate -> COg + H2O
Such a sequence may also operate in animal tissues, since lactate was iden-
tified chromatographically after incubation of mouse liver slices with pro-
pionate (Daus et al., 1952). If the principal pathway of propionate is via
succinate, malonate would be expected to inhibit its oxidation readily but,
if acetyl-CoA is formed, the inhibition will vary with the conditions as
discussed for the effects of malonate on pyruvate oxidation.
Malonate has been shown to inhibit strongly the C^^Oo formation from
labeled propionate in mouse liver slices (Daus et al., 1952), rat liver slices
(Katz and Chaikoflf, 1955), suspensions of rabbit liver particles (Wolfe,
1955), and peanut mitochondria (Giovanelli and Stumpf, 1958), as anticipat-
ed. In the rabbit liver particulate preparation, 10 mM malonate suppresses
the formation of C^^O, from both propionate-l-C^* and propionate-2-C^^
almost completely, and at the same time leads to the accumulation of suc-
cinate, and in rat liver slices malonate also causes succinate accumulation.
From these data alone it is impossible to say whether the succinate arises
directly from propionate or is formed via the cycle, but the marked inhibi-
tion of C^^O, formation would indicate the former. This is substantiated
by the demonstration of labeled methylmalonate in the rat liver slices.
Another possible site for malonate inhibition is suggested by the work of
Flavin et al. (1955) on rat tissues. The intercon version of methylmalonate
and succinate was shown to be inhibited completely by 5 mM malonate and
thus malonate leads to the accumulation of methylmalonate during propion-
ate metabolism. However, it is not known if malonate can inhibit the
methylmalonyl-CoA isomerase, which catah^zes the interconversion in the
normal pathway, or if malonate only inhibits the formation of methyl-
malonyl-CoA from methylmalonate. The latter is reasonable because mal-
onate could compete with methylmalonate for the active site on the enzyme.
In peanut mitochondria the situation may well be different. Malonate at
6 mM inhibits the formation of C^^Oa from propionate- 1-C^* 41% (Giovanelli
and Stumpf, 1958). It was felt that this inhibition is not as much as would
be expected if the pathway from propionate leads to succinate. Further-
more, fluoride, which inhibits the carboxylation of propionyl-CoA, does
not depress the C^^Oa significantly. The pathway through methylmalonyl-
CoA to succinate may not be operative here, and the following pathway was
proposed:
Propionate ->• propionyl-CoA -> acrylyl-CoA -> ^-hydroxypropionyl-CoA ->•
/?-hydroxypropionate -> malonic semialdehyde -> malonyl-CoA -> acetyl-CoA
The COo formed in the last step derives from the carboxyl group of propion-
146 1. MALONATE
ate so that an inhibition of C^^02 formation from proprionate-l-C^* would
imply an action of malonate somewhere along this pathway. It is possible
that malonate, or malonyl-CoA formed from it, could compete with the
malonyl-CoA from propionate and in this way reduces the formation of
0^*02- It is known that the addition of methylmalonate simultaneously
with propionate depresses the propionate utilization strongly (Feller and
Feist, 1957). Lactating rat mammary gland preparations convert propion-
ate to fatty acids in part, the principal pathway being direct condensation
with malonyl-CoA to form the odd-chain fatty acids. Malonate at 10 vnM
inhibits this incorporation 50-65%, whatever the position of C^^ in pro-
pionate, and simultaneously C^^Og is depressed around 30% from proprion-
ate-l-C^* and propionate-2-C^^, and nearly 50% from priopionate-3-C^^
(Cady et al., 1963). This was interpreted not as a direct action on the pro-
pionate pathway but as a reduction of ATP or NAD(P)H, these being nec-
essary for fatty acid synthesis, as a consequence of inhibition of the cycle.
In Rhodospirillum rubrum, both the oxidation (Clayton et al., 1957)
and the photosynthetic dissimilation (Clayton, 1957) of propionate are
inhibited by malonate to approximately the same extent as are the similar
reactions of succinate, and this was given as evidence to support the meta-
bolism of propionate to succinate in these organisms.
Effects on Synthesis of Fatty Acids
There are at least three systems for the synthesis of fatty acids; one is the
reversal of the /^-oxidation in the helix and the other two involve the for-
mation of malonyl-CoA from acetyl-CoA, one mitochondrial and the other
nonmitochondrial (Green and Wakil, 1960). There are obvious relationships
between fatty acid synthesis and oxidative metabolism of various sub-
strates. The controls that establish the rates of fatty acid synthesis, or the
balance between oxidation and synthesis, have not been elucidated and it is
difficult to determine in a particular case what the effect of a cycle block
would probably be. The level of acetyl-CoA and the availability of the var-
ious pathways for its metabolism must be an important factor, but the
concentrations of coenzyme A, ATP, NADH and NADPH could also play
a significant role.
Malonate has been found to produce a variety of effects. Most of the
studies have involved the incubation of tissue preparations with acetate-
1-C^* and the subsequent determination of labeled fatty acids formed from
the acetate. In some cases a marked stimulation of fatty acid synthesis
in the presence of malonate has been observed. Malonate at 50 milf inhib-
its the O2 uptake of nonparticulate extracts of rat mammary gland and
yet increases the formation of fatty acids, sometimes as much as 10-fold
(Popjak and Tietz, 1955), The addition of oxalacetate or a-ketoglutarate
with the malonate increases the synthesis aven more (see accompanying
EFFECTS OF MALONATE ON LIPID METABOLISM 147
tabulation). The stimulating action of a-ketoglutarate was attributed to
the generation of NADH by which hydrogen atoms are provided for fatty
acid synthesis. Dils and Popjak (1962) claimed that malonyl-CoA is not
Incorporation of acetate into
fatty acids (m/< moles/ 100 mg)
Control 18.2
Malonate 118
Oxalacetate 72 . 7
Malonate + oxalacetate 241
a-Ketoglutarate 51.7
Malonate + a-ketoglutarate 517
formed from malonate in these extracts and that the stimulation of fatty
acid synthesis must be an indirect effect, possibly the suppression of the
deacylation of malonyl-CoA formed from acetyl-CoA, or the inliibition of
the decarboxylation of malonyl-CoA. Kallen and Lowenstein (1962) pointed
out that if this were the mechanism by which malonate acts, it should also
stimulate the synthesis of fatty acids from malonyl-CoA, which it does not;
indeed, malonate at 10 niM inhibits the conversion of malonyl-CoA into
fatty acids 33%. There is actually a stimulation of the formation of fatty
acids from acetyl-CoA. Furthermore, Spencer and Lowenstein (1962) found
that malonate is incorporated into fatty acids in an extramitochondrial
extract from rat mammary gland; acetate stimulates malonate incorpora-
tion just as malonate stimulates acetate incorporation. All of the stimula-
tion by malonate, however, cannot be explained by its conversion to mal-
onyl-CoA since several times more acetate than malonate is incorporated.
The varying effects of malonate on different preparations from a single
tissue are well illustrated in the studies of Abraham et al. (1961) with rat
mammary gland, where malonate stimulates fatty acid synthesis markedly
in cell-free systems (maximal stimulation around 130% at 17 mM malon-
ate), inhibits the synthesis 63% in slices, and has very little affect when
glucose is present. Glucose was assumed to provide NAD(P)H by forming
cycle substrates and also to augment the ATP level, which in the absence
of glucose might have been reduced by malonate. In the homogenates ATP
was added so that this aspect of the action of malonate could not be exhibit-
ed. The response to malonate is markedly dependent on the experimental
conditions, as shown by Hosoya and Kawada (1961) with human placental
slices, additions of estradiol, ATP, NAD, or bicarbonate altering the fatty
acid synthesis and its modification by malonate. It may be noted that fatty
acid synthesis in particulate preparations from the locust fat body occurs
rapidly only in the presence of malonate (Tietz, 1961).
148 1. MALONATE
So far we have considered total fatty acid synthesis. Separation of the
different fatty acids from animal tissues in malonate experiments has not
been done, but in mycobacteria malonate shifts the incorporation of acetate
into the higher fatty acids (Kusunose et al., 1960). The synthesis of total
fatty acid is moderately increased and this was attributed to the formation
of malonyl-CoA (see accompanying tabulation). Differential effects of mal-
Acetate-l-C* incorporation
Fatty acid % Change
Control Malonate 3.3 mM
Palmitate 7627
Stearate 2887
Arachidate 1144
Behenate 1089
Lignocerate 1862
Total acids 14609 17234 + 18
onate on the synthesis of short-chain and long-chain fatty acids are also
seen in rat liver slices metabolizing octanoate-1-C^* (Lyon and Geyer, 1954).
Although the over all effect of malonate on lipid synthesis in a particulate
preparation from avocado mesocarp is not marked, the incorporation of
acetate is shifted from stearate to oleate (see accompanying tabulation)
3488
- 54
1497
- 48
1332
+ 16
2110
+ 94
8807
-f373
Malonate
Acetate incorporation
into lipid
Distribution of label
(mM)
Palmitate
Stearate
Oleate
0
8.20
26
33
41
5
8.50
23
12
65
10
8.15
24
14
62
30
7.10
21
9
70
(Mudd and Stumpf, 1961). Although malonate may inhibit the cycle, this
may be counteracted by the formation of malonyl-CoA, which dilutes the
labeled malonyl-CoA formed from labeled acetate. It is interesting that
malonate is formed from acetate in avocado and this may be one regulatory
factor in fatty acid synthesis. Malonate has been found in three instances
to exert only inhibitory effects on fatty acid synthesis: in cell-free prepara-
tions from pigeon liver, 20 mM malonate inhibits the incorporation of
acetate into fatty acids 32% (Brady and Gurin, 1952); in various tumor
tissues (mammary and testicular carcinomata, and a sarcomatoid ovarian
EFFECTS OF MALONATE ON LIPID METABOLISM 149
tumor), 30 mM malonate inhibits 8-73% (van Vals and Emmelot, 1957);
and in rat liver extracts containing mitochondria, 10 mM malonate in-
hibits 51% (Iliffe and Myant, 1964).
These divergent observations are difficult to explain satisfactorily and
one must conclude that the final effects of malonate must depend on many
factors. In cellular preparations malonate may alter the levels of ATP,
NAD(P)H, and coenzyme A, as well as divert the metabolism of acetyl-
CoA by its inhibition of the cycle. In nonmitochondrial soluble enzyme
systems these actions would be minimized or absent, and the most important
factors might be the facilitation of fatty acid synthesis through the forma-
tion of malonyl-CoA or direct effects on the enzymes involved, although
there is no evidence for such direct affects at present. When malonate is
itself incorporated into fatty acids, as in several examples above and in
spinach chloroplasts, where malonate incorporation occurs at about half
the rate for acetate (Mudd and McManus, 1964), additional complications
must be considered. Since the incorporation of acetate- 1-C^* into lipid in
chloroplasts is reduced 71% by 0.67 mM malonate (Mudd and McManus,
1962), it would appear that malonate also inhibits some step or steps in the
pathway. The compartmentalization of the pools of acetyl-CoA, malonyl-
CoA, acetoacetate, and the various enzymes and cofactors within the cell
must be borne in mind in trying to explain certain differential affects of
malonate.
Effects on the Metabolism of Fats, Phospholipids, and Sterols
Several observations on total lipid response to malonate are interesting
even though it is impossible to assign a mechanism. In rat liver homogenates
incubated with palmitate-1-C^*, the lipids other than phosphohpids increase
22% in the presence of 10 mM malonate compared to controls (Jedeikin
and Weinhouse, 1954). Whether this is direct utilization of palmitate or
lipid synthesis with the C^^Og formed from palmitate is difficult to say.
Malonate at 50 mM also increases the total lipid content of potato tuber
slices some 230% (Table 1-19) (Romberger and Norton, 1961) and this
could be due mainly to an increased synthesis of fatty acids. On the other
hand, lipid synthesis from glucose-C^* in human placenta is depressed 75%
by 20 mM malonate (Hosoya et al, 1960). These results again show that
the action of malonate on lipid metabolism is quite variable. It will be more
profitable to turn to the synthesis of particular lipid fractions.
Injections of malonate lead to elevation of the free and esterified cho-
lesterol in the liver, kidney, and blood of the rat (see tabulation) (Mook-
erjea and Sadhu, 1955). Injections of 800 mg/kg of sodium malonate were
made daily for 3-4 weeks, some toxic effects being noted, and the animals
then sacrificed. Simultaneously, the blood glucose rose from 92 mg% to
196 mg% and the blood acetoacetate from 0.8 mg% to 3.6 mg%. Kidney
150
1. MALONATE
Free cholesterol
(mg/100 g wet wt.)
Esterified cholesterol
(mg/100 g wet wt.)
Contro:
Malonate
% Change
Control
Malonate % Change
Liver
Kidney
Blood
205
360
43
426
724
83
+ 107
+ 101
+ 93
73
72
68
104 + 42
228 +216
80 + 17
and liver slices showed impaired respiration with succinate, acetate, and
acetoacetate as substrates. This augmentation of tissue cholesterol is clear
and is reasonable on the basis of diversion of acetyl-CoA metabolism by a
block of succinate oxidase. However, in vitro work has shown only inhibi-
tion of cholesterol synthesis. The formation of labeled cholesterol from
octanoate-1-C^* in rat liver slices is consistently depressed by 5.84 mM
malonate, and fumarate was very ineffective in counteracting this inhibition
(Lyon and Geyer, 1954). The total lipids rise and this is partly attributable
to the increased synthesis of short-chain fatty acids. The formation of
labeled cholesterol from acetate- l-C^* in the same tissue is inhibited 73%
by 50 mM malonate (Kline and DeLuca, 1956) and 78% by 30 mM mal-
onate (van Vals and Emmelot, 1957). Cholesterol synthesis in rat tumors is
even more strongly depressed. The discrepancy between the in vivo and in
vitro results might be due to several factors. In the intact animal many
secondary effects may occur, e.g. as a result of the marked rise in blood
glucose. Also the malonate concentration in the tissues of the rats is undoubt-
edly less than in the work with slices. It is unfortunate that most of the
studies have been made with unreasonably high malonate concentrations
so that a specific inhibition of succinate oxidation is doubtful. The catabolism
of cholesterol, as determined by the formation of C^'^Og from the labeled
terminal methyl groups of cholesterol, in suspensions of rat liver mitochon-
dria is inhibited 78% by 10 mM malonate (Whitehouse et al., 1959), so
that this factor must also be considered in explaining changes in tissue levels
over longer periods of time. The synthesis of other sterols has been studied
very little. Pieces of rat adrenal form corticosteroids in the presence of
glucose and this is markedly stimulated by the addition of ACTH. Malonate
at 10 mM stimulates the formation of sterols in the absence of ACTH
from 17 to 22 //g/100 mg/2 hr ( + 29%) but depresses the synthesis in the
ACTH-activated preparations from 81 to 72 //g/100 mg/2 hr (-11%)
(Schonbaum et al., 1956). Fluoroacetate also inhibits very little and it was
concluded that the cycle does not play a major role in sterol synthesis,
glucose metabolism and particularly the pentose phosphate pathway being
of more importance. The bearing of such studies on the metabolic basis of
cholesterol and hormonal sterol levels in animals, especially the relationship
EFFECTS ON AMINO ACID AND PROTEIN METABOLISM 151
to the activity of the cycle and the other pathways for the utilization of
acetyl-CoA, warrants further investigations of the actions of malonate and
other cycle inhibitors both in vitro and in vivo. One approach to the met-
abolic defect in hypercholesteremia could be made in this way.
The incorporation of inorganic P^^ into phospholipids is almost invari-
ably inhibited strongly by malonate. This has been shown in peanut mito-
chondria (Mazelis and Stumpf, 1955), mycobacteria (Tanaka, 1960), guinea
pig brain dispersions (R. M. C. Dawson, 1953), rat liver mitochondria
(Marinetti et al., 1957), and other tissues. In cat brain slices, the effects of
malonate are very slight and it is possible that malonate does not penetrate
well (Strickland, 1954). Yet 3 mM malonate inhibits such incorporation
87% in K+-stimulated rat brain slices, although this may be due to a more
active cycle participation in the active tissue, inasmuch as respiration is
93% inhibited (Yoshida and Quastel, 1962). The phosphorylation of phos-
pholipid precursors probably involves the formation of high-energy phos-
phate compounds and malonate could depress this as the result of a block of
the cycle. A direct effect on the phosphorylation is unlikely. On the other
hand, the incorporation of activity into phospholipids from palmitate-1-C^^
in rat liver homogenates (Jedeikin and "Weinhouse, 1954) or from acetate-
1-C^* in rat liver slices (Kline and DeLuca, 1956) is affected scarcely at aU by
malonate. The phospholipids comprise a very heterogenous group and the
response to malonate probably depends on which type of phospholipid is
under investigation.
EFFECTS OF MALONATE ON AMINO ACID
AND PROTEIN METABOLISM
The pathways of amino acid metabolism often lead to or from the cycle
so that malonate would be expected to influence amino acid utilization and
formation by its inhibition of succinate oxidase. The intracellular accumu-
lation of amino acids and their incorporation into proteins are processes
requiring energy and consequently malonate could depress these important
reactions involved in cellular growth by a depletion of high-energy phosphate
derived from the cycle. Finally, malonate might act directly on the enzjones
catalyzing amino acid transformations. Information on these matters is
fragmentary but enough work has been done to demonstrate some interest-
ing effects of malonate on this phase of metabolism.
Effects on Amino Acid Metabolism
None of the enzymes involved directly in amino acid metabolism seems
to be very sensitive to malonate (Table 1-12) but a number of important
reactions have never been studied. Enz>Tnes catalyzing the reactions of
152 1. MALONATE
the dicarboxylic amino acids particularly might be expected to bind mal-
onate to some extent, but there is only indirect evidence for this. The
dehydrogenation of glutamate with methylene blue as an acceptor in
toluene-treated E. coli is inhibited about 20% by 71 vaM malonate (Quastel
and Wooldridge, 1928), but it may be that all of the hydrogen atoms do
not arise from glutamate here and that some other reaction is inhibited.
In Walker carcinosarcoma, kidney, and liver, glutamate is metabolized
readily to succinate in the presence of 6.3 milf malonate (Nyhan and
Busch, 1957), but no controls are avilable so that some inhibition is pos-
sible. Aspartate and glutamate are metabolized by Hemophilus parain-
fluenzae; malonate does not interfere with the oxidation of the former but
does inhibit glutamate oxidation (Klein, 1940). Contrary to these results,
malonate completely inhibits aspartate oxidation in rat liver homogenate
(Nakada and Weinhouse, 1950). It was believed rightly that this could not
be entirely attributed to an inhibition of succinate oxidase.
Glutamate may be converted to a-ketoglutarate by either glutamate
dehydrogenase or transamination, or it may be decarboxylated to y-amino-
butyrate; the decarboxylase is limited mainly to certain bacteria and the
nervous system of animals, so the major product is usually a-ketoglutarate,
which can be oxidized through the cycle or participate in transaminations
whereby it is reconverted to glutamate (this occurs also with y-amino-
butyrate so that the net reaction forms succinic semialdehyde, ammonia,
and CO2 from glutamate). The pattern of glutamate metabolism will depend
on the relative activities of these various enzymes, the availability of other
amino acids for transamination, and the supply of NAD for the glutamate
and a-ketoglutarate dehydrogenases; likewise, the response to malonate
inhibition will depend on these factors. If malonate selectively inhibits
succinate oxidation, the O2 uptake due to glutamate should be reduced
moderately (perhaps around 25-50%) unless much of the a-ketoglutarate
formed is transaminated and does not enter the cycle. Malonate, however,
occasionally inhibits the formation of ammonia from glutamate, indicating
some effect on the oxidative deamination. Malonate also inhibits the oxi-
dation of glutamate by guinea pig mammary gland mitochondria completely
(Jones and Gutfreund, 1961), which would not be the case if only succinate
oxidation were blocked. The glutamate respiration of rat brain mitochon-
dria is depressed 88% by 17.3 mM malonate (see note in Table 1-14)
(Levtrup and Svennerholm, 1963), which would indicate that glutamate
is being converted mainly to a-ketoglutarate by transamination (glutamate
decarboxylase is not present in brain mitochondria). Similar high inhibi-
tions by 20 mM malonate are observed in the mitochondria from pigeon
muscle, rat heart, rat liver, and ascites cells (64-99%) (Borst, 1962). Con-
clusions as to the pathway of glutamate catabolism based on the results
with malonate depend on the assumption that the inhibition is specifically
EFFECTS ON AMINO ACID AND PROTEIN METABOLISM 153
on succinate oxidation, and at these high concentrations this may not be
true. On the other hand, Das and Roy (1961, 1962) claim that transamina-
tion contributes little to the metabolism of glutamate in mitochondria from
Vigna sinensis, and since the decarboxylase is absent, oxidation by gluta-
mate dehydrogenase would seem to be the major route. Glutamate is con-
verted primarily to aspartate in rat brain homogenate via the pathway
glutamate -^ a-ketoglutarate -^ succinate -^ oxalacetate -^ aspartate
(Haslam and Krebs, 1963). The addition of fumarate removes this inhi-
bition, as expected.
Certain amino acids appear to be involved in the functioning of nerve
tissue and the effects of inhibitors on the metabolism of these substances
are of particular interest in this connection. Glutamate is accumulated in
brain and plays a role in the active transport of ions, while y-aminobutyrate
and A^-acetylaspartate have recently attracted attention because of their
ability to modify central nervous system activity. Glutamate and K"*" are
taken up by retina and brain slices in approximately equivalent amounts.
Malonate at 20 raM depresses the formation of glutamate + glutamine
only 12% while it reduces K+ uptake 40% (Terner et al., 1950), indicating
that the major effect of malonate is not mediated through interference with
glutamate. When guinea pig brain slices are incubated with glucose-u-C^*,
a good deal of the C^'* appears in amino acids, the most important of which
is glutamate (Tsukada et al., 1958). Malonate at 10 mTlf inhibits glutamate
formation around 25%, ^-aminobutyrate formation around 75%, and the
formation of aspartate appreciably. The total C^* incorporation into amino
acids from glucose-u-C^* in rat brain slices is inhibited 64% by 10 mM
malonate at normal K+ concentration but 83% in the presence of 105
mM K+, which produces an activation of brain metabolism (Kini and
Quastel, 1959). Such results can be readily explained on the basis of a mal-
onate-reduced pool of amino acid precursors due to the reduction in cycle
activity. Glutamate is a central substance in amino acid formation through
transaminations and anything which decreases the formation of a-keto-
glutarate would be expected to impair these pathways. Cremer (1964) has
recently found that 40 mM malonate not only reduces drastically the
incorporation of glucose-u-C^* into glutamate, aspartate, y-aminobutyrate,
and protein in brain slices, but also causes a loss of amino acids from the
cells. This concentration, of course, is probably not specifically inhibiting
succinate oxidation. A disputation type of reaction occurs in certain
tissues:
2 a-Ketoglutarate + NH3 + ADP + P, ^ glutamate + succinate + CO, + ATP
Tager (1963) used malonate to block succinate dehydrogenase and surpris-
ingly found that it augments the formation of glutamate in suspensions
of rat liver mitochondria (see accompanying tabulation). It was suggested
154
1. MALONATE
Control
Malonate 20 mM
A O2 (/^ atoms)
-1.2
- 1.2
A a-Ketoglutarate (^ moles)
-3.9
-10.4
A Glutamate (/< moles)
+2.0
+ 4.9
A Esterified phosphate (/< moles)
+ 1.9
+ 4.8
that malonate is converted to oxalosuccinate via malonyl-CoA. The oxalo-
succinate might function in the NAD- and NADP-dependent isocitrate
dehydrogenase systems to form a transhydrogenase so that NADPH is
the eventual donor for the formation of glutamate, oxalosuccinate acting
catalytically. Such studies show how complex the effects of malonate on
amino acid metabolism can be. The oxidation of certain amino acids pro-
ceeds via an initial transamination followed by degradation of the deaminat-
ed acids. The oxidation of y-aminobutyrate is completely inhibited by
1 n\M malonate in rat brain mitochondria (Sacktor et al., 1959) but in
Bacillus pvmilus is not affected even by 40 mM malonate (Tsunoda and
Shiio, 1959). Whether the former inhibition is the result of an indirect
suppression of transamination by cycle block or a direct effect on the oxi-
dative pathway of this amino acid is not known.
iV-Acetylaspartate occurs at a relatively high concentration in mamma-
lian and avian brain, increasing rapidly after birth. Its formation involves
direct acetylation of aspartate and the brain has little ability to metabolize
this substance. When acetate-1-C^* is injected intracerebrally, some of the
C^* is later found in A^-acetylaspartate (Jacobson, 1959). The injection of
malonate with the acetate reduces the incorporation of the C^* about 50%.
The injection of acetate depresses the level of total A^-acetylaspartate and
malonate counteracts this. These effects are quite complex and difficult to
interpret. The concentration of malonate injected was high (1.34 M) and
could have caused a severe fall in ATP so that acetate activation prior to
acetylation would be depressed. The rise in ^V-acetylaspartate seen with
malonate might have been due to a cycle block counteracting the effect
of the injected acetate, whereby cycle intermediates involved in trans-
aminations would be decreased, the level of aspartate being maintained with
more aspartate available for acetylation. There are so many pathways
associated with aspartate metabolism and acetylation reactions that the
final effects of a cycle block are difficult to predict. A good example of the
complex effects of malonate on amino acid metabolism is seen in Table
1-19, where certain types of amino acid in potato slices increase and other
types decrease during incubation with malonate (Romberger and Norton,
19C1).
EFFECTS ON AMINO ACID AND PROTEIN METABOLISM 155
Effects on Protein Synthesis
The intracellular synthesis of protein requires the simultaneous opera-
tion of many metabolic pathways and thus is susceptible to inhibition on a
variety of reactions. Some of the processes involved in protein synthesis
are: (1) the active uptake or accumulation of exogenous amino acids, (2)
the production of high-energy substances such as ATP from the oxidative
reactions of the cycle (except in anaerobes), (3) the formation of amino acid
precursors, again mainly by the operation of the cycle, and (4) all the com-
plex reactions for the activation and assemblage of the amino acids into
proteins. There are thus a multitude of possible sites for malonate action
but, at reasonable concentrations, the most important mechanism must
be a cycle block leading to both depletion of energy supplies and decrease
in amino acid precursors. There is no evidence that malonate can interfere
significantly either with the proteases or peptidases involved in the break-
down of proteins to amino acids or with the terminal assembling reactions
for the formation of protein.
The effects of malonate on the uptake and accumulation of amino acids
by cells have been studied in three types of tissue. Excised diaphragm main-
tains the same tissue/medium ratio for glycine as in the whole animal, and
the marked effects of 2,4-dinitrophenol indicate that glycine is concentrated
actively (Christensen and Streicher, 1949). Malonate, however, at concen-
trations of 3-55 TaM does not uniformly alter the tissue/medium ratio. It
is possible that malonate does not penetrate adequately, because muscle
is often rather impermeable to anions. The situation is different in Ehrlich
mouse ascites carcinoma cells. Glycine is accumulated so that tissue/medium
ratios are often 10-15. In two experiments, malonate at 37 mM decreased
this ratio from 13.0 to 5.1 and at 40 mM from 13.9 to 4.0 (Christensen and
Riggs, 1952). This occurred despite the fact that malonate increased the
synthesis of glycine. In cell suspensions of Gardner lymphosarcoma the
uptake of labeled glycine is inhibited 73% and the uptake of alanine 56%
by 10 mM malonate (Kit and Greenberg, 1951). These studies demonstrate
that malonate can interfere with protein s>Tithesis, at least in some cells,
by inhibiting the initial process of amino acid uptake.
The synthesis of protein is usually strongly inhibited by malonate, but
no analyses of the block have been made and the mechanisms are un-
known (see accompanying tabulation). The formation of adaptive enzymes
has often been taken as indicative of the synthesis of general cell proteins,
but this is not necessarily so, as pointed out by Mandelstam (1961). In E.
coli any substance acting as a substrate and source of energy represses en-
zyme synthesis, whereas inhibitors, such as malonate and 2,4-dinitrophenol,
counteract such effects and stimulate the synthesis. Furthermore, under
conditions in which /5-galactosidase synthesis is inhibited, the incorporation
of leucine-C^* into cell protein is not affected. The lack of inhibition in
156
1. MALONATE
Process
Malonate
(mi/)
% Change
Reference
Formation of induced /5-galacto-
sidase in E. coli
33.5
+ 31
Mandelstam (1961)
100
- 8
134
- 42
167
- 79
Incorporation of leucine-C*
into tobacco leaf proteins
10
0
Stephenson et al. (1956)
Protein formation in chick embryo
tissue culture
10
-100
Gerarde et al. (1952)
Incorporation of glycine-2-C^*
into rat liver homogenates
45
- 86
Peterson and Greenberg
Incorporation of glycine-C''
(1952)
into antibody in rabbit lymph
nodes
1.2
- 67
Ogata et al. (1956)
Incorporation of acetate- l-C^^
into rat liver slices
30
- 50
van Vals and Emmelot
Incorporation of acetate- l-C*
(1957)
into various tumor slices
30
— 74 to —90
van Vals and Emmelot
Incorporation of glutamate-u-C^*
(1957)
into Walker carcinosarcoma
6.25
-23 to -56
Nyhan and Busch (1957)
Incorporation of glycine- 1-C"
into chick embryo proteins
20
- 57
Quastel and Bickis (1959)
Incorporation of glycine- 1-C^*
into ascites protein
20
- 92
Quastel and Bickis (1959)
tobacco leaves was attributed to the presence of preformed precursors or
energy donors, so that interference with metabolism during the 2-hr in-
cubation does not modify the assembling of the proteins (Stephenson et al.,
1956). In two instances, glucose is able to partially reverse the effects of
malonate. Glucose addition to the Walker carcinosarcoma slices reduces the
inhibition by malonate, sometimes restoring the normal rate of protein
synthesis (Nyhan and Busch, 1957), and in ascites cell suspensions glucose
decreases the malonate inhibition from 92% to 14% (Quastel and Bickis,
1959), although the inhibition, is even increased slightly in chick embryo.
The marked glycolytic activities of tumor tissue may be responsible for this
phenomenon, sufficient energy for protein synthesis being obtained from
noncycle pathways.
The inhibition of amino acid uptake and the synthesis of proteins and
enzymes by malonate must be considered in long-term experiments or in
whole animal experiments, since this could secondarily affect many other
EFFECTS ON AMINO ACID AND PROTEIN METABOLISM 157
metabolic systems. Most enzymes are probably in a state of simultaneous
formation and degradation, so that an inhibitor of synthesis would induce
a steady fall of the enzyme level in the cells. This could apply, of course,
to all inhibitors of protein synthesis.
Effects on Urea Formation
The terminal product of much protein and amino acid metabolism is urea
and it has been found that under certain circumstances malonate inhibits
the formation of urea quite potently. The inhibition has been mentioned in
connection with its antagonism by fumarate (page 116). The most important
reactions of the urea cycle comj^rise the following, assuming that glutamate
is the immediate amino-group donor:
(1) Glutamate + oxalacetate ->• aspartate + a-ketoglutarate (transaminase)
(2) Aspartate + citrulline -j- ATP ->■ argininosuccinate + ADP + P,
(argininosuccinate synthetase)
(3) Argininosuccinate -> arginine + fumarate {argininosuccinase)
(4) Fumarate -> oxalacetate (fumarase and lyialate dehijdrogenase)
(5) Arginine -> ornithine + urea (arginase)
(6) Ornithine + NH3 + CO2 ~> citrulline {citrulline synthetase)
Glutamate + NH3 + CO2 + ATP -► a-ketoglutarate + urea + ADP + P,
This urea cycle thus makes contact with the tricarboxylate cycle at several
points. The a-ketoglutarate formed in the over all reaction can be oxidized
through succinate to oxalacetate or can be transaminated to regenerate glu-
tamate. The operation of the urea cycle thus requires sources for oxalace-
tate and ATP, both of which may be blocked by malonate.
Cohen and Hayano (1946) found that 5.7 raM malonate inhibits the con-
version of citrulline to arginine 90% in liver homogenates when glutamate
is the amino donor. The mechanism of the inhibition was not apparent
at that time. These results were confirmed by Fahrlander et al., (1947)
and, in addition, they showed that fumarate or malate, can counteact the
inhibition, indicating a block of succinate oxidation. They interpreted
the mechanism as a depletion of ATP and a consequent inhibition of reac-
tion (2). Subsequently, they showed that low malonate concentrations (1-2
ToM) inhibit urea formation as much as 75% and felt this was evidence for
a specific action on succinate dehydrogenase (Fahrlander et al., 1948).
The ATP level in the homogenates drops from 130 to 49.6 in the presence
of 2.5 yrM malonate and fumarate restores the ATP level to normal. It
was believed that glutamate not only furnishes the amino group but
also cycle substrates from which the energy is derived; a block by malonate
at the succinate level would reduce the amount of ATP formed. Krebs and
158
1. MALONATE
Eggleston (1948) then demonstrated a differential effect of malonate on
the formation of urea depending on whether glutamate or aspartate is
used as the amino donor, the inhibition being less in the latter case. An
elucidation of the true mechanism of the inhibition was presented by
Ratner and Pappas (1949), who showed a very definite differential effect
of malonate when glutamate and aspartate are used (see tabulation). The
% Inhibition
by malonate 20 laM
Substrate
Aspartate
Glutamate
Arginine synthesis Oj
uptake
Arginine synthesis Oj uptake
None
Pyruvate
Oxalacetate
Fumarate
a-Ketoglutarate
6
11
8
1
Stim 6
27
32
20
7
16
73 38
57 37
Stim 22 5
Stim 2 9
66 40
transamination forming aspartate from glutamate is not inhibited by mal-
onate so the mechanism must be sought elsewhere. It was proposed that
malonate prevents the formation of oxalacetate and thus indirectly blocks
the formation of aspartate; fumarate would, of course, overcome this block.
They opposed the idea that ATP depletion is important and felt that the
ATP derived from a-ketoglutarate oxidation would be sufficient. However,
they did not by any means disprove the ATP depletion hypothesis and it is
quite possible that it also plays a role in assigning an over all mechanism
for the inhibition. Miiller and Leuthardt (1950) extended these observations
by showing chromatographically that malonate inhibits the formation of as-
partate by reducing the formation of oxalacetate from a-ketoglutarate, and
also demonstrated conclusively that the transamination reaction itself is
not sensitive to malonate. It should be noted that in the reactions written
above, oxalacetate appears to be regenerated in the arginosuccinase reac-
tion followed by the hydration and oxidation of fumarate. but this is appar-
ently not sufficient to maintain the cycle, probably because much of the
oxalacetate disappears in other reactions. This is why an external source
of oxalacetate is necessary.
EFFECTS OF MALONATE ON PORPHYRIN SYNTHESIS
The pathway for the synthesis of porphyrins in both animals and plants
originates in the cycle in the condensation of succinyl-CoA with glycine
(Fig. 1-16). The succinyl-CoA can be formed either from a-ketoglutarate
EFFECTS OF MALONATE ON PORPHYRIN SYNTHESIS 159
or from succinate; the latter reaction requires ATP and is catalyzed by
succinyl-CoA synthetase (P-enzyme). A total of 8 molecules of succinate and
8 molecules of glycine is required for the synthesis of a molecule of proto-
porphyrin. The close connection between this pathway and the succinate
steps of the cycle, and the great iniportance of porphyrin synthesis in all
tissues, make the study of the action of malonate on this system interesting.
We may speculate on the various ways in which malonate could modify
porphyrin synthesis. (1) If the succinyl-CoA is formed in the cycle through
a-ketoglutarate, malonate could restrict its formation by blocking the cycle
Acetyl -Co A
Fumarote
(Malonate)
I
a-Ketoglutarote Succmote
-Succinyl -Co A
^^ Glycine
a -Ammo- /9 - ketoadipote
S - Aminolevulinote
I
Porphobilinogen
Protoporphyrin
Fig. 1-16. The pathways involved in
porphyrin biosynthesis.
and reducing the rate of acetyl-CoA entry, especially if no noncycle source
of oxalacetate is available. (2) If succinyl-CoA can be formed through the
cycle readily in spite of a malonate block, malonate might divert more suc-
cinate into the synthesis of porphyrin by inhibiting succinate oxidation.
(3) If the succinyl-CoA arises from succinate, this requires ATP and mal-
onate could deplete the system of ATP. (4) Malonate might deplete the
system of coenzyme A by the formation of malonyl-CoA. (5) It is possible
in some way that malonate might inhibit the formation of glycine, although
this is rather unlikely because there are usually several pathways avail-
able for glycine synthesis. The effects of malonate will thus depend on the
type of preparation used and the conditions of the experiment.
Duck erythrocytes (intact or hemolyzed) incubated with succinate and
glycine form porphyrin. Succinyl-CoA could be formed from succinate either
directly or through the cycle and the relative importance of these pathways
may be demonstrated by the use of succinate-C^'* with subsequent deter-
160 1. MALONATE
mination of the porphyrin labeling (Shemin and Kumin, 1952). Succinate-
C^^OO" when oxidized through the cycle gives rise to a-carboxyl-labeled
a-ketoglutarate and hence to unlabeled succinyl-CoA; therefore, no porphy-
rin labeling should result from this pathway. However, succinate-C"00~
could also directly form succinyl-CoA, which in this case would be labeled
and C^* would be found in porphyrin. On the other hand, succinate-C^^Hg
would form labeled succinyl-CoA by both pathways. If it is assumed that
malonate inhibits the oxidation of succinate and the cycle pathway only,
malonate should not inhibit any porphyrin labeling after incubation with
succinate-C^*00~, but should inhibit appreciably the porphyrin labeling
from succinate-C^^Hg. This was found by Shemin and Kumin, as the aver-
aged results in the accompanying tabulation show (figures are counts/
Hemin Control Malonate 20 mif % Change
From succinate-C'^OO- 215 210 - 2
From succinate-Ci^Hj 1025 421 -59
minute for intact erythrocytes). It would appear that both pathways are
operative in these cells. The failure of malonate to increase the porphyrin
labeling from succinate-C^*00~ is rather surprising because one might
expect malonate to divert some of the succinate from the cycle into the
formation of succinyl-CoA. It is possible that this effect is somewhat coun-
teracted by an inhibition on succinyl-CoA synthetase.
Further information on porphyrin synthesis and the effects of malonate
were obtained by Wriston et al. (1955) by the use of labeled acetate. Different
malonate effects were obtained when methyl-labeled and carboxyl-labeled
acetate were incubated with glycine, the inhibition of porphyrin labeling
being much greater with the former (see tabulation). This is the expected
Hemin Control Malonate 20 mM % Change
From acetate-Ci^Ha 376 174 -54
From acetate-C'^OO- 72.5 67 - 8
result, because the formation of labeled succinyl-CoA from acetate-C^*00~
does not involve the complete cycle and the C^* pathway does not go through
the succinate oxidation step, whereas porphyrin labeling from acetate-
C^^Hg depends on the operation of the entire cycle (except for the contri-
bution from the y-C atom of a-ketoglutarate). Furthermore, the labeling
EFFECTS OF MALONATE ON PORPHYRIN SYNTHESIS 161
in the porphyrin from acetate-C^^Hg should be altered by malonate.
Labeling in the carbon atoms of the A and B pyrrole rings of protoporphyrin
occurs after incubation of duck erythrocytes with acetate-C^^Hg and glycine.
CH, 9
II
6 H3C CH 8
' I I
' fi^N \
I
H
Control
Malonate 10
mM
% Inhibition
Total porphyrin
186,000
54,000
71
Pyrroles A and B
88,000
27,500
69
Carbon 4
13,000
3,700
72
Carbon 5
12,000
900
93
Carbon 6
20,000
9,400
53
Acetate-Ci^Hg will lead directly to -OOC-C^^HaCHa-CO-CoA and if the
cycle is blocked completely by malonate, carbons 6 and 9 only will be
labeled, except for some labeling of carbons 4 and 8 due to the reversible
reaction succinyl-CoA :^ succinate (as long as ATP is available). Carbons 2,
3, and 5 should not be labeled. This is essentially seen in the tabulation.
The cycle, of course, is not blocked completely so that some labeling in
carbon 5 occurs. The over all inhibition is due to a depression of the entry
of acetate into the cycle. These experiments not only show the variable
effects of malonate on a pathway associated with the cycle but well illus-
trate the use of an inhibitor to elucidate a metabolic pathway.
The analysis of the action of malonate on porphyrin synthesis was ex-
tended by Granick (1958) in his work with chicken erythrocytes. The for-
mation of protoporphyrin is innibited 90% by 10 mM malonate when only
glycine is present, 85% when succinate is added, and 80% when a-keto-
glutarate is added. The effects of different concentrations of malonate are
shown in Fig. 1-17. Malonate could decrease the incorporation of succinate
into porphyrin by blocking the cycle and reducing the ATP level, and thus
inhibit both pathways of succinate-CoA formation from succinate. However,
the quite strong inhibition of protoporphyrin formation from glycine
-f a-ketoglutarate is surprising. Inhibition of the step a-ketoglutarate — *
succinyl-CoA is not likely as the only explanation, because 1 raM malonate
inhibits protoporphyrin synthesis 32% and there is no reason for thinking
that this low concentration would inhibit a-ketoglutarate oxidase. The for-
162
1. MALONATE
mation of protoporphyrin. from S-aminolevulinate is not inhibited by mal-
onate so that an action on this part of the pathway is excluded. Granick
suggested that malonate reacts with coenzyme A and thus depletes the
system so that succinyl-CoA cannot be so readily formed. These effects
were confirmed in lysed chicken erythrocytes by Brown (1958). Porphyrins
are not formed in these preparations but (5-aminolevulinate is formed from
glycine and succinyl-CoA derived from a variety of cycle substrates. Mal-
onate at 10 mM inhibits this reaction 30% when the incubation is with
glycine and citrate, substantiating the action on this region of the pathway.
1 50
2 I
A Arsenite
B DNP
C Fluoroacetote
D Malonate
pi ►
Fig. 1-17. Effects of four inhibitors on the sjti thesis
of protoporphyrin in chicken erythrocytes with the
substrates as indicated. (From Granick, 1958.)
It is interesting that the addition of succinate to glycine and citrate in the
incubation medium leads to an inhibition of (5-aminolevulinate synthesis.
This was shown to be due to the formation of oxalacetate, which inhibits
a-ketoglutarate oxidase. Malonate is able to overcome this inhibition by
succinate through the prevention of oxalacetate formation. This indicates
another minor mechanism for the effect of malonate on porphyrin synthesis,
namely, the reduction in oxalacetate concentration and a consequent release
from any inhibition on the oxidation of a-ketoglutarate. Finally, we may
note that coproporphyrin synthesis from glycine and a-ketoglutarate in
EFFECTS ON METABOLIC PATHWAYS 163
Rhodopseitdomonas spheroides is inhibited 50% by 20 mif malonate and
75% by 40 mM malonate (Lascelles, 1956), indicating again some inhibition
of the formation of succinyl-Co A.
The incorporation of iron into heme, as demonstrated with Fe^^, is not
inhibited by 10 mM malonate in canine reticulocytes (Yoshiba et al., 1958)
but is inhibited 26% in chicken erythrocytes (Kagawa et al., 1959). It is
possible in the latter case that the inhibition is due to the chelation of part
of the Fe^^, making it unavailable for incorporation.
EFFECTS OF MALONATE
ON MISCELLANEOUS METABOLIC PATHWAYS
There have been many reports on the actions or lack of action of malonate
on enzyme reactions or metabolic pathways of varying degrees of impor-
tance. Some of these are worth mentioning, either because they indicate
areas where further study might be profitable or because they provide some
evidence for noncycle actions of malonate.
One might expect very little effect of malonate on photosynthesis but,
although very little work has been done, in every case some effect has been
observed. Even the Hill reaction is susceptible to inhibition (Ehrmantraut
and Rabinowitch, 1952). This reaction is the photochemical oxidation
of water with the production of oxygen and the reduction of a substance,
usually quinone, other than COg. In Chlorella this reaction is inhibited
30% by 6 mM malonate and 50% by 60 mM malonate. The inhibition is,
surprisingly, prevented by fumarate, indicating that the site of action is
succinate dehydrogenase and that this enzyme takes part in the transport
of hydrogen in the Hill reaction, which would not be the case if quinone
were serving as the immediate hydrogen acceptor. It may also be that
malonate does not inhibit the Hill reaction directly, but depletes the cells
of cycle intermediates or other cycle products necessary for the Hill reaction
to proceed. Malonate not only inhibits the total incorporation of C^^Og in
Scenedesmus by about 20%, but almost completely blocks the formation of
labeled malate (Bassham et al., 1950). This was taken as evidence that mal-
ate is not on the direct line of phosphoglycerate synthesis, but it also dem-
onstrates that by some mechanism malonate can inhibit COg incorporation.
An inhibition of glucose formation in Chlorella by malonate has also been
reported (Kandler, 1955), although there is less inhibition in the light than
in the dark. The synthesis of glucose was believed to be closely related to
the formation of high energy phosphate intermediates, and it is thus inter-
esting that malonate inhibits the photosynthetic phosphorylation of ADP
in Rhodospirillum, although the inhibition is only 17% at the very high
concentration of 100 mM (Smith and Baltscheffsky, 1959). With these lim-
ited observations on the effects of malonate, it must be admitted that it
164 1. MALONATE
is difficult to fit the data into the modern concepts of the carbon pathway in
photosynthesis, which does not directly involve the cycle, and particularly
to understand the mechanism whereby the Hill reaction is inhibited.
Malonate is also able to interfere in the metabolism of glycerol. Glycerol
is fermented to succinate, accompanied by the uptake of CO2, in Propioni-
hacterium pentosaceum, and 30 mM malonate inhibits both the glycerol
fermentation and the CO2 uptake around 10% (Wood and Werkman, 1940).
It is impossible to attribute this to an action on succinate dehydrogenase.
The oxidation of glycerol may involve an initial phosphorylation with
subsequent formation of pyruvate and entry into the cycle:
Glycerol + ATP -> glycerophosphate -> pyruvate -> cycle
The phosphorylation is inhibited in rat liver homogenates (Ruffo and D'A-
bramo, 1952), but the total oxidation is not markedly affected in the
mycobacteria (G. J. E. Hunter, 1953). Malonate at 10 mM inhibits 5% in
M. stercoris and stimulates 4-6% in M. smegmatis and M. butyricum. It is
surprising that greater inhibition is not observed if the oxidation does
involve the cycle. In castor bean cotyledons, malonate at high concentra-
tions has very marked effects on the utilization of glycerol (Beevers, 1956).
At 70 mM the C^^Oa formation from labeled glycerol is inhibited 97% and
the sucrose formation is inhibited 82%, while at 130 mM the oxygen
uptake is inhibited 60%. Such high concentrations may interfere with the
formation of ATP and hence depress phosphorylation and also block the
cycle CO2 release.
The stimulation of muscle respiration by insulin is inhibited potently by
malonate (Stare and Baumann, 1940). Insulin almost doubles the respiration
of minced breast muscle from depancreatized pigeons and 1 mM malonate
inhibits this increase 92%. Fumarate is able to overcome both this inhibi-
tion and the inhibition of nonstimulated respiration completely. This
interesting action in vitro led to a study of the antagonism in the whole
animal. Solutions of sodium malonate were injected subcutaneously in
rabbits either before or with insulin and the drop in blood glucose was much
less than with insulin alone. Insulin (4 units) decreases the blood glucose 65%
in 4 hr, whereas with malonate present the reduction is only 13% and none
of the rabbits goes into convulsions. Malonate alone increases the blood
glucose 26%. The respiratory inhibition in the muscle mince could be
explained on the basis of a typical cycle block (although the degree of inhi-
bition is surprising for a concentration of 1 mM), but the inhibition of glu-
cose utilization in the animal is more complicated. The initial phosphoryla-
tion of glucose and its uptake could have been inhibited indirectly by a re-
duction of the available ATP, or it could have resulted, at least in part,
from a hyperglycemic action of malonate unrelated directly to the insulin
stimulation.
EFFECTS ON METABOLIC PATHWAYS 165
Succinate and propionate are formed anaerobically in Ascaris muscle
from glucose and lactate, presumably by the following pathway:
Glucose ^
^>fc^ +CO, +4H „ -COj ^
Pyruvate »=- Oxalacetate *- Succinate *~ Propionate
Lactate -""^
Malonate at 20 mM does not appreciably inhibit the decarboxylation of suc-
cinate to propionate (about a 13% reduction in total radioactivity) but the
small inhibition indicates a possible competition with succinate for the
enzyme. However, the incorporation of lactate-2-C'-^ into succinate is inhibited
almost 90%. If succinate is formed by reduction of fumarate derived from
oxalacetate, malonate would be expected to inhibit well, not only because
of the effect on the succinate dehydrogenase but also by an inhibition of
oxalacetate formation. Malonate inhibits the formation of labeled propionate
from lactate-2-C^* 65%. The smaller inhibition compared to that for suc-
cinate formation implies another less important pathway for the formation
of propionate, perhaps by direct reduction, as shown in several bacteria.
The metabolism of glyoxylate by Kver mitochondria is rather complex;
it is decarboxylated to formate by a devious route, it may be oxidized to
oxalate, or it may be aminated to glycine (Crawhall and Watts, 1962).
Malonate inhibits the decarboxylation competitively but does not interfere
with the formation of oxalate or glycine; indeed, the latter may be stimul-
ated slightly due to diversion in a branched chain. The decarboxylation
reaction, which requires glutamate, is quite sensitive to malonate, around
50% inhibition occurring at 0.15 mM, both substrates being at 3 mM.
This would certainly appear to be one system in which a marked effect can
be exerted by malonate at low concentrations and which is unrelated to
succinate oxidation.
The synthesis of acetylocholine is an endergonic process and is related
to the cycle both for the supply of energy and with respect to the utilization
of acetyl-CoA. The effects of inhibitors on acetylcholine synthesis and hy-
drolysis are particularly important when considering the mechanisms by
which malonate can alter nerve and muscle function. Unfortunately, only
one study of the action of malonate has been made (Torda and Wolff,
1944 a). The formation of free acetylcholine in minced frog brain, in the
presence of physostigmine to prevent hydrolysis, is inhibited 32% by 0.08
mM, 46% by 0.8 mM, and 49% by 8 mM; the inhibition of total acetyl-
choline is about the same. Succinate, fumarate, and citrate increase acetyl-
choline formation. It would be interesting to investigate the effects of mal-
onate on the purer enzyme systems now available for acetylcholine synthe-
sis to determine if the inhibition is a direct effect or secondary through
ATP depletion. The effects of malonate in the intact cell may be quite
complex, because malonate might suppress the incorporation of acetyl-CoA
166 1. MALONATE
into the cycle and thereby lead to a greater availability of acetyl-CoA for
choline acetylation. However, it is not known if the acetyl-CoA pool is
common to both the cycle and the synthesis of acetylcholine. In many
cases the effects of malonate are due only to a depression of the cycle oper-
ation and a decreased formation of ATP. For example, in the synthesis of
chondroitin sulfate in tibial condyles of chick embryos, the fixation of sulfate
is inhibited by malonate in a parallel fashion to the inhibition of respiration
(Boyd and Neuman, 1954). The fixation of sulfate requires ATP, as shown
by the marked inhibition with 2,4-dinitrophenol, so that here the mechanism
of malonate action is simply an inhibition of energy formation. Other proc-
esses, such as calcium deposition in tibial cartilage (Hiatt et al., 1953),
do not require energy and are not inhibited by malonate.
EFFECTS OF MALONATE
ON THE ENDOGENOUS RESPIRATION
The alterations of the most important metabolic pathways by malonate
have been discussed, and we shall now conclude this aspect of the subject
with a survey of the effects on the total oxygen uptake of cells respiring in
the absence of any external substrate. Although the interpretation of the
results of such studies is very difficult, the changes in the endogenous respi-
ration have been examined more frequently than any other response to
malonate. There is, thus, a vast and variable mass of data, some of which
is summarized in Table 1-26. The aim of most of these investigations has
been to demonstrate the absence or presence of the cycle in the types of
cells tested, and we must attempt to assess the validity of conclusions based
on the response to malonate. The most unsatisfactory work has been done,
and the most unjustified conclusions have been drawn, in studies of this
type, inasmuch as the inherent complexities of the situations have seldom
been appreciated. Although the cycle has a wide distribution in the cells of
microorganisms, plants, and animals, its operation during the metabolism
of endogenous substrates is quite variable and dependent on the state and
past history of the cells.
Factors That May Determine the Degree of Malonate Inhibition
Certain basic factors should be considered in every investigation of the
susceptibility of the endogenous respiration to malonate. Although some
of these have been mentioned previously and some will be taken up in
greater detail later, it may be convenient to enumerate here the most im-
portant.
(a) Intrinsic susceptibility of succinate dehydrogenase to malonate. This en-
zyme from different species varies a good deal in its ability to bind mal-
EFFECTS ON THE ENDOGENOUS RESPIRATION 167
onate, as is evident from the range of K^ values observed (page 33). It is,
perhaps, too often assumed that in every organism the succinate dehydro-
genase wiU be readily blocked by malonate and that the inhibition of the
endogenous respiration depends only on the importance of the enzyme in
the total oxygen uptake. It is mandatory to demonstrate the sensitivity
of succinate dehydrogenase to malonate in the preparation being studied.
(b) Degree to which intracellular succinate dehydrogenase is inhibited. In
addition to the intrinsic susceptibility, there are other factors which can
alter the inhibition occurring within the cell. The concentration of succinate,
both initially and following the accumulation resulting from the inhibition,
may be high enough to oppose the malonate effect appreciably. The relative
stability of plant respiration to malonate has been attributed to the high
concentrations of succinate and other organic anions in plant cells. It is
probably very seldom that an inhibition even approaching completeness
can be achieved in cells at concentrations likely to be specific.
(c) Specificity of malonate inhibition. Inhibition of the endogenous res-
piration can, of course, arise from actions other than on succinate dehydro-
genase. If the object of the study is to evaluate the contribution of the
cycle to the oxygen uptake, inhibitions on noncycle pathways must be
eliminated. At the high malonate concentrations often used (see Table 1-26),
there is certainly no assurance that the inhibition is specific.
(d) Intracellular concentration of malonate. Malonate does not penetrate
readily into most cells, especially at physiological external pH values, so
that the internal concentrations of malonate may be far below those in
the surrounding medium (see page 190). The degree of respiratory inhibition
observed is probably often more of a measure of malonate penetrability
than of the nature or susceptibility of the metabolic systems. There are
several instances in Table 1-26 in which the inhibition rises with destruction
of the normal tissue structure or the removal of permeability barriers. In
general, the inhibition is greater in homogenates than in minces, and greater
in minces than in slices or intact cells. The results of Bonner (1948) on
Avena coleoptiles are interesting in this regard. Soaking in water for 24 hr
increases the susceptibility to malonate and removal of the endosperm
further increases the inhibition. The many observations that a lowering
of the pH augments the inhibition also provide evidence of the importance
of permeability.
(e) Metabolism of malonate. Many tissues and organisms can metabolize
malonate to acetyl-CoA, the oxidation of which contributes to the oxygen
uptake (see page 228). Some of the respiratory stimulations noted with
malonate must be due to this, and it is likely that the experimentally de-
termined inhibition in other cases is reduced from that which would be ob-
served if malonate were not metabolized. The best way to test for and
168
1. MALONATE
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180 1. MALONATE
correct for this phenomenon is to determine the C^^02 formed from labeled
malonate.
(f) Nature of the cycle operation and the presence of alternate pathways.
The inhibition of the oxygen uptake associated with the cycle will depend
on a number of factors in addition to the inhibition of succinate dehydro-
genase. The availability of a large pool of organic acids to form oxalacetate,
or the presence of pathways from which oxalacetate may arise (e.g., by
carboxylation of pyruvate, or from aspartate by transamination), will re-
duce the inhibition of oxygen uptake from that which would be observed
if all the oxalacetate had to be derived from the cycle. Other pathways
for the metabolism of succinate may circumvent the block to some extent.
These matters have been discussed in some detail (see pages 72-88).
(g) Adaptive changes in the presence of malonate. Inhibition of the cycle
may accelerate other pathways. The increased uptake and metabolism of
glucose brought about by malonate have been noted in several types of
cells, and such a phenomenon will tend to counteract the malonate inhi-
bition on the oxygen uptake. Adaptive changes in enzyme concentrations
probably are seldom important in short-term experiments but cannot be
completely ignored in work with certain microorganisms. Inhibitions by
malonate have occasionally been noted to decrease with time, and adaptive
changes are the most obvious explanation.
These and other more subtle factors determine the effect of malonate on
the total oxygen uptake of a preparation, and it should be apparent that
deductions based exclusively on the inhibitions of endogenous respiration
are frequently untenable. A definite inhibition with a reasonable malonate
concentration is more significant than a negative result, because there are
many factors which can reduce or abolish the action of malonate even
though the cycle is present and active.
The Time Course of Malonate Inhibition
The inhibition of respiration by malonate may occur fairly rapidly and
remain constant, or it may increase slowly to a level at which it is main-
tained, or it may gradually disappear, or it may vary in quite complex
fashion with time. A slowly developing inhibition would not be unexpected
with a substance which does not penetrate readily. The inhibition of suc-
cinate dehydrogenase is essentially instantaneous so that an approximately
linear increase in the inhibition to a constant level would imply that the
rate of inhibition is determined by the penetration. On the other hand,
secondary effects, such as would result from the depletion of ATP, may
also contribute to a progressive inhibition. Greville (1936) noted that mal-
onate does not immediately inhibit the respiration of rat diaphragm but
EFFECTS ON THE ENDOGENOUS RESPIRATION 181
that the inhibition developes over 1-2 hr. A very similar time course was
observed in barley roots by Laties (1949 a). After the addition of 10 mM
malonate, the respiration drops linearly for 60-90 min and then becomes
relatively constant at about 40%. However, over the next 5 hr, the inhi-
bition lessens somewhat. This was postulated to be due to increasing suc-
cinate concentration, but actually such changes in succinate take place
much more rapidly in most cases. It must be admitted that in spinach
leaves the maximal succinate accumulation occurs at 4 hr (Laties, 1949 b),
so that this explanation can by no means be eliminated. The inhibition
of the spinach leaf respiration by malonate is greater at 6 hr than at 3 hr
but not enough data are available to correlate the changes in inhibition
with succinate levels. Malonate requires about 1 hr to produce its maximal
inhibition of sea-urchin egg homogenate respiration (Yeas, 1950). This is
the only report of such a slowly developing inhibition in subcellular pre-
parations and no explanation is evident. Another situation seems to exist
in bovine kidney culture cells, where the inhibition by 100 mM malonate
slowly increases from 23% at 1 hr to 35% at 5 hr (Polatnick and Bachrach,
1960). It is more likely here that secondary changes are responsible.
Definite decrease in the respiratory inhibition of pigeon brain dispersions
(probably similar to homogenates) with time was reported by Banga et al.
(1939). The inhibition is 41.7% at 10 min, 29.5% at 30 min, and 20.5%
at 50 min. Since the malonate concentration was 24 mM, it seems unlikely
that metabolism of malonate could have reduced its concentration signifi-
cantly. Adaptive changes in homogenates are improbable and sufficient
accumulation of succinate from the relatively limited substrate supply is
scarcely possible. Besides, the inhibition of pyruvate oxidation increased
over this interval. Sometimes changes in tissue metabolism occur during
the course of an experiment independent! j^ of malonate action. When mal-
onate is added to human brain slices immediately, the endogenous respira-
tion is inhibited around 25% at 5 mM, but if malonate is added after
90 min incubation of the slices, there is no inhibition (Elliott and Suther-
land, 1952). The role of succinate oxidase in the respiration must change
as a result of the slicing or the abnormal medium.
The inhibition of the respiration of rat ventricle slices by malonate
5-20 mM follows a more complex course (Webb et al., 1949). Following
an initial inhibition, the respiration rises for approximately 1 hr and then
begins to fall again. There is thus a maximum or hump in the respiration
curve. After 1 hr, the respiratory level with malonate is higher than in
the controls. The reversal of the inhibition is inhibited by fluoride, which
would indicate that the hump is due to augmented glucose oxidation or to
the metabolism of malonate. Calcium is also necessary for the typical re-
sponse to malonate. Very complex effects of malonate were also found by
Turner and Hanly (1947) and Hanly et al. (1952) in carrot slices. The var-
182 1. MALONATE
iation of the inhibition depends on the pH. At pH 4, the inhibition de-
velops over 1 hr and remains constant, but at higher pH's the inhibition
may disappear or only stimulation may be seen. The value of these interest-
ing experiments is greatly reduced by the unaccountable use of potassium
malonate rather than the sodium salt. Potassium at 50 mM (which is the
concentration of malonate generally used by them) stimulates the respira-
tion and alters its character, so that aU the results must be the summation
of two usually opposing actions. This illustrates how the incorrect choice
of an inhibitor salt can vitiate the results of an otherwise excellent in-
vestigation.
Most of the work on the malonate inhibition of endogenous respiration
has been done without regard for possible alterations in the inhibition
with time. The inhibitions have simply been determined over an arbitrary
interval. Inasmuch as changes in the inhibition by malonate occur quite
frequently, it is likely that over all inhibitions, such as are presented in
Table 1-26, are often mean values and do not reflect either the initial inhi-
bition or the maximal inhibition. As was pointed out in Chapter 1-12, the
value of many studies on inhibitors would be increased by determinations
of the variation of the inhibitions with time.
Effects of Different Conditions on the Inhibition of Endogenous Respiration
One of the most important variables affecting the response of the endog-
enous respiration to malonate is the age of the tissue, particularly as it
relates to the stage of development or the interval between the preparation
of the tissue and the experimental testing. The respiration of plant tissues
usually becomes more sensitive to malonate with time. This indicates a
progressive change in the metabolic pattern in the direction of a greater
participation of the cycle. The changes in the inhibition during malonate
inhibition, discussed in the previous section, can be due to the effects of
the malonate or to inherent metabolic alterations. It is thus important in
such studies to determine both the changing inhibition in the presence of
malonate and the changing susceptibility as malonate is added at various
intervals. The malonate inhibition rises with time in the Avena coleoptile
(Bonner, 1948), rose petals (Siegelman et al., 1958), chicory root slices (La-
ties, 1959 a), Arum spadix slices (Simon, 1959), and potato tuber slices
(Romberger and Norton, 1961). These changes are usually associated with
an increase in the total uninhibited respiration. For example, in potato
slices there is a 4-fold rise in the respiration during incubation for 30 hr;
the malonate resistant fraction doubles and the malonate-sensitive fraction
increases 10- to 15-fold. Carrot slice inhibition by malonate, on the other
hand, decreases steadily up to 376 hr after cutting the sections (Hanly
et al., 1952), although the uninhibited respiration first rises and then falls.
The results obtained with animal tissues are less striking and more variable.
EFFECTS ON THE ENDOGENOUS RESPIRATION 183
The inhibition of rabbit ova respiration at various times postcoitum does
not change significantly (Fridhandler et al., 1957), while the inhibition of
trematode respiration decreases with time from excystment (Vernberg and
Hunter, 1960). When 20 raM malonate is added to rat ventricle slices 1 hr
after slicing, the inhibition of the respiration is only 25%, whereas initially
the inhibition is near 50% (Webb et al., 1949), and this relationship holds
for all malonate concentrations up to 100 vaM. We have seen that in the
presence of malonate the inhibition has been replaced by stimulation at
1 hr. Therefore the metabolism changes differently during the action of
malonate and the maxima in the time curves cannot be explained by
inherent alterations of the metabolic pattern. All of these results point to
the importance of considering the time factor in studies of malonate inhi-
bition.
Another apparently important factor is the ion and buffer composition of
the medium, although no thorough studies have been done and the mecha-
nisms are not understood. Many years ago Annau (1935) observed that the
inhibition of the respiration of both rabbit liver and kidney slices is less
in Ringer than in phosphate medium. The results were quite variable but
on the average the inhibition is 30% in phosphate and 15-20% in Ringer
medium. Unfortunately, Annau did not state what form of malonate was
used nor did he mention pH control, so the results are perhaps unreliable.
The presence of bicarbonate abolishes the inhibition by malonate in ox
retina homogenates (Burgess et al., 1960), and it is probable that inhibition
in intact cells would also vary with the bicarbonate concentration. Bicarbon-
ate can, of course, facilitate the formation of oxalacetate through carboxyla-
tion reactions. When Ca++, Mg++, or K"*" is removed from the medium, the
inhibition of rat brain slice respiration by 10 mM malonate is not altered,
but in the presence of fumarate the inhibition becomes progressively greater
as these ions are successively removed (Greville, 1936). The addition of
Ca+"'" to nematode minces increases both the respiratory inhibition and the
inhibition of succinate oxidation (Massey and Rogers, 1950). The sensitivity
of chicory root slice respiration to malonate is markedly affected by K+ and
Li+ (Laties, 1959 b). Slices incubated with 50 milf K+ are inhibited more
and with 50 mM Li+ inhibited less than the fresh slices. It is also interesting
that increase in CO2 tension results in progressive disappearance of the mal-
onate inhibition, whereas increase in Og tension augments the malonate-
sensitive fraction of the respiration. It is thus clear that the medium can
play an important role in the response to malonate. Much work has been
done in quite nonphysiological media and the results are thus difficult to
apply to the actions of malonate in situ. Much more effort should be di-
rected at creating approximately physiological conditions.
The functional activity of the tissue determines the level and type of
respiration, and therefore is often a major factor in the sensitivity to mal-
184 1. MALONATE
onate. This has been shown particularly clearly in brain slices, stimulated
both electrically (Heald, 1953) and by K+ (Kimura and Niwa, 1953; Yoshida
and Quastel, 1962). The stimulated respiration is readily inhibited by mal-
onate (Fig. 1-14) whereas the resting respiration is insensitive. This be-
havior is probably exhibited by many tissues. It is often very difficult to
determine exactly the functional state of isolated tissues, such as slices,
but where possible this should be attempted. We shall find later that
active tissues are more easily functionally depressed by malonate and
this may have a metabolic basis. Indoleacetate stimulates the growth of
Avena coleoptiles and increases the respiration simultaneously. This ad-
ditional respiration brought about by indoleacetate is readily inhibited by
malonate (Bonner, 1949), and it is likely that the respiration of rapidly
growing tissue is generally inhibited more strongly by malonate than that
of resting or slowly proliferating tissue.
Consideration must also be given to the history of the tissue. Bonner (1948)
has shown that the nutritional state of the Avena coleoptile determines the
inhibition by malonate, and it is probable that the same applies to animal
tissues. The inhibition of wheat seedling respiration by malonate depends
on a number of factors, including the type and duration of irradiation,
the nutrition, and the region from which the plants come (Farkas et al.,
1957 a, b). This is a field that has been very little explored. The changes
in the respiratory inhibition of animal tissues with nutrition might not only
provide information on the metabolic patterns under various conditions,
but be important in the use of the inhibitor to selectively depress the me-
tabolism and growth of neoplastic tissues.
Effects on the Respiratory Quotient
The effects of an inhibitor on the respiratory quotient (R.Q. = COg form-
ed/Og uptake) are often indicative of shifts in metabolic pathways. Let us
first consider the theoretical values of the R.Q. for the metabolism of
various substrates (see tabulation) in the presence and absence of malonate,
assuming that malonate is able to block succinate oxidation completely.
Cases in which oxalacetate is formed in the cycle and from noncycle sources
must be separated. Summarizing these results, one would expect malonate
to increase or decrease the R.Q., depending on the substrate and the nature
of the cycle operation. Since a complete block of succinate oxidation would
prevent the formation of oxalacetate through the cycle, malonate may
shift the pathway from cycle oxalacetate to externally formed oxalacetate,
if the latter reaction is possible. If this is so, the R.Q. should rise in every
case.
This prediction is quite consistently borne out experimentally. The R.Q.
of rat liver slices rises from 0.72 to 0.77 in the presence of 20 mM malonate,
at which concentration the respiration is inhibited 14% (Elliott and Greig,
EFFECTS ON THE ENDOGENOUS RESPIRATION 185
Substrate metabolism R.Q.
Glucose -> CO2 + H2O 1.0
Glucose -> succinate -f CO2 + H^O 0.8
Glucose + 2 oxalacetate ->• CO2 + HgO 1.27
Glucose + 2 oxalacetate -> 2 succinate + CO2 + H2O 1.5
Pyruvate -> COj + HjO 1.2
2 PjTuvate -> succinate + CO2 + H2O 1 . 33
Pyruvate + oxalacetate -> COj + H2O 1.4
Pyruvate + oxalacetate ->■ succinate + COj + O2 2.0
ButjTate -^ CO2 + H2O 0.8
ButjTate -r oxalacetate -> COj + HjO 1 . 23
Butyrate + oxalacetate -> succinate + CO2 + HjO 1.0
Butyrate + 2 oxalacetate -> CO2 + HjO 1.2
But>Tate + 2 oxalacetate -> 2 succinate + CO2 + H2O 1.33
1937). However, malonate decreases the R.Q. of kidney slices, both en-
dogenous and with pyruvate as the substrate. Malonate elevates the R.Q.
of rat adipose tissue from 1.0 to 1.13 endogenously and from 1.14 to 1.41
in the presence of glucose (Haugaard and Marsh, 1952). In frog muscle,
the R.Q. first rises from 0.9 to 0.97 at 10 mM malonate, but then progres-
sively decreases as the malonate concentration is raised so that at 200 mM
malonate the R.Q. is 0.39 (Thunberg, 1909). In plant tissues, the effects
are less variable. Malonate has been shown to increase the R.Q. of barley
roots from 0.97 to 1.14 (Machlis, 1944), of maize roots (Beevers, 1952),
of carrot roots up to values as high as 3 (Hanly et al., 1952), of chicory
roots from 1.03 to 1.14 (Laties, 1959 a), and of rhubarb leaves at pH 5.3
(Morrison, 1950).
Of course, there are many factors which must be taken into account,
since malonate can secondarily alter several metabolic pathways. A stimula-
tion of glucose uptake could change the R.Q. in either direction, depending
on the nature of the substrates used in the uninhibited tissue; in the pre-
sence of a significant cycle block, this would usually depress the R.Q. and
counteract the more direct effects described above. On the other hand,
metabolism of malonate would tend to elevate the R.Q. since the complete
oxidation would give R.Q.'s of 1.50-1.55 and the oxidation to succinate
3.0-4.0. A final factor of importance is the relative dependence of glucose
and fatty acid metabolism on the operation of the cycle and the levels of
ATP, since malonate could alter the oxidative contribution from these sub-
strates secondarily.
186 1. MALONATE
Significance of Respiratory Inhibition
Does the degree of malonate inhibition indicate the contribution of the
cycle to the total oxygen uptake? This must certainly be answered in the
negative. Lack of inhibition can be due to a failure to penetrate, the me-
tabolism of malonate, a source of oxalacetate external to the cycle, met-
abolic adaptations of the cells, and many other factors. Positive evidence
of inhibition is more valuable than absence of inhibition, but even when
definite inhibition is observed the possibility of actions other than in the
cycle must be considered, especially when the malonate concentration must
be high to achieve an effect. It is doubtful if anyone examining Table 1-26
would attempt to correlate the inhibitions with the importance of the
cycle in the organisms and tissues. For example, in general there is greater
inhibition of mammalian endogenous respiration than of the respiration of
microorganisms or plants. This might indicate a greater role of the cycle
in mammalian tissues, but it could also be attributed to a poorer penetration
in the plants and microorganisms, or to a greater metabolic flexibility and
adaptability in these more resistant forms. It must also be clear that the
degree of respiratory inhibition bears no necessary relationship to the de-
gree of inhibition of succinate dehydrogenase. A significant inhibition by a
reasonable concentration of malonate is evidence for the operation of the
cycle, but the quantitative aspects of the contribution cannot be derived
from these data alone. The effects of malonate on the endogenous respira-
tion are sometimes of greater physiological significance than effects on the
oxygen uptake in the presence of high concentrations of often abnormal
substrates, since the endogenous metabolism may be representative of a
more normal balance of substrates. In this connection, studies of inhibitors
would often be improved if the attempt were made to provide the cells
with a mixture of physiologically pertinent substrates at the concentrations
normally occurring in the cellular environment.
PERMEABILITY OF CELLS TO MALONATE
One of the major problems in the use of malonate has always been the
degree of penetration of the inhibitor into the cells or tissues, and it has
been frequently stated that this is the primary factor responsible for the
low inhibitions observed in many cases. It is true that the plasma mem-
brane is relatively impermeable to most ions, particularly anions and those
carrying two or more charges, but if this is so how can one explain the
marked respiratory stimulations usually seen with succinate or other di-
carboxylate ions? Furthermore, malonate is often metabolized readily by
tissues and this presupposes entrance into the cells. Since there are other
possible reasons for a resistance to malonate, the permeability hypothesis
must be examined critically.
PERMEABILITY OF CELLS TO MALONATE 187
Experimental Evidence Relating to the Penetration of Malonate
Malonic acid is more than 10 times as lethal on injection into frogs as
is sodium malonate (Heymans, 1889). No explanation was offered for this
observation but it could have been due to the greater permeability of the
cells to the acid or, on the other hand, to a nonspecific acidification of the
animals. Malonate administered to rabbits circulates initially in a volume
equivalent to the extracellular compartment and the intracellular transfer
occurs slowly (Wick et al., 1956). The failure of malonate to alter the me-
tabolism of labeled acetate was attributed to both the slow penetration into
the tissues and the simultaneous metabolism of the malonate, both factors
keeping the intracellular concentration at low levels. The inability of mal-
onate to alter gastric acid secretion in frogs, even at lethal doses, was sim-
ilarly attributed to these factors (Davenport and Chavre, 1956). Inasmuch
as succinate oxidase is present in the secretory cells and the cycle is im-
portant in secretion (as shown by the inhibition with fluoroacetate), the
lack of action must be due to an insufficient concentration within the cells.
Turning to isolated tissues and cell suspensions, the augmentation of
malonate effects by procedures designed to reduce or abolish the perme-
ability barriers has been demonstrated many times (Table 1-26). Rat dia-
phragm respiration is inhibited slowly by malonate, but if the diaphragm
is cut into small pieces, and hence presumably damaged, the inhibition is
immediate (Greville, 1936). The oxygen uptake of pigeon brain brei is
inhibited rather poorly by 24 mM malonate, but when the brain is dispersed
more completely in the form of a homogenate the inhibition is more marked
(Banga et al., 1939). The succinate dehydrogenase of yeast incubated for
5 hr in liquid nitrogen is much more susceptible to malonate than in normal
cells (Lynen, 1943), and the same holds for E. coli treated with toluene
(Ajl and Werkman, 1948). Sensitivity to malonate can be induced by liquid
nitrogen treatment in the fungus Zygorrhynchus (Moses, 1955) and by dry-
ing Pseudomonas (Gray, 1952). Malonate does not inhibit glutamate oxida-
tion in intact cells of Pasteurella, but inhibits well in sonic lysates (Kann
and Mills, 1955). All of these phenomena have been interpreted in terms of
permeability. This is certainly the most obvious explanation and it is prob-
ably generally correct, but it must be admitted that such drastic treat-
ments could affect many other things; for example, alter the organized
enzyme structure so that the attacked enzyme is more exposed, or reduce
the ability of the cells to metabolize malonate.
Only one investigation of the relative permeabilities of the dicarboxy-
late anions has been made. Giebel and Passow (1960) determined the half-
times for penetration of these ions into bovine erythrocytes and the results
are given in Table 1-27. Giebel and Passow attempted to correlate the per-
meabilities with the ionic sizes and the acidic ionization constants. The
ionic volumes and lengths presented in the table, which are somewhat
188
1. MALONATE
Table 1-27
Erythrocyte Permeabilities and Molecular Properties of Dicarboxylate
Anions "
Anion
Relative
permeability
Ionic
volume
Ionic
length
(A)
(H,B)
(HB-)
Oxalate
100
54
5.1
0.00015
276
Malonate
17
66
7.5
0.112
6590
Maleate
14
75
7.9
0.051
252
Fumarate
1.4
75
7.6
0.014
445
Succinate
0.45
78
7.5
2.56
6410
Malate
0.31
91
7.5
0.135
1860
Glutarate
0.085
90
8.5
1.99
3780
Tartrate
0.045
104
7.5
0.0103
338
" The permeabilities were determined in bovine erythrocytes by Giebel and Passow
(1960). The values given here for the relative permeabiHties are the reciprocals of the
entrance rate half-times (ti/^) multipHed by 100. Tlie calculations of the ionic volumes
and lengths are approximate and usually depend on the configuration of the ions.
The concentrations of HjB and HB are given for a total concentration of 1 M.
different than those given by Giebel and Passow, must be considered as
only relative values, neglecting hydration and special configurations of the
ions. Their experiments were run at a pH of 7.35 so the concentrations of
the undissociated and singly dissociated forms of the acids are given for
this pH. The ionization constants are sometimes quite different from those
assumed by Giebel and Passow and are those in Table 1-2. There is certainly
little or no general correlation between permeabilities and the concentra-
tions of either H2B or HB". Succinate, for example, penetrates one-thirty
eighth as rapidly as malonate and yet (H2B) for succinate is 23 times higher
than for malonate. This does not necessarily invalidate the assumption
that for a single substance the unionized forms penetrate more rapidly than
the ionized, but it shows that there are other factors which are quite im-
portant. There is also no correlation with the ionic length and it is unlikely
that one would be expected. However, there is some correlation with ionic
volume, leading Giebel and Passow to suggest that the dicarboxylate anions
penetrate through pores in the membrane, whereas the monocarboxylates
pass through the lipid phase of the membrane. They calculate the pore
radius to be between 3.8 and 4.5 A. If these ions pass through the pore
channels in the extended form, which is likely, there are two major factors
which may contribute to the permeability: the cross-sectional area perpen-
PERMEABILITY OF CELLS TO MALONATE 189
dicular to the direction of passage and the degree of interaction of the
molecules with the walls of the pores. This interaction may be of various
types and includes steric repulsion and attractive forces (such as van der
Waals' forces or hydrogen bonds). The configuration of the ion must be
important in this connection. Maleate penetrates 10 times faster than fu-
marate and this must be mainly due to the structure of maleate wherein
the carboxylate groups are much closer than in fumarate. That both of
these ions penetrate more rapidly than succinate may be due to the greater
rigidity of the former, energy perhaps being required for succinate to change
from its statistically most probable configuration to that necessary for
penetration. It is evident, however, that none of these explanations satis-
factorily fits the experimental data and that we need to know much more
about the membrane before accurate interpretations can be made. It should
be mentioned that these results on erythrocytes do not apply to other
types of cells or tissues, inasmuch as erythrocyte permeability is in some
senses unique.
Malonate inhibits the succinate dehydrogenase of calf thymus nuclei and
yet at 10 mM has no effect on the respiration or ATP level of intact nuclei
(McEwen et al., 1963 b). This indicated a failure to penetrate and it was
shown with labeled malonate that this is indeed the case, which is some-
what surprising in view of the usual concepts of the nuclear membrane.
Malonic acid does not enter organic solvents from water readily, due
probably to the dipolar nature of the unionized carboxyl groups. The par-
tition ratios for malonic acid are given as (concentration in solvent/concen-
tration in water): oleyl alcohol 0.049 (Collander, 1951), ether 0.083, iso-
butanol 0.62, methylisobutylketone 0.15, and methylisobutylcarbinol 0.37
(Pearson and Levine, 1952). The partition ratios for succinic acid are always
somewhat higher, as expected. These data would indicate that even the
unionized malonic acid would not penetrate through the lipid phase of the
membrane too readily. The fact that the un-ionized forms penetrate better
than the ionized does not imply that passage through a lipid phase occurs.
The negative charge on the ions could impede movement through pores,
especially when it is considered that in most cells these ions must move
up an electrical potential gradient to cross the membrane.
Variation of Malonate Inhibition with pH
One of the strongest arguments for the preferential uptake of the less
ionized forms of malonic acid is the rise in the inhibition observed with a
lowering of the pH. This has been examined particularly in plant tissues
and the results are quite uniform. Such effects have been observed in to-
mato stem slices (Link et al., 1952), maize roots (Beevers, 1952), rhubarb
leaves (Morrison, 1950), barley roots (Laties, 1949 a), spinach leaves (Bon-
ner and Wildman, 1946), carrot root slices (Hanly et al, 1952), and Avena
190 1. MALONATE
coleoptile (Cooil, 1952). These results are plotted in Fig. 1-14-19. On the
other hand, rather insignificant effects of pH have been noted in fungi,
such as Microsporum, Trichophyton, Epidermophyton (Chattaway et al.,
1956), and Pullularia (Clark and Wallace, 1958). In Pullularia malonate
is readily oxidized; the oxygen uptake from malonate increases with a
lowering of the pH along with the inhibition of the cycle and the effects
tend to cancel one another. The effects of malonate in tobacco leaves are
not changed greatly by lowering the pH from 7 to 4 (Vickery and Palmer,
1957), although down to pH 5 the uptake of malonate becomes progressively
greater. The incubation here was very long (48 hr) and hence there was
more opportunity for malonate to penetrate than in shorter experiments.
It is unfortunate that no quantitative work on pH has been done with
animal tissues.
It is also regrettable that in those instances in which malonate inhibits
more strongly at low pH values the reversibility of the inhibition has not
been adequately tested. Lowering the pH of the medium in the presence
of a weak acid increases the amount of unionized acid entering the cells
and can decrease the intracellular pH to a degree causing cell damage. It
was noted in carrot root tissue that injury to the cells occurred, including
loss of turgor and release of some of the cell contents, with malonate at a
pH around 4 (Hanly et al., 1952). It is very difficult in experiments of this
type to distinguish between a specific malonate effect and a nonspecific
acid damage. Examination of the reversibility of the inhibitions might
provide some evidence on this point.
In Volume I it was shown that the intracellular concentration of a di-
carboxylate anion is related to the total external concentration in the follow-
ing manner (Eq. I-i4-178):
(i=), = I J; (!,)„ (1-4)
oJi \n. )'i
where the subscripts o and i refer to outside and inside the cells and ^f-'
is the appropriate pH function for the external inhibitor (see Eq. 1-14-12).
Two assumptions are involved in this formulation: (1) only the Hgl form
of the inhibitor penetrates, and (2) the cells are internally completely buf-
fered. The variation of the intracellular active 1= form with external pH
is very marked, as shown in the accompanying tabulation, assuming an
intracellular pH, of 6.8. Of course, cells are not completely buffered and,
as the internal pH, drops, the entrance of the inhibitor is slowed, so that
with decrease in the buffering capacity the ratios given will be lessened.
Here one is presented with the dilemma that at low pH values one must
either assume a high internal inhibitor concentration or a significant fall
in pH,. Since the inhibitions observed are not as high as would be predicted
on the basis of the above equation and tabulation, one is forced to conclude
PERMEABILITY OF CELLS TO MALONATE 191
that the intracellular pH^ must fall. This may not only damage the met-
abolic systems but wiU tend to reduce the inhibition on succinate dehydro-
genase by decreasing the concentration of dicarboxylate anion.
pHo
a-)i/(it)o
8.2
0.0016
7.4
0.063
7.0
0.39
6.0
34.8
5.0
1610
If one assumes that the HI~ form can also penetrate, the internal con-
centration of the active 1= form will not be so strongly dependent on the
external pH^, and the inhibition will increase significantly as the pH^ is
lowered from 6 to 5, but otherwise the same behavior will be expected. It
is usually dif&cult to distinguish between penetration by the Hgl form only
and penetration by the HI" form also. Some arguments have arisen on this
point, Bonner and Wildman (1946) believing that the HI~ form penetrates
and Beevers (1952) holding that only the Hgl penetrates. A Simon-Beevers
plot of the data from maize roots (see Chapter 1-14) shows that (Hgl) does
not remain constant for 50% inhibition of respiration over the pH range
3-6.5, which would indicate a possible contribution from the HI~ form.
Actually, it might be better to express the total entry rate of malonate as:
Entry rate = Phj(H2I)o + Phi-(HI-)o + Pi=(I=)<, (1-5)
where the P's represent the permeabilities to the various forms of the inhi-
bitor. Although Pjj I > Pri- > Pi-^ above pH 4 (HI~) is much greater
than (Hoi) so that the contribution of the second term to the total rate may
be significant. One would like to know the relative values of the P's for a
particular tissue and these could be determined if the entry rate of mal-
onate were determined (for example, with labeled malonate) at different
pH values. One must also bear in mind that a change of pH could alter
the permeability properties of the membrane.
The only data on the effects of pH on malonate action in animal tissues
were obtained on rat ventricle strips by Masuoka et at. (1952). Malonate
stimulates the amplitude of hypodynamic strips much more at pH 6.2
than at 7.4. This positive inotropic action may be unrelated to the inhi-
bition of succinate dehydrogenase, since succinate at pH 6.2 gives essenti-
ally the same response, and could depend on the oxidation of malonate
(see page 216). Whatever the mechanism, these results indicate that mal-
onate penetrates more readily at the lower pH.
192 1. MALONATE
In comparing the actions and penetrations of malonate with succinate,
or with other dicarboxylate anions, it is necessary to consider that the
relative concentrations of the ionic species can be quite different. Thus
the HgB form of succinate is at a much higher concentration than the same
form of malonate at the same total concentrations (Table 1-3). It is also
possible that some cells possess active transport or carrier systems for the
substrate dicarboxylate anions, allowing succinate to penetrate more readi-
ly than malonate. Permeability in some cases seems to be as specific as
are enzyme reactions, and can be quite dependent on the configurations
and charge distributions of the transported substances. This could be due
to either the structure of the membrane pores or the nature of a carrier.
For these reasons one must anticipate striking differences between different
tissues with regard to the relative permeabilities to the various ions, and
it is particularly important not to apply the results on plants unreservedly
to animal tissues or microorganisms.
GROWTH, DEVELOPMENT, AND DIFFERENTIATION
The responses of growth, cleavage, and histogenesis to inhibitors are
interesting because they often demonstrate the nature of the metabolic
basis for these important biological processes. The results may also have
bearing on the possible use of the inhibitors for the selective depression
of the growth of organisms or abnormal cells which are detrimental to the
host. These processes all require energy from the metabolism so that any
reduction of either the exergonic reactions or their coupled phosphoryla-
tions would be expected to interfere in some manner. In addition, more
specific effects may occasionally be observed. The selective inhibition of
the growth of certain cells can result from different rates of growth of the
cells involved, or from differences in the metabolic requirements for growth.
It is generally true that rapidly proliferating cells are more readily affected
by inhibitors than are the same or other cells growing or multiplying at
slower rates. It has also been demonstrated that various types of cells may
utilize different enzymes or metabolic pathways to support proliferation.
With respect to malonate, one might anticipate that cells whose growth
is in one way or another significantly dependent on the cycle would be
inhibited more than cells not requiring the operation of the cycle. However,
other factors, such as the degree of penetration of the malonate or the
susceptibility of the succinate dehydrogenases to malonate, may be im-
portant.
Virus Multiplication
Malonate is able in some instances to suppress the intracellular formation
of virus without permanently damaging the host cells. The results obtained
GROWTH, DEVELOPMENT, AND DIFFERENTIATION
193
with influenza type A virus isolated from man and cultured in chick chorio-
allantoic membrane illustrate this well. Ackermann (1951) found that mal-
onate inhibits the formation of virus and simultaneously reduces the oxygen
uptake of the host cells (see accompanying tabulation). Malonate is not
Malonate
% Inhibition
Final
Infectivity
(mi/)
of respiration
virus titer
titer
None
213
107.8
20
3
192
107.8
40
18
69
106.5
60
47
0
103.3
virucidal since the original virus can be recovered from the infected cul-
tures. The major effect of malonate is not to prevent infection of the cells,
since essentially the same results were obtained when malonate was added
4 hr after the inoculation of the virus. The inhibition is thus exerted on
the synthesis of new virus material. Furthermore, the host cells are not
damaged; if infected chorioallantoic tissue is exposed to 60 mM malonate
for 24 hr, virus proliferation is completely inhibited but, if the tissue is
washed free of malonate, becomes susceptible to infection and supports
virus multiplication. These results were confirmed and extended by Eaton
(1952), who reported that 42 niM malonate completely inhibits virus mul-
tiplication, inhibits respiration around 50%, and does not alter aerobic
glycolysis. The formation of virus in minced chorioallantoic membrane is
inhibited 85% by 27 mM malonate when glucose is the substrate and 95%
when glutamate is the substrate (Eaton and Scala, 1957). The injection
of 1 mg sodium malonate into chick embryos 1 hr after inoculation with
virus reduces the amount of virus after 48 hr by 55% (Hannoun, 1952).
The energy for the synthesis of new virus material must come mainly from
the host cell tricarboxylate cycle. It is impossible to reduce the energy
supply for virus synthesis without simultaneously restricting the energy
for host cell activities. However, it appears that the virus propagation is
selectively depressed because it is the most endergonic process occurring,
the chorioallantoic cells otherwise not being very functionally active. As
long as the low maintenance energy requirement is satisfied, the cells are
not damaged (see Fig. 1-9-6). If one were dealing with virus in an active
tissue, such as nerve or muscle, it would be much more difficult, or im-
possible, to selectively block virus multiplication.
Other viruses growing in animal cells have been studied with various
results. Malonate at 6.7 mM has a definite depressant effect on the proli-
feration of vaccinia virus in chick embryo cultures (R. L. Thompson, 1947)
194 1. MALONATE
and similar results were reported for psittacosis virus (Morgan, 1954). In
neither case does malonate depress the growth of the tissue cultures, noi
does it inhibit the cellular contractions of chick heart cultures. Very slight
effects of malonate at concentrations from 10 to 100 mM were observed
with foot-and-mouth disease virus in bovine kidney culture cells (Polatnick
and Bachrach, 1960). Since there is little inhibition of the respiration, it is
possible that malonate does not penetrate adequately. At 100 mM, respira-
tion is inhibited 23%, virus yield perhaps 20%, and there is a 10 min delay
in the appearance of virus. Finally, malonate was found to actually stimu-
late feline pneumonitis virus proliferation in the isolated chick embryo yolk
sac, 10 mM increasing the virus titer about 33% (Moulder et al., 1953).
Thus a wide variety of effects have been observed with different viruses
and host cells, no general conclusions being possible at this time. I am not
aware of any studies on the effects of malonate on the course of virus in-
fections in whole animals.
Plant viruses have been inadequately investigated and results on the
tobacco mosaic virus only are available. Although malonate at 0.5 mM de-
creases the number of lesions/cm^ in detached tobacco leaves from 28.9
to 21.9, Chiba et al. (1953) felt that this result is statistically insignificant.
Ryzhkov and Marchenko (1954, 1955) reported that malonate inhibits
multiplication of this virus and that this is reversed by fumarate, but
Schlegel (1957) found only variable effects of 3 mM malonate on the yield
of virus in leaf discs. The spraying of 10 mM malonic acid solutions (pH
2.7-3.6) onto the leaves of bean plants decreases the number of virus lesions
68% without leaf damage (Matthews and Proctor, 1956), but this may be
unrelated to the action of malonate on the cycle inasmuch as succinic acid
is even more inhibitory. This is probably a nonspecific acid effect because
the cycle intermediates usually increase the virus yield when they are added
to leaf cultures.
It is somewhat surprising that malonate has no demonstrable effect on
the multiplication of E. coli phage. Spizizen (1943) found no effect at 10 mM
under any conditions of virus growth, and Czekalowski (1952) reported no
actions on either T2 phage or host cells at concentrations from 0.1 to 100 mM.
It may well be that phage proliferation is not directly dependent on the
energy derived from the cycle, but inhibition by 2,4-dinitrophenol, cyanide,
and fluoride is observed. In fact, Czekalowski stated that all the inhibitors
that depress phage selectively seem to act in some manner on the cyto-
chrome system and are able to inhibit succinate oxidase; yet the most
specific inhibitor for this enzyme is inactive. Lack of penetration is an
unlikely explanation and this failure of malonate to inhibit deserves further
study.
GROWTH, DEVELOPMENT, AND DIFFERENTIATION 195
Multiplication of Bacteria, Fungi, and Other Microorganisms
Bacterial growth is apparently fairly resistant to malonate, despite the
many observations of enzyme and metabolic inhibitions in these organisms.
The absence of any effect on E. coli at malonate concentrations up to 100 mM
reported by Czekalowski was mentioned above, but Loveless et al. (1954)
found 50% inhibition of growth at 19.2 ml/ malonate, with no effect on cell
size. Malonate at 300 roM produces somewhat elongated E. coli cells and
at 800 mM they are markedly lengthened and often U-shaped: division is
abolished but the cells continue to elongate slowly (Schweisfurth and
Schwartz, 1959). The effects of malonate must depend on many factors,
including the nutrient medium, the duration of the growth phase studied,
and the pH. Although Rosenberg (1948) found the growth of Clostridium
saccharobutyricum to be inhibited by malonate, a concentration of 100 mM
was used, so that the mechanism of the effect is not clear. His observation
that the inhibition is overcome by meso-inositol and borate must be inter-
preted as indicating a unique approach to the study of malonate inhibition.
Malonate at 10 mM has a slightly depressant action on rate of germination
of Bacillus subtilis spores but does not affect growth in culture (Hachisuka
et al., 1955). The bacteriostatic concentrations of malonate were given as
3.2 mM for Pseudomonas fluorescens and 7.7 mM for B. aerogenes (Cooper
and Goddard, 1957) but the acid was used, the pH being 2.5 and 1.5, re-
spectively, so that a nonspecific acid effect is the most likely explanation,
especially as succinic acid is as inhibitory. It is clear that the investigations
of the effects of malonate on bacterial growth leave everything to be desired.
The sporulation and growth of yeast are inhibited by malonate quite
potently and in parallel fashion, 50% depression of each occurring at 5 mM
(Miller and Halpern, 1956). On the other hand, Hensenula ellipsoidospora,
a vellum-forming yeast, grows more rapidly in the presence of malonate
(Luteraan, 1953). It would certainly not be too surprising to find certain
microorganisms stimulated by malonate inasmuch as many can oxidize
malonate readily (page 228). Malonate arrests sporulation of Aspergillus
niger without suppressing growth (Behal, 1959) but germination of Neuro-
spora ascospores is not blocked by 10 mM malonate even at pH 2.3 and
after 24 hr (Sussman et al., 1958), nor is the germination of Puccinia uredo-
spores affected significantly by 20 mM malonate at pH 4.8, although the
respiration is inhibited around 37% (Farkas and Ledingham, 1959). The
formation of conidia is often essential for the spread of the scab disease
of apple caused by Venturia inaequalis and so Kirkham and Flood (1963)
investigated the effects of various respiratory inhibitors on ascosporulation.
Malonate was found to inhibit at high concentrations, the inhibition sur-
prisingly increasing with increase in the pH (see accompanying tabulation).
This might imply an action on or within the membrane; this is supported
by the relative lack of effect on the respiration and the rather potent inhi-
196 1. MALONATE
bition produced by ^ra ws-aconitate. The injection of 50 milf malonate into
the leaf petioles, however, increases infectivity so that a more significant
Malonate
(mil/)
Initial pH
0/
/o
Inhibition of
sporulation
50
4.2
4
100
39
40
5.0
Stim 7
100
71
50
6.2
68
100
96
effect on the host tissue is evident. The development of Puccinia rust on
wheat seedling leaves is inhibited by 10 mM malonate, but the leaf tissue
is damaged (Samborski and Forsyth, 1960). In this particular case the
phytotoxicity is greater than the rust suppression so that malonate could
not be used commercially. Malonate esters have been tested for inhibition
of mold growth in syrups, but 13 mM does not have much effect over 144 hr
(Lord and Husa, 1954); however, these esters are used as fungistatic agents
in soy sauce in Japan (Tsukamoto, 1951). Another instance of growth stim-
ulation by malonate was reported for Endamoeba histolytica (Nakamura
and Baker, 1956); the average cell count per field at the end of 3-4 days
was 1 in the control and 16 in the presence of 12.8 mM malonate, indicating
possible metabolism of the malonate by these organisms.
Plant Growth
Avena coleoptile growth is sometimes stimulated and sometimes depressed
by malonate, the response depending on the strain of oats used, the pH,
and whether the sodium or potassium salt of malonate is used. The marked
inhibition (61%) reported by Commoner and Thimann (1941) for 10 mM po-
tassium malonate over 24 hr has not been observed by others. Albaum and
Eichel (1943) found only stimulation (around 30%) from 1-5 niM potassium
malonate over a period of 160 hr, and it was felt that malonate was serving
as a substrate, which was substantiated by the higher respiration in the
presence of malonate. Thimann and Bonner (1948) provided further evi-
dence for this by the finding that malonate at 1 mM, having little effect by
itself, antagonizes the marked inhibition produced by iodoacetate. How
much of this is due to malonate and how much to potassium is difficult
to say. Cooil (1952) confirmed the counteraction of iodoacetate inhibition
by potassium malonate, but found that the sodium salt is not nearly as
potent, implicating the potassium ion. The failure of malonate to inhibit
the growth is probably the result of poor penetration, as shown by the
GROWTH, DEVELOPMENT, AND DIFFERENTIATION 197
effects of pH (see accompanying tabulation); the pH alone has little effect
on growth. The malonate inhibition is satisfactorily reversed by fumarate.
pH Mean growth (mm) in malonate 3 mM
6.5 1.56
6.0 1.70
5.5 1.55
5.0 0.22
4.5 0.16
The mitotic activity of the excised roots of the garden pea {Pisum sativum)
is stimulated by glucose. This is very strongly blocked by malonate at pH
5.5. A concentration of 0.01 mM delays the action of glucose 2 hr but does
not inhibit mitosis; 0.1 mM inhibits mitotic activity around 50%; 0.5 milf
almost completely inhibits mi^-^ses; and 1 mi!f not only inhibits completely
but produces some toxic effects (Wilson et al., 1959). It was postulated
that the initiation of mitosis is dependent on the cycle, since once mitosis
began it proceeded to telophase normally. The progression through mitosis
may be coupled with a more anaerobic type of metabolism. The growth
and cell proliferation in tissue cultures of the crown galls of various plants
(marigold, Paris daisy, periwinkle, and sunflower) are inhibited to different
degrees by malonate. Cultures from normal tobacco stem are inhibited
similarly. At 10 mM, the following inhibitions may be estimated from the
curves given by Hildebrandt et al. (1954): sunflower 0%, marigold 29%,
tobacco 40%, Paris daisy 60%, and periwinkle 67%. At 80 mM malonate,
all are inhibited completely. A question arises again as to whether these
effects are related to cycle inhibition, since succinate, pyruvate, acetate,
and other organic anions inhibit also. The pH was 6.0 so that acid effects
should not be important.
Egg Cleavage and Embryogenesis
The best and most interesting work on the growth responses to malonate
has been done with marine invertebrate eggs and embryos. Since this work
was done in sea water, we must bear in mind that the concentration of
free malonate is much less than the total concentration due to the high
amounts of Ca++ and Mg+"'". When malonate is added at a total concen-
tration of 25 mM, it is likely that the free malonate is around 4 mM (see
Table 1-5). In addition, the pH of sea water is near 8.2 and this is unfavor-
able to malonate penetration into the cells. Considering these factors, it is
surprising that such definite and characteristic effects of malonate have
been observed.
198 1. MALONATE
Egg cleavage is generally rather sensitive to malonate. Although Arbacia
eggs divide normally in 1 mM malonate (Krahl and Clowes, 1940), Dend-
raster eggs are inhibited quite well at this concentration (Pease, 1941). The
development of bilaterality in Dendraster, seen with many inhibitors, does
not occur with malonate even at 10 mM, demonstrating a true differential
effect on cleavage. Division of Arbacia and Chaetopterus eggs is inhibited
completely by malonate at 70 mM, although 40 mM is essentially without
action, and this is completely reversed by fumarate, indicating a block of
the cycle (Brust and Barnett, 1952; Barnett, 1953). This is not a sodium
effect since NaCl does not inhibit. Such high concentrations of malonate
are not unreasonable in work in sea water and the inhibitions here may be
quite specific. Egg cleavage seems to depend on the ATP generated in the
cycle, because concentrations of malonate that completely inhibit the cleav-
age of Chaetopterus and Lytechinus eggs cause an immediate drop in the
high-energy phosphate to the unfertilized levels, and inorganic phosphate
is actually lost from the cells (Barnett and Downey, 1955).
The effects of malonate on the development of Arbacia eggs were
thoroughly investigated by Rulon (1948), who demonstrated abnormal dif-
ferentiation with low concentrations of malonate. If fertilized eggs in the
1-2-cell stage are placed in 1.2 mM malonate, a slight retardation of cleav-
age is observed, and at 13 hr (when the controls are swimming bilateral
gastrulae) there is no evidence of gastrulation, development having pro-
gressed only to spherical blastulae with no ventral flattening. After 24 hr
(when the controls are plutei), abnormal gastrulae with thickened apical
ends and long active cilia are seen. At 48 hr some had developed into ab-
normal plutei with ciliated apical knobs rather than normal arms. When
malonate is removed after 13 hr, quite normal plutei are formed, showing
that the effect is readily reversible. It is interesting that eggs exposed to
malonate for varying times, then washed free of malonate and fertilized,
show abnormal development, demonstrating that malonate can so disturb
egg metabolism that the effects are made evident later after the malonate
is no longer present. Exposure of unfertilized eggs to 1.44 mM malonate
for 12 hr, for example, leads to only 30 40% cleavage with irregular cleav-
age furrows and cells of unequal sizes, development not progressing
beyond irregular blastulae. Rulon postulated a gradient of succinate de-
hydrogenase throughout the cells and embryos, paralleling the physiological
activity gradient, but it is not necessary to assume this to explain effects on
differentiation. In connection with our work on parapyruvate (Montgomery
and Bamberger, 1955), we examined the development of Strongylocentrotus
purpuratus in 25 mM malonate. Up to 24 hr no discernible differences from
the controls are seen, but at 44 hr a slight inhibition of development can be
detected, with less formation of the primary mesoderm. Incoordination of
ciliary activity is also evident, most of the blastulae simply rotating rather
GROWTH, DEVELOPMENT, AND DIFFERENTIATION 199
than swimming. At 64 hr, when the controls are beginning to gastrulate,
the treated embryos are still spherical blastulae, and at 72 hr have not
progressed beyond this point. In this species, malonate would appear to
be a rather specific inhibitor of gastrulation without altering cleavage pri-
marily. It may be noted that other cycle inhibitors, such as parapyruvate
and fluoroacetate, also specifically block gastrulation in Strongylocentrotus.
It is difficult to understand these marked differences between the behaviors
of various sea urchin eggs, and especially the striking effects obtained by
Rulon with such low malonate concentrations. The free malonate concen-
trations in his work would have been around 0.17 milf (he used Ca++-free
sea water) and the intracellular concentration presumably much less.
There is evidence that insect spermatogenesis is an aerobic process with
the terminal electron transport through the cytochrome system, and that
the cycle is the primary pathway for substrate oxidations. Yet no inhi-
bition of meiosis or differentiation into spermatids and spermatozoa by
50 ToM malonate is observed in hanging-drop cultures from the Cecropia
silkworm (Schneiderman et at., 1953). These experiments were carried out
at pH 6.8-7.2 and it is possible that malonate failed to penetrate.
Amphibian gastrulation is inhibited by high concentrations of malonate.
Frog embryos at the early dorsal lip stage were dissected to give explants
which were exposed to 40 raM malonate at pH 8.0 for 18 hr. Development
does not proceed beyond the next stage (Ornstein and Gregg, 1952; Gregg
and Ornstein, 1953). There is no differential effect on the respiration of
dorsal and ventral explants, both being inhibited about 60%. Unfortunately,
there is some doubt that the block of development is due to any specific
effect of the malonate since 45 vciM NaCl apparently inhibits to the same
degree. Thus the mechanism of the block could have been osmotic or a Na"*"
effect. The chick embryo seems to be much more sensitive to malonate.
Explants of chick embryo in the presence of glucose undergo morphogenesis
and differentiation to the formation of the central nervous system and an
actively beating heart. Malonate at 1-2 mJf exerts striking differential ef-
fects (Spratt, 1950). Although no differences were noted during the first
20 hr, afterwards the central nervous system degenerates completely while
the heart forms normally and beats. Malonate is the most specific inhibitor
for the development of the nervous system. Some antagonism of this effect
was seen with succinate but none with fumarate. No inhibition of mitoses
in cultures of chick bone is observed with malonate from 65 to 138 nxM
(A. F. W. Hughes, 1950).
Mitoses in Mammalian Tissues
Epidermal mitotic activity in mouse ear fragments was determined over
4 hr periods at pH 7.4 and 38^ by BuUough and Johnson (1951). From the
effects of anaerobiosis and various substrates it was concluded that the
200 1. MALONATE
energy for mitosis is derived from cycle oxidations. Malonate at 20 mM
prevents the cell from entering mitosis for 3 hr but evidence of recovery is
seen after this time. However, at 4 hr, although some cells are progressing
through mitosis, the number of mitoses is definitely less than in the controls.
There is no evidence that malonate can stop mitosis once it has begun.
Epidermal cells adjacent to the wound show a higher number of mitoses
than normal epidermis and malonate depresses both strongly (see accom-
panying tabulation). (Bullough and Laurence, 1957). Malonate can thus
Malonate
Average number of mitoses
(milf) Epidermis adjacent to wound Normal epidermis
0 8.20 0.96
10 1.23 0.22
20 0.05 0.18
30 0.05 0.01
inhibit healing without significant damage to the tissue, since no necrosis
is seen following incubation with malonate. Similar results were obtained
by Gelfant (1960) and concentrations as low as 0.5 mM are definitely inhi-
bitory. After 4 hr 50 m M malonate produces some necrosis. The mitotic
rate in the germinal epithelium of rat ovaries is also suppressed by malo-
nate, 2 mM having variable effects but inhibiting 21% on the average and
10 mi/ inhibiting 59% (Weaver, 1959). The relatively high sensitivity of
mammalian tissue mitosis to malonate would implicate the cycle as an
important source of energy for this process.
Growth of Neoplastic Tissues
The respiration of various types of tumor cells is inhibited by malonate
(Table 1-26) but comparisons with the appropriate normal tissues have
seldom been made. Amino acid uptake is also inhibited (Kit and Greenberg,
1951) and high-energy phosphate compounds reduced (Greaser and Schole-
field, 1960), but whether tumor tissue is more or less sensitive than normal
tissue to malonate is not known. Fishgold (1957) obtained evidence that
hepatoma succinate oxidase is inhibited more readily than the enzyme from
normal mouse liver, but it is not certain if this is a true difference in the
affinities of the dehydrogenase active center for malonate or the result of
other factors. Other than this, there is no demonstrated metabolic difference
between tumor and normal tissues with respect to inhibition by malonate.
The time course of the accumulation of succinate in the Flexner-Jobling
tumor is essentially the same as in other tissues (Fig. 1-11), but the relation-
GROWTH, DEVELOPMENT, AND DIFFERENTIATION
201
ship of the accumulation to malonate concentration is not linear for the
tumor (Fig. 1-12). As pointed out by Busch and Potter (1952 b), the ac-
cumulation of succinate in the tumor may be superficially indistinguishable
from that in normal tissues, but there is reason to believe that the succinate
in the tumor arises by somewhat different pathways, mainly from gluta-
mate and related compounds. There is little reason to believe from the
known metabolic characteristics of tumor tissues that malonate would se-
lectively depress their growth; indeed, one might expect them to be less
sensitive to malonate, except for the fact that tumor cells are often more
rapidly proliferating and more active metabolically than normal tissues.
Neoplastic growth in general seems to be relatively resistant to malonate.
Malonate at 30 mM is not toxic to cultures of various tumors but 50 mM
is toxic to all types of cells in a few hours (Chambers et al., 1943). It is
interesting that no differences in susceptibility of lymphocytes and other
wandering cells, whether from normal or neoplastic tissues, are seen. Eagle's
KB strain of human carcinoma cells is more sensitive, the 50% inhibitory
concentration of malonate being around 4 ml/ (Smith et al., 1959). Malo-
nate, like many metabolic inhibitors, is capable of producing acentric blebs
on Sarcoma 37 ascites cells (Belkin and Hardy, 1961). Although this indi-
cates some alteration of the membrane properties and the permeability to
water, the relationship to growth inhibition is unknown.
Studies of the action of malonate on tumors growing in whole animals
are more pertinent to the problem of the possible value of this inhibitor
in therapy. The earliest work was done by Boyland (1940) following his
investigations of the effects of malonate on tumor respiration. Definite sup-
pression of growth was observed with malonate at a dose well below the
lethal, the carcinoma being more sensitive than the sarcoma (see accom-
panying tabulation). Actual regression of the carcinomata was observed.
Inhibitor
Dose
% Inhibition of growth of
LD,„
(mg/day) Qrafted sarcomata Spontaneous carcinomata
Malonate
20
32
Malonamide
25
36
Ethylmalonate
40
27
Glutarate
25
8
Adipate
25
0
31
100
44
150
25
160
48
150
35
200
as indicated by the 131% inhibition. Whether ethylmalonate and malon-
amide are metabolized to malonate and are active for this reason is not
known, but the lesser potencies compared to malonate do not suggest that
these uncharged substances penetrate better. Malonate, fluoride, iodoacetate
202 1. MALONATE
and azide were administered to patients with advanced neoplasia and tem-
porary suppressive effects were noted (Black and Kleiner, 1947; Black et at.,
1947). Sodium malonate was given orally at doses of 1-1.5 g/day. It is
difficult to state clearly the effects of malonate, since the inhibitors were
usually given sequentially or together, but it was stated that hematological
remissions occurred in acute myeloblastic leukemia and that shrinkage of
solid tumors, with relief of pain, was evident. The tumor cells usually be-
come refractory to these inhibitors. After resistance to fluoride and iodo-
acetate has developed, a beneficial effect is seen with malonate. It would
seem that these results are encouraging enough to warrant further study,
particularly with combinations of the inhibitors to prevent or reduce the
development of resistance. Several derivatives of malonate were tested
against mammary Carcinoma 755 in mice and suppressive action was dem-
onstrated (Freedlander et al., 1956). Malonic acid at 1.2% in the diet did
not affect either the tumor size or the growth of the mice. The most active
ethyl ester was diethylethoxymethylenemalonate (the group =CH — 0 —
— CH2CH3 on C-2), which at 1.2% in the diet reduces the surface area of
the tumors 83% while causing minimal loss of body weight. This substance
is as effective as 8-azaguanine and is less depressant on the total body
growth. Some diamides are also active, iV-dimethylmalondiamide being the
most active, reducing tumor area 68% with no effects on body growth.
It was thought that these substances may be inhibitory to succinate de-
hydrogenase, probably after hydrolysis, but there is no evidence at present
for this and it is quite possible that the mechanism is entirely different.
Despite the lack of evidence for a high susceptibility of the metabolism or
growth of isolated neoplastic cells to malonate, the in vivo work has brought
out interesting effects that deserve more thorough investigation.
CELLULAR AND TISSUE FUNCTION
Many studies of the effects of malonate on physiological function with
the object of relating the cellular activity to succinate oxidase or the cycle
have been reported, but in only a few instances have the necessary data
been obtained and a relationship adequately established. The general rela-
tions between enzyme inhibition and changes in cellular function were
discussed in Chapter 1-9, and several methods for demonstrating correla-
tions were presented. The complexities of such studies were emphasized
and the difficulties commonly encountered are well illustrated in the results
of malonate inhibition. In addition to the various possible metabolic effects
of malonate, one must bear in mind that malonate, or the cations added
with it, can directly alter functional processes, actions which can be dis-
tinguished by the proper controls.
CELLULAR AND TISSUE FUNCTION 203
Single Cell Motility
A thorough study of the respiration and motility of the ciliate Paramecium
caudatum was made by Holland and Humphrey (1953). Malonate at 20 mM
inhibits the endogenous respiration 31% but has no effect on the ciHary
activity over 1 hr. The oxidation of cycle substrates, such as citrate, a-
ketoglutarate, fumarate, and malate, is well inhibited, and it is likely that
the cycle is present. Furthermore, the succinate oxidase in homogenates
is quite sensitive to malonate. One must conclude either that malonate
does not penetrate sufficiently or that the motility is not entirely dependent
on the cycle for an energy supply. The answer to this problem may lie in
the observation that malonate does not inhibit the oxidation of pyruvate
or acetate. Holland and Humphrey point out that there are many alternate
pathways for metabolism in paramecia. Thus it is possible that normally
energy is derived from the cycle but during a cycle block energy is provid-
ed by other alternate reactions. In the human parasitic ciliate Balantidium
coli, malonate depresses both respiration, which is probably endogenous,
and motility (Agosin and von Brand, 1953). The flagellar motility of Ba-
cillus brevis is not affected by 10 mM malonate (De Robertis and Peluffo,
1951). The motility of bull sperm is reduced appreciably by 10 mM mal-
onate and the endogenous respiration is simultaneously inhibited 55%
(Lardy and Phillips, 1945). However, the results are quite different when
various substrates are present and motility is depressed very little or not
at all. Since motility is not depressed with glucose or pyruvate as sub-
strate, whereas it is with acetate, it is likely that energy sources other
than the cycle can be used. Ciliary and flagellar activities in different or-
ganisms are thus affected in various ways by malonate and no uniform
picture emerges from the data at present available.
Chemotaxis and phagocytosis in guinea pig leucocytes are partially inhi-
bited by malonate but the concentration used (140 mM) was so high that
little significance can be attached to these results (Lebrun and Delaunay,
1951). Bacterial phagocytosis by human neutrophiles is depressed moder-
ately by malonate from 33.5 to 100 mM but no effect is seen with 6.7 mM
(Berry and Derbyshire, 1956). Malonate at 1 mM has no effect on the mi-
gration of amphibian chromatophores in cultures of the neural crest (Flick-
inger, 1949). At 10 mM, malonate is toxic to these preparations and not
specifically depressant to the motility. These limited results point to a
relative insusceptibility of ameboid-type movements to malonate, which is
not surprising considering the known metabolic characteristics of such cells.
Renal Tubular Transport
Most of the studies of the effects of malonate on the kidney have involved
the active transport of p-aminohippurate. The accumulation of this sub-
204 1. MALONATE
stance by kidney cortex slices, which is quite marked (slice/medium ratios
around 10), is reduced by malonate: in slices from dogs, 5 raM inhibits 25%,
10 mM inhibits 35%, and 20 mM inhibits 55% (Shideman and Rene, 1951 b)
and in slices from rabbits 20 mM inhibits 72% while reducing the respiration
with acetate by 53% (Cross and Taggart, 1950). This action can be shown
to occur in the whole animal also. Dominguez and Shideman (1953, 1955)
removed one kidney from rats, administered solutions of sodium malonate,
removed the other kidney, and determined the accumulation of j^-amino-
hippurate. When approximately 10 millimoles/kg of malonate are injected
subcutaneously, the uptake of p-aminohippurate is depressed 52%, the
average slice/medium ratio falling from 9.28 to 4.47. The decrease in the
slice/medium ratio is linear with intravenous doses of 4-7 millimoles/kg.
Some effect occurs at 15 min after the injections, the maximal inhibition
is around 60 min, and the transport mechanisms have returned to normal
by 150 min, indicating the ready reversibility of the action. The transport
inhibition can also be demonstrated by the renal clearance technique in
dogs (Shideman and Rene, 1951 b). Malonate at a dose of 0.96 millimole/kg
depresses the p-aminohippurate Tm* 73%. It was stated that 50% inhi-
bition of renal succinate dehydrogenase is produced by 1.32 mM malonate
and thus the dose given would be expected to inhibit in vivo, but the con-
centration of malonate in the tubular fluid is probably much higher than
in the blood and permeability factors must also be important. Farah and
Rennick (1954, 1956) studied the effects of many inhibitors on the p-amino-
hippurate uptake in guinea pig kidney slices and the results are summarized
in Fig. 1-18. Malonate is one of the weakest inhibitors but yet exhibits a
marked effect at 10 mM. Koishi (1959 a) confirmed the inhibition by mal-
onate on p-aminohippurate accumulation in rat kidney slices, obtaining
slight inhibition at 1 mM and 65% inhibition at 5 mM. The effects of mal-
onate are expressed in the following equation:
log (S/M) = 1.452 + 0.457 log (/) (1-6)
where S/M is the slice/medium ratio and (/) is the molar concentration of
malonate. The active transport of p-aminohippurate is thus definitely relat-
ed in some manner to succinate dehydrogenase and the cycle, assuming a
specific action of malonate, which is likely at the generally low concentra-
tions used.
This conclusion is somewhat substantiated by the findings that the renal
transport of other substances is frequently not inhibited potently by mal-
onate. The accumulation of tetraethylammonium ion is scarcely affected
by malonate up to 40 mM (Farah and Rennick, 1956; Farah, 1957), the
metabolic requirements apparently being different than for jj-aminohippu-
* Tm is the tubular transport maximal rate for a substance.
CELLULAR AND TISSUE FUNCTION
205
rate. Phenol red accumulation is also not depressed (Shideman and Rene,
1951 b), although in isolated flounder tubules it is suppressed by 10-20 roM
malonate (Forster and Goldstein, 1961). It was claimed that there is a cor-
relation between succinate oxidase activity and transport in different spe-
cies. Clearance studies with glucose and phosphate show that there is little
effect of malonate on their transport: e.g., when p-aminohippurate trans-
port is inhibited 73%, glucose Tm is decreased only 10%. On the other
hand, Malvin (1956) reported that malonate quite definitely depresses the
phosphate Tm, although no data were given. Since malonate interferes
with the uptake of inorganic phosphate in kidney homogenates, it was
concluded that malonate in some manner suppresses the esterification of
phosphate during its transport.
Fig. 1-18. Effects of inhibitors on the slice/medium (S/M) ratio
for p-aminohippurate in kidney shces. DNP = 2,4-dinitrophenol,
FA = fluoroacetate, lA = iodoacetate, DHA = dehydroacetate,
CN = cyanide, and F = fluoride. (Modified from Farah and
Rennick, 1956.)
Renal electrolyte transport is disturbed by malonate (Mudge, 1951).
Rabbit renal cortex slices were leached for 2.5 hr in 0.15 M NaCl, this
lowering the tissue K+ concentration and reducing the endogenous respir-
ation. The slices were then incubated for 30 min in medium containing
10 mM K+, 10 mM acetate, NaCl to provide a constant osmotic pressure,
and phosphate buffer, during which time K+ enters the cells. Malonate at
50 mM almost completely blocks this return of K+ into the cells and re-
verses the movement of water (see accompanying tabulation). The Na+
changes are not so significant because the substitution of divalent anions
for chloride increases the external, and presumably the internal, Na+ con-
centration. If fresh slices are treated with malonate, a loss of cell K+ and
206 1. MALONATE
Initial Control Malonate 50 xnM % Change
espiration (Qo^)
—
0.69
0.37
'^ater content (%)
79.0
78.2
81.1
+ (meq/kg wet wt.)
24.7
58.0
28.1
a+ (meq/kg wet wt.)
112
93.8
139
+ + Na+
136.7
151.8
167.1
- 46
- 90
+ 100
an equivalent gain of Na+ would be expected due to the inhibition of the
transport mechanism responsible for K+ accumulation. The effect on water
transport is really quite marked and greater than with some fifty other
inhibitors used. Mudge suggested that malonate acts by depression of
aerobic metabolism in general and not necessarily by a specific inhibition
of succinate oxidase, which at the high concentration used is quite possible.
The results on malonate in vivo occasionally do not correspond to the
in vitro experiments, nor do they always correspond to each other. The in-
jection of 10 millimoles/kg malonate into rats leads to a considerable diu-
resis which lasts for several days (Angielski et al., 1960 a). On the other
hand, infusion of malonate into the renal artery of a dog at a concentration
of 8.7 mM causes no significant change in creatine, p-aminohippurate, or Na+
clearances, and produces a 15% suppression of urinary volume (Strickler
and Kessler, 1963). The effects in intact animals are related to acid-base
imbalance in addition to direct renal action, as shown by unpublished ex-
periments by Goldberg (1963) in which rats were water loaded and received
17 millimoles/kg sodium malonate with 17 ml 10% mannitol/kg, this pro-
ducing certain toxic symptoms (e.g., respiratory difiiculty, sluggishness,
and mild cyanosis). The results are summarized in the accompanying tab-
ulation and it is clear that a systemic acidosis was produced. A rather
Determination Control Malonate
Urinary flow (ml/min) 0.054 0.058
Urinary pH
Titratable acidity (meq/liter)
Creatinine clearance (ml/min)
Na+ excretion (meq/liter)
K+ excretion (meq/liter)
NH4+ excretion (meq/liter)
Plasma pH
Plasma Na+ (meq/liter)
Plasma K+ (meq/liter)
6.72
5.45
17.0
46.7
1.1
0.89
9.5
110
10.3
57.5
25.7
16.3
7.39
7.23
50
198
5.4
9.5
CELLULAR AND TISSUE FUNCTION 207
clear increase in phosphate excretion with a fall in plasma phosphate was
also observed.
The effects of malonate on the renal excretion of malate are interesting,
but it is not known if this phenomenon is related to an action on transport
mechanisms or to more general metabolic effects. Vishwakarma (1957,
1962) showed that malonate has no effect on the excretion df malate in-
duced by malate infusion. However, when succinate is infused, the in-
creased malate excretion is due to both an increased filtration and a marked
tubular secretion. Malonate inhibits the tubular secretion of malate and
converts the excretion to a purely filtration process or causes a net resorp-
tion. Vishwakarma and Lotspeich (1960) continued this study in chickens
and found that when malonate is infused with succinate, instead of block-
ing the formation of malate, it further increases the malate excretion.
Malonate was infused at a rate of about 6.8 //moles/kg/min. This could
mean that malonate (1) enhances the formation of malate from succinate,
(2) facilitates the tubular secretion of malate, or (3) gives rise to malate by
metabolic conversion. Since malonate infused alone did not significantly
increase malate excretion, the last explanation is unlikely. In the dog, mal-
onate inhibits the tubular secretion rather than stimulating it and does not
inhibit the resorption of malate. The mechanism for this paradoxical effect
is unknown. Reference may be made to studies of Lotspeich and Woron-
kow (1964), who unilaterally perfused chicken kidneys and found complex
effects on the excretions of various organic acids, and concluded that the
cycle must be involved in some manner.
Transintestinal Transport
Quastel has studied the effects of various inhibitors on the transfer of
glucose across the guinea pig intestinal wall. This is an active transport
and depends strongly on the aerobic metabolism (as shown by the marked
inhibition by cyanide and azide) and the associated phosphorylations (as
shown by the 2,4-dinitrophenol inhibition). When malonate at 20 mM is
present in the lumen, there is 18.5% inhibition of the glucose transported,
but if malonate is present both inside and outside, the inhibition is 44.3%
(Darlington and Quastel, 1953). An increase of K"'" from 6 to 15.6 n\M ac-
celerates glucose transport about 50%. Malonate inhibits the K^-stimulated
transport completely at concentrations as low as 2 mM (Riklis and Quastel,
1958). This result may be related to the greater sensitivity of K+-stimulated
brain slices to malonate. It was claimed that 20 mM malonate depresses
both the accumulation of L-monoiodotyrosine-I^^^ in the intestinal cells and
its transport across the intestine, but no data were given (Nathans et al.,
1960). There is a marked difference between transintestinal transport and
tissue accumulation of triiodothyroacetate, the former being inhibited much
208 1. MALONATE
more readily by a number of substances; malonate at 10 mM inhibits up-
take 16% and transport 70% (Herz et al., 1961).
Everted segments of the rat duodenum transport iron from the mucosal
to the serosal surface against concentration gradients and this process is
dependent on oxidative phosphorylations (Dowdle et al., 1960). Malonate
at 50 mM reduces the inside/outside ratio of Fe++ from 4.0 to 0.6. Ca++ is
also transported actively and malonate at 20 mM reduces the inside/outside
ratio of Ca*^ from 5.0 to 2.5 (Schachter and Rosen, 1959). Ca++ is also ac-
cumulated by the intestinal cells, the tissue/medium ratio being 5.8, which
is decreased by 20 mM malonate to 3.4 (Schachter et al., 1960). The question
of the chelation of Ca++ and Fe+++ by the malonate arises, since the high
malonate concentrations would certainly reduce the free ions appreciably.
This must play some role but in the case of Ca++ cannot explain the reduc-
tion in the transport, inasmuch as the inside/outside ratio is increased as
the Ca++ concentration is lowered.
Gastric Acid Secretion
The secretion of hydrochloric acid by the parietal cells is dependent on
oxidations and the formation of ATP, since it is strongly inhibited by cya-
nide, antimycin, and 2.4-dinitrophenol. However, the secretion in isolated
rat stomachs is not affected by 10 mM malonate (Patterson and Stetten,
1949). Injection of malonate in mice subcutaneously inhibits the accumu-
lation of p-aminohippurate in the kidney but does not inhibit acid secretion:
4.8 millimoles of malonate reduces the kidney/medium p-aminohippurate
ratio from 6.1 to 2.9 but inhibits the secretion of hydrochloric acid only
6% (Davenport and Chavre, 1956). This is near the fatal dose of malonate
and many of the mice did not live long enough to perform the test. Inas-
much as succinate oxidase activity is high in the stomach and is readily
inhibited by malonate, and since fluoroacetate inhibits acid secretion, the
most likely explanation for the lack of a malonate effect is a failure to
penetrate into the parietal cells sufficiently. Some evidence for the partici-
pation of succinate oxidase in acid secretion was obtained by Vitale et al.
(1956), who showed that stimulation of guinea pig or human gastric mucosa
with histamine leads to significant increases in the succinate oxidase activity,
although histamine has no such effect in liver or duodenum. Furthermore,
succinate oxidase is concentrated in those regions of the stomach where
the parietal cells are abundant.
Active Transport of Ions in Various Cells and Tissues
The effects of malonate on nerve and muscle, to be discussed in the fol-
lowing sections, depend in part on the modification of the active transport
of ions in these tissues. Malonate depresses many types of active transport.
CELLULAR AND TISSUE FUNCTION 209
as we have seen for kidney and gastric mucosa, and the mechanism may be
either a simple reduction in the energy supply or a more direct interference
with electron transport associated with ionic movements across the mem-
brane. One must also attempt to distinguish between effects on the active
transport and increases in permeability. If the permeability to an ion is
significantly increased, its intracellular accumulation may drop because the
ion pump is no longer able to maintain the normal concentration; such an
action would appear superficially to be an inhibition of active transport.
Malonate up to 10 mM has no effect on the transport of Na+ and K+
across the human erythrocyte membrane (Maizels, 1951), which is not sur-
prising since the principal energy source in such cells is glycolytic. In ascites
tumor cells, substrates (for example, glucose and succinate) increase the
efflux of Na+. Cyanide at a concentration inhibiting respiration 70% has
no effect on either the influx or efiiux of Na+, presumably because the rate
of aerobic glycolysis is simultaneously doubled, this compensating for the
oxidative depression (Maizels et al., 1958). Malonate at 12.5 mM, on the
other hand, inhibits respiration 35% but produces only a 5% increase in
the glycolysis, which may explain why the rate coefficient for Na+ efflux
drops from 6.5 to 5.1 hr-^ The accumulation of intramitochondrial K+ in
preparations from rabbit heart is dependent on oxidative phosphorylation
but is unaffected by 0.2 mM malonate (Ulrich, 1960). Inasmuch as a-keto-
glutarate was the substrate used, even a complete block of succinate oxida-
tion might not be expected to have much effect on ion movements because
sufficient ATP may be generated in the single-step oxidation of a-keto-
glutarate. Ca++ uptake and binding by kidney mitochondria depend on an
oxidizable substrate and ATP; it is depressed 75% by 10 mM malonate,
which suggests interference with the operation of the cycle, but could
relate to the chelation of the Ca++ by the malonate (Vasington and Murphy,
1962). The uptake of iodide is inhibited by rather high concentrations of
malonate in the brown alga Ascophyllum, nodosum (79% inhibition at 25 mM)
(Kelly, 1953), the rabbit ciliary body (50% inhibition at 50 mM) (Becker,
1961), and the rabbit choroid plexus (50% inhibition at 20 mM) (Welch,
1962), but 1 mM malonate has no effect on the uptake or incorporation of
iodide in sheep thyroid particulate fractions (Tong et al., 1957).
The accumulation of P,^^ in the roots of the loblolly pine Pinus taeda
during a 3 hr incubation is inhibited 5% at pH 4.75 but stimulated 54%
at pH 5.75 (Kramer, 1951). Similar results are obtained in mycorrhizal
root tips but the inhibition is somewhat greater. It is possible that at the
concentration (25 mM) of malonate, the stimulation is an ionic effect which
is partially counteracted by a malonate inhibition at the lower pH. The
uptake of K+ and Br~ by barley roots is quite strongly inhibited by mal-
onate at pH 4.5 (see accompanying tabulation) (Ordin and Jacobson, 1955).
The inhibition is overcome to some extent by malate and fumarate; sue-
210 1. MALONATE
cinate, however, actually increases the inhibition. It is likely in this tissue
that ion accumulation is obligatorily coupled to the operation of the cycle.
Malonate
% Inhibition of:
(mM)
K+ uptake
Br- uptake
Respiration
5
10
39
92
42
70
30
55
Effect of Malonate on Mitochondrial Swelling
Eat liver mitochondria swell quite readily, as measured by changes in
light scattering or optical density, when treated with various substances,
and the effects of malonate on this phenomenon are interesting. Raaflaub
(1953) established that succinate and phosphate promote swelling. This
swelling is counteracted by ATP in both cases, but malonate prevents the
swelling from succinate only. This was confirmed by Tapley (1956), who
extended the list of substances causing swelling to fumarate, malate, gluta-
mate, acetate, and a-ketoglutarate. Swelling is prevented by citrate, pyru-
vate, and oxalacetate, as well as malonate. Since malonate can prevent the
swelling from substrates other than succinate, there is some question as to
the specificity of the effect. It was claimed that the same results are obtain-
ed in the absence of oxygen and thus that the swelling is not related to the
utilization of these substrates. Quite different conclusions were reached by
Chappell and GreviUe (1958) inasmuch as they found a good correlation
between swelling and utilizable substrates. Malonate prevents the swelling
from succinate but not from a-hydroxybutyrate, and, in general, inhibitors
blocking oxidations reduced swelling. Matters were further complicated by
the results of Keller and Lotspeich (1959 b). They found that phlorizin
caused swelling of kidney mitochondria and that this could be counteracted
by Mg++, ATP, 2,4-dinitrophenol, and malonate. Hunter et al., (1959 a, b)
considered the possibility that swelling is related to the fraction of NAD in
the oxidized form, since amobarbital prevents oxidation of NADH and
prevents swelling. However, succinate in the presence of amobarbital causes
swelling and this is blocked by malonate. Glutamate-induced swelling is not
prevented by malonate. It was concluded that swelling depends on electron
flow between the substrate and oxygen, and whether or not an inhibitor will
prevent swelling is determined by where in the electron transport chain
the substrate and the inhibitor act. This does not very well explain the
prevention of swelling by 2,4-dinitrophenol and it was suggested that there
are at least two different types of mitochondrial swelling. Further confusion
was introduced by Sabato and Fonnesu (1959), who found that swelling is
CELLULAR AND TISSUE FUNCTION 211
prevented by oxidizable substrates such as succinate, glutamate, and a-
ketoglutarate, and that malonate counteracts this preventive action. These
results were confirmed by Kaufman and Kaplan (1960), who again ob-
served, in contrast to the earlier workers, that succinate inhibits swelling
and malonate reverses this protection. They believe that swelling is correl-
ated with the mitochondrial release of pyridine nucleotides (see tabulation).
Pyridine nucleotide released
(//g/20 min)
Optical density
No substrate 64 —0.710
Succinate (20 mM) 14 —0.095
Malonate (20 mM) 68 -0.740
Succinate + malonate 38 — 0.520
Succinate reduces the loss of the pyridine nucleotides and malonate anta-
gonizes this effect. One must conclude that there must be different mecha-
nisms of swelling and that the mitochondrial behavior perhaps depends on
the metabolic state and the nature of the suspension medium. The effects
of malonate and other anions on the concentrations of free Ca++ and Mg++
should also not be ignored, inasmuch as EDTA has usually been shown to
modify the swelling.
Conduction and Membrane Potentials of Nerve
Penetration of malonate into nerve axons in the physiological pH range
must be very poor. This may account for the failures of Shanes and Brown
(1942) to observe an eft'ect of 20 mM malonate on the resting potential of
frog nerve, and of Greengard and Straub (1962) to find an effect of 10 mM
malonate on nonmyelinated nerve posttetanic hyperpolarization, despite
the fact that this phenomenon is quite sensitive to other inhibitors. How-
ever, Jenerick (1957) reported some effect of 10 mM malonate on frog sciatic
nerve although the action was presumably slow in developing. When the
action potential spike amplitude is reduced by 80-90%, the threshold for
stimulation begins to rise rapidly. Conduction block occurs when the rest-
ing potential has fallen by 30-40%. It is doubtful if the decrease in external
Ca++ concentration, which was 1.3 mM initially, resulting from chelation
by malonate could be held responsible for these effects, and it was felt
that metabolic interference must occur. The preganglionic and postgan-
glionic action potentials in preparations of cat sympathetic ganglia are
depressed equally (75-80%) by 14 mM malonate and transmission through
the ganglia is reduced (Larrabee and Bronk, 1952). The excitability of the
212 1. MALONATE
isolated cat carotid body is lowered by perfusion with malonate (Anichkov,
1953). These meager data are all we have to understand the actions of mal-
onate on nerve function. Unfortunately, little has been done on junctional
transmission, inasmuch as it might be predicted that the synapses would
be more sensitive to malonate than are the axons, because of both a higher
permeability of such regions to anions and a greater energy requirement
for the synthesis of acetylcholine.
Skeletal and Smooth Muscle Function
Essentially nothing is known of the effects of malonate on skeletal muscle.
Beckmann (1934) claimed that 6.7 mM malonate causes a swelling of muscle,
indicating an alteration of permeability. This was termed a membrane-
loosening effect. In the initial work of Ling and Gerard (1949) with intra-
cellular microelectrodes, it was observed that 10 toM malonate drops the
resting potential of frog sartorius muscle from 78 mv to 65.3 mv over a
period of 3 hr. This may be correlated with the suppression of Na+ ex-
trusion observed by Kernan (1963) in the same muscle, 30% inhibition
being produced by 1 mM malonate over 2 hr, an effect similar to that oc-
curring in brain slices (Bilodeau and Elliott, 1963). No direct work on the
contractile response to malonate has been done.
The contractions of isolated rabbit intestine are not inhibited by 10 mM
malonate, whether in the absence of substrate or in the presence of either
acetate or glucose (Weeks and Chenoweth, 1950; Weeks et al., 1950). In-
deed, there is a tendency for malonate to increase the contractile activity
slightly, especially with glucose as the substrate. There is also no interfer-
ence with the recovery of substrate-depleted strips produced by the ad-
dition of acetate or pyruvate. It was suggested that a lack of penetration
of malonate into the smooth muscle cells might be responsible. Fluoroacetate
is quite inhibitory under the same conditions so that some relationship of
the contractility to the cycle is likely. The contractile properties of the
vascular smooth muscle in the cat hind limb are not affected by 1 mM
malonate (Hitchcock, 1946), and the behavior of electrically stimulated pig
carotid artery is not altered by 10 mM malonate (Jacobs, 1950).
It would be important to know more about the possible effects of mal-
onate on the formation and release of the neurohormones, such es acetyl-
choline and the catecholamines, but the data are not available. It is in-
teresting to note, however, that malonate is reasonably effective in inhi-
biting the release of histamine from guinea pig lung slices during an ana-
phylactic reaction (Moussatche and Prouvost-Danon, 1958). The inhibition
is 10% at 20 mM, 40% at 40 mM, and 50% at 60 mM. The inhibition was
attributed to the effect on succinate dehydrogenase. Nevertheless, malonate
at 40-60 ToM has virtually no effect on the release of histamine brought
about by the application of the histamine-releaser Compound 48/80 (Mous-
CELLULAR AND TISSUE FUNCTION 213
satche and Prouvost-Danon, 1957), although respiration of the lung slices
is depressed fairly strongly. It would appear that malonate interferes with
the anaphylactic release of histamine by a mechanism other than a direct
effect on the formation or release of histamine. It may be noted that suc-
cinate accelerates the oxygen uptake but has no effect on the release of
histamine.
Cardiac Membrane Potentials and Function
The physiological disturbances produced by malonate have been most
thoroughly studied in the heart. Although the effects are often very slight,
despite the evident importance of the cycle in the myocardium, under cer-
tain conditions the responses to malonate are very interesting. The earliest
investigation was made by Forssman and Lindsten (1946) at Lund, who
noted a marked discrepancy between the effects of malonate in the whole
animal and on isolated hearts. Intravenous injections of malonate at doses
around 3.7-7.5 millimoles/kg to cats and rabbits lead to an increase in the
venous blood pressure and a fall in the arterial blood pressure, indicating
cardiac depression. In rabbits the cardiac failure begins about 20 min after
the injection whereas in cats the changes are immediate. In rabbits the
heart may stop after 40 min but in cats recovery is the rule. At autopsy
the heart is found to be dilated. The effects of malonate on the isolated
perfused rabbit heart, however, are rather small and inconsistent (see ac-
companying tabulation). Moreover, succinate at the same concentrations
Malonate °''» ^^^^^^ ^^
(mM)
Amplitude
Coronary flow
Rate
10
-14
-23
- 5
20
-19
-16
- 8
40
-34
-29
0
acts similarly. These are the immediate effects of malonate and it is possible
that the heart would recover from this depression after several minutes, as
do rabbit atria (Webb, 1950). It is doubtful if these effects are related to
inhibition of succinate oxidase; they are more likely ionic actions on the
membrane. The reduction in the coronary flow may result from a vascular
constriction, but is more probably the response to the decreased functional
activity.
Isolated rabbit atria are depressed only by high concentrations of mal-
onate, 30-40 mM producing a 30% decrease in contractile amplitude and
rate at 2 min; the depression is temporary, complete recovery being ob-
served after 8-10 min (Webb, 1950b). Atria can continue to beat normally
214 1. MALONATE
for hours in 50 mikf malonate. The temporary depression brought about
by malonate is not counteracted by fumarate added either before, with, or
after the malonate. In fact, fumarate, along with pyruvate, acetate, suc-
cinate, and malate, has an action very similar to that of malonate on the
atria. It is not known if this inhibition and recovery are related to the
somewhat slower but similar time course of ventricular respiration under
the influence of malonate (page 181) (Webb et al, 1949). The depression is
not due to chelation of Ca++ or Mg++ since reduction in the concentrations
of these ions produces a different response. Gardner and Farah (1954) con-
firmed the resistance of rabbit atria to malonate, finding that 10-20 milf
has no significant effects on contractility, spontaneous rate, excitability
threshold, refractory period, and conduction rate.
The effects of malonate were investigated more thoroughly on rat atria
(Webb and Hollander, 1959). The contractility is depressed 21% imme-
diately but slow recovery occurs: the inhibition is 13% during 5-25 min,
9% during 25-45 min, and 5% during 45-60 min. The malonate concen-
tration used was 15 mM. The addition of 15 mM NaCl produces a rapid
contractile depression about half as great as from malonate, so that at least
part of the initial malonate effect is attributable to the Na+ ion. A slight
initial rise in the magnitude of the action potential is observed with both
malonate and NaCl, but in the case of malonate this is soon replaced by a
small depression. There is also some shortening of the action potential and
a decrease in its area after the first 5 min, which could be responsible for
the fall in tension. In summary, the addition of 15 ml/ malonate produces
a rapid initial effect attributable mainly to the Na+ and this is progressively
replaced by changes due to the malonate, these latter changes being mod-
erate depressions of the action potential and contractility. The importance
of the cycle in the atrial function is indicated by the marked changes brought
about by fluoroacetate, and thus the resistance to malonate is probably due
to a low intracellular concentration of malonate. Greater effects on the con-
tractility of rat atrium were observed by Venturi and Schoepke (1960),
5 mM depressing 22%, 10 mM 44%, and 20 mM 90%. It was stated that
NaCl at these concentrations does not alter the contractility. Furthermore,
succinate is as inhibitory as malonate. The greater depression observed here
compared to my work is difficult to explain. Venturi and Schoepke used
Locke solution at pH 7 whereas I used Krebs-Ringer-bicarbonate medium
at pH 7.4. Part of the larger inhibition seen by Venturi and Schoepke thus
might be due to the lower pH. In any event, these effects seem to be un-
related to the inhibition of succinate oxidase and again must be attributed
to some action directly on the membrane. Venturi and Schoepke found that
increasing Ca++ concentration can completely overcome the depressant ac-
tions of malonate, succinate, and the other organic anions used, leading
them to suggest that the negative inotropic action is due to the chelation
CELLULAR AND TISSUE FUNCTION
215
of Ca+"'". However, there are some reservations in accepting this explanation
completely. In the first place, Ca++ stimulates atrial contractility and would
be expected to counteract most depressants in a nonspecific manner. In
the second place, reducing the Ca++ from 1.22 mM to 0.91 mM does not
alter the contractility, although further reduction to 0.61 mM depresses
43%. Malonate at 15 mM would reduce the Ca++ from 1.22 mM to 0.82 mM
and a small contractile depression may result from this. The total Ca++ in
Locke solution is 2.16 mM and 20 mM malonate would reduce the free
Ca++ to 1.31 mM, which alone could not produce the 90% depression seen
by Venturi and Schoepke. In the third place, monocarboxylate ions, such
as acetate, lactate, and pyruvate, at 20 mM depress the contractile am-
plitude 40-45% and these do not deplete the Ca++.
The modifications in the electrocardiogram following intravenous injec-
tions of malonate into turtles were studied by Lenzi and Caniggia (1953).
At a dose of 4.4 millimoles/kg malonate the following occurred: brady-
cardia, slowing of the a-v conduction, a tendency for the shortening of
repolarization and electric systoles, with eventually a total a-v block
and a prolongation of the depolarization time (see accompanying tabula-
tion). Pacemaker and conduction depression are thus evident, and it is
Control
Malonate
at 30-35 min
Control
at
Malonate
57-65 min
Rate
50
23
78
32
pq interval
0.30
0.465
0.24
block
qrs interval
0.15
0.18
0.10
0.18
st-t interval
0.57
0.895
0.44
1.06
qt interval
0.72
1 . 075
0.54
1.24
quite possible that similar changes would be observed in mammals, con-
sidering the general behavior of the heart in cats and rabbits treated with
malonate (Forssman and Lindsten, 1946). The electrocardiogram from the
embryonic chick heart is not altered by 4 mM malonate (Boucek and Paff,
1961).
In contrast to the depressant effects of malonate on the whole heart and
isolated atria, the rat ventricle strip is usually strongly stimulated, as first
noticed by Masuoka et al. (1952). Substrate-depleted and hypodynamic strips
recover to almost the initial contractile amplitude upon addition of 10 mM
malonate at pH 6.2 (which was used to facilitate penetration of the mal-
onate), and simultaneously the stimulatory action of succinate is blocked.
The ability of glucose to induce recovery is augmented by malonate, and
that of pyruvate is slightly reduced (Berman and Saunders, 1955). This
interesting positive inotropic action was analyzed in detail because it was
216 1. MALONATE
felt that such an action might have bearing on the mechanisms whereby
the cardioactive glycosides stimulate the failing heart. Only the major
results will be summarized here. The positive inotropic action occurs most
strongly when glucose is present, less in substrate-free medium, and not at
all with pyruvate or a-hydroxybutyrate as substrate (Covin and Berman,
1956). These results suggested that malonate might stimulate the Embden-
Meyerhof glycolytic pathway, resulting in an accelerated conversion of glu-
cose and glycogen to pyruvate. If this were so, pyruvate should produce a
comparable positive inotropic effect and it does in both substrate-depleted
and glucose-supplemented strips. Furthermore, iodoacetate at 0.2 mM
blocks the stimulation by malonate, whereas it does not affect the response
to pyruvate. The chelation of Ca++ was shown to contribute to the depression
produced by malonate at high concentrations (20-50 mM), and it probably
reduces the amount of stimulation seen at the lower concentrations since
lowering the Ca++ to the degree calculated to occur in 10 mM malonate
depresses the contractile activity 23%. The effects of malonate on the oxida-
tion of C^*-labeled substrates by ventricle strips were then studied in cham-
bers in which the respiration and contractile activity could be determined
simultaneously (Rice and Berman, 1961). Malonate at 5.6 mM under con-
ditions in which a positive inotropic effect is observed has very little effect
on the utilization of glucose- 1-C'^, glucose-6-C^*, and pyruvate-2-C^*, slight
inhibition of glucose oxidation being noted although this is possibly not
significant. These results indicate that the stimulatory action is not related
to (1) acceleration of glucose metabolism, (2) inhibition of the cycle, or (3)
stimulation of the pentose-phosphate pathway. It was found that C^^Og is
produced from malonate-2-C^* in ventricle strips, and possibly part of the
positive inotropic action in substrate-depleted strips is related to the oxida-
tion of malonate via formation of acetyl-CoA and its incorporation into
the cycle. However, the explanation for the greater effect of malonate in
the presence of glucose and the inhibition of its action by iodoacetate is
not immediately evident. It may be noted that other metabolic inhibitors,
such as fluoride, arsenate, fluoroacetate, and dehydroacetate, can exert po-
sitive inotropic actions under the appropriate conditions, so this paradoxical
effect of malonate is not unique.
Wenzel and Siegel (1962) determined the dose-response curves for the
positive inotropic effects of malonate and ouabain on the rat ventricle strip,
and then constructed an isobologram, plotting the malonate concentration
against the ouabain concentration for a chosen contractile stimulation.
Since the isobol sags, i.e., is concave upwards, they claimed it is clear that
potentiation occurs and that this indicates the sites of action of malonate
and ouabain are different. There is some doubt that a moderately sagging
isobol can be interpreted as potentiation, inasmuch as pure summation
often elicits such a curve (see Figs. 1-10-7, 8).
EFFECTS OF MALOXATE IN THE WHOLE ANIMAL 217
The response of the heart to neurohormones is not altered by malonate.
The positive chronotropic effect of epinephrine on the frog heart is not
changed by 0.1 roM malonate (Nickerson and Nomaguchi, 1950), which is
not surprising considering the concentration. A brief report by Ellis and
Anderson (1951 a) stated that malonate does not affect the stimulation by
epinephrine except after prolonged treatment when the frog heart is de-
pressed. The malonate concentration was not given. It would seem likely
that any severe metabolic disturbance producing marked cardiac depression
would prevent, or at least reduce, any type of stimulation, since the ad-
ditional functional activity would demand more energy, so that an an-
tagonism of epinephrine by malonate is not of much significance unless it
occurs when the heart is not too much depressed. Malonate has no demon-
strable effect on the response of the heart to acetylcholine, with respect to
reduction of either rate or contractility (Webb, 1950 b), whereas fluoroace-
tate alters the response markedly, indicating again that the cycle is of im-
portance in these myocardial functions but that malonate does not reach
sufficiently high intracellular concentrations.
EFFECTS OF MALONATE IN THE WHOLE ANIMAL
A summary of the miscellaneous results relating to toxic and lethal ef-
fects of malonate is given in Table 1-28. One may conclude that in mammals
injected doses of 1.5 2.5 g/kg (10 17 millimoles/kg) of sodium malonate are
generally lethal. Such doses, especially when given intravenously, probably
produce plasma levels in excess of 10 inM malonate at peak concentra-
tions. A dose of 12 millimoles/kg subcutaneously in rats leads to a plasma
concentration of 4.5 mill at 30 min, and another similar dose raises the
plasma level to around 8 mM (Busch and Potter, 1952 a). Intravenous
injection would give higher peak levels. When compared in the same ex-
periments, the acid is more toxic than the sodium salt. It is difficult to
know if this is due to a nonspecific acid effect or to better penetration into
the tissues. It would appear that malonate is more toxic to mice at 38° than
at 30° environmental temperature (Gruber et al., 1949).
The sequence of symptoms resulting from the injection of malonate into
rats and mice was described by Gruber et al. (1949) as: champing, air hunger,
maintenance of the head in a dorsally flexed position, and clonic convulsions.
Busch and Potter (1952 a) found dyspnea and convulsions in rats follow-
ing injection of toxic doses. The cause of death has been attributed to var-
ious actions. Forssman (1941) and Forsmann and Lindsten (1946) believed
that death is due to cardiac failure (the cardiovascular effects observed were
discussed in the previous section). Handler (1945) also favored a cardiac
mechanism for death and found the succinate oxidase to be inhibited 50-
75% in homogenates prepared from poisoned animals. He also noted that
218
1. MALONATE
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EFFECTS OF MALONATE IN THE WHOLE ANIMAL 219
the heart is dilated at death and that there is an accumulation of ascitic
and pleural fluid. Forssman and Lindsten, from their failure to obtain appre-
ciable direct action on the isolated heart, suggested an indirect mechanism,
possibly mediated through the effect of malonate on liver metabolism. The
dyspnea and various central nervous system effects may result from a direct
action of malonate but could also arise from the ionic and acid-base im-
balances produced. Handler (1945) showed that malonate causes a marked
fall in CO2 capacity in the blood from 53 to 6 vol%. Although no study of
the alterations in the plasma electrolytes undoubtedly produced by mal-
onate has been made, Wick et at. (1956) noted that the ability of the blood
to coagulate is reduced, and attributed many of the actions of malonate to
the chelation of divalent cations. Other changes in the blood composition
may also be responsible for some of the toxic effects. Handler (1945) re-
ported that 1.6 g/kg malonate given subcutaneously to rabbits increases
the blood glucose 368%, blood lactate 545%, blood pyruvate 163%, serum
inorganic phosphate 262%, and serum organic phosphates 155%. (The keto-
nemia produced by malonate was discussed previously). There is thus much
opportunity for secondary mechanisms to play a role in the toxicity of
malonate. At the present state of our knowledge it is even difficult to evaluate
the importance of succinate oxidase inhibition in these effects.
The kidneys have the highest concentration of malonate after administra-
tion and therefore the renal effects and nephrotoxicity have been investigat-
ed. Early arguments about the renal toxicity of glutarate were published
between 1907 and 1912. Rose (1924) "reinvestigated this and found that
glutarate is a nephrotoxic substance in rabbits, as indicated by the increases
in nonprotein nitrogen, urea, and creatinine, and the almost complete disap-
pearance of renal function as measured by the phenolsulfonphthalein test.
A single experiment with malonate was reported. No renal damage was
observed after 2 g given on successive days and 3 g 2 days later, nor was
there a change in the rate of dye excretion. Corley and Rose (1929) reported
that methylmalonate and ethylmalonate are slightly toxic to the kidneys
at doses of about 1 g/kg in rabbits, there being a definite increase in the
nonprotein nitrogen and some reduction in dye secretion, although both
effects are transitory. Extensive renal damage was observed by Becker and
Rieken (1954) following the intraarterial injection of 20 mg potassium mal-
onate (Fig. 1-19 a). The vessel walls become edematous, potocytosis is evi-
dent, and many perinuclear vacuoles appear in the loops of Henle. However,
it requires much higher doses to depress the respiration of kidney slices
prepared from injected animals, 80 mg potassium malonate giving no effect
and 150 mg inhibiting 30.5%. Similar histological changes occur after in-
cubating kidney tissue in vitro with 110 mil/ potassium malonate for 20 min
(Fig. 1-19 b), vacuolization being intense. Tinacci (1953) found not only
kidney damage but widespread degenerative changes 2-8 days after sub-
220
1. MALONATE
Fig. 1-19. Renal damage resulting from (a) intraarterial injection of 20 mg potas-
sium malonate into rabbits, and (b) incubation of slices with 110 mif malonate for
20 min. (From Becker and Rieken, 1954.)
EFFECTS OF MALONATE ON BACTERIAL INFECTIONS 221
cutaneous injections of 0.4-0.8 g/kg of sodium malonate, almost all organs
being involved. Malonate is diuretic when subcutaneously injected into
hydrated rats at a dose of 11.5 millimoles/kg, this dose being sufficient to
inhibit kidney succinate oxidase (Fawaz and Fawaz, 1954). However, the
effect is probably osmotic rather than due to enzyme inhibition by the mal-
onate, since KNO3 at the same dosage induces the diuresis, and also be-
cause diethylmalonate, which inhibits kidney succinate oxidase, has no
diuretic activity. Summarizing these results, one may conclude that renal
damage may occur from high doses of malonate, but that minimal changes
in the kidney occur after the usual toxic doses. It seems unlikely that the
renal action is of major importance in poisoning or death from malonate.
EFFECTS OF MALONATE ON BACTERIAL INFECTIONS
The influence of enzyme inhibitors on the course of bacterial infections
is of great interest because the results have bearing on the fundamental
question of the metabolic basis of the resistance to infection. The effects
of malonate on Salmonella typhimurium infection in mice have been studied
by Berry and co-workers at Bryn Mawr in a series of excellent investiga-
tions. Mice injected intraperitoneally with a Salmonella suspension show
evidence of the infection on the third day and most succumb by the sixth
day. If mice are given intraperitoneal injections of 20 mg sodium malonate
in saline every hour for 8 hr, they die much more rapidly from the infection
than animals given only saline injections (Berry and Mitchell, 1953 a).
The striking effects is illustrated in Fig. 1-20. Noninfected mice show no
effects of the malonate. Thus sublethal doses of malonate are able either
to accelerate bacterial proliferation or to decrease the resistance of the
host markedly. These are the basic observations and the later work attempts
to elucidate the mechanisms by which these effects are brought about.
The reduction of the survival time in mice infected with Salmonella is
not unique. Similar effects of malonate on infections with Proteus morganii,
Staphylococcus aureus, Streptococcus pyogenes (Berry and Mitchell, 1954),
Diplococcus pneumoniae, and Klebsiella pneumoniae (Berry et al., 1954 a)
have been observed. Furthermore, reduced survival times have been found
in Salmonella infected rats, guinea pigs, and chickens (Berry and Beuzeville,
1960). Finally, the phenomenon has been seen with other inhibitors, such
as fluoroacetate and arsenite (Berry et al., 1954 a, b). This, then, is a gen-
eral effect of certain types of inhibitor, especially those affecting the cycle
in some manner, and the problem is thus more important because it must
relate to some fundamental metabolic relationship between host and bacteria.
We shall now examine in greater detail some of the characteristics of
the malonate effect. Malonate not only can reduce survival time, but in
some instances can change a nonlethal infection into a lethal one, this being
222
1. MALONATE
observed with Diplococcus, Staphylococcus, and Proteus. That is, an infec-
tion which does not kill plus a dose of malonate which is non-toxic may
cause the death of all the animals within several hours (Berry et al., 1954 a).
This is a true case of synergism. It has been found that Salmonella bactere-
mia in mice is much greater in malonate-treated animals than in the con-
trols (Berry and Mitchell, 1953 b, 1954). The bacteremia in the controls
reaches a low peak value soon after inoculation and then falls off, whereas
in the presence of malonate it continues to progress. At 9 hr, the blood of
No. 10
Dead
/ Malonate
/
j
/Saline
/ controls
.
y
24 30 36 42 48
Time after injection ( Hours)
Fig. 1-20 Effect of malonate given in 8 hourly injections of 20 mg
sodium malonate on the mortality of mice intraperitoneally in-
jected with suspensions of Salmonella typhimurium . (From Berry
and Mitchell, 1953 a.)
the controls contains 10,000 to 20,000 bacteria/ml whereas the blood of
malonate-treated animals has a count of around 3,000,000 bactcria/ml.
Thus the bacteremia is over 100-fold as severe in the treated animals as in
the controls. This bacteremia is a reflection of the situation throughout
the body. The total number of bacteria in the body 6.5 hr after the injec-
tions is 1.65 X 10^ in the controls and 32.1 X 10^ in the malonate-treated
mice (Berry, 1955). The ratio of counts in the treated and control series is
20 for the whole body and 19 for the blood at this time. Now, the interesting
thing is that, at the time of death, the number of bacteria in the body is
the same in both the controls and malonate-treated mice. This clearly shows
that malonate does not alter the susceptibility of the mice to the bacteria,
but reduces the time required for the bacteria to multiply to that number
necessary to kill.
EFFECTS OF MALONATE ON BACTERIAL INFECTIONS 223
The rapid proliferation of bacteria could be due to a weakening of the
body defenses for disposing of the bacteria. This does not seem to be the
case, inasmuch as malonate has no effect on the uptake of thorotrast by
the reticuloendothelial system (Berry, 1955) nor does it depress phagocytosis
except at very high concentrations (Berry and Derbyshire, 1956). Instead it
would appear that malonate disturbs metabolism in such a way that it
creates a more favorable environment in the host for bacterial growth.
Malonate itself may be metabolized slightly by Salmonella, but not to the
extent required to explain the explosive proliferation (Berry and Beuzeville,
1960). Growth medium was prepared from the eviscerated carcasses of con-
trol and malonate-treated animals and it was found that the bacteria grow
more rapidly in the latter (Berry, 1955). It has also been shown that Sal-
monella grows more rapidly in the peritoneal fluid of malonate-treated mice
than in the controls (Berry and Beuzeville, 1960). Citrate is known to ac-
cumulate following the administration of malonate. This was confirmed in
mice given the doses of malonate capable of reducing survival times of
infected animals (Berry et al., 1954 b). Both malonate and endotoxin from
Salmonella increase citrate levels in most tissues, and together the increases
are often greater than with either alone (see accompanying tabulation). It
Treatment
Citrate (//g/g) in
None
42
94
76
HI
Malonate
43
245
225
120
Endotoxin
50
273
190
187
Malonate
+ endotoxin
70
173
173
407
Blood Spleen Kidney Heart Duodenum Liver
132 109
115 170
170 443
543 240
is thus possible that a summation of effects on the cycle could be partially
responsible for the increased mortality. However, Salmonella infection does
not increase citrate levels (Berry and Beuzeville, 1960). Could the increased
citrate be favorable to the growth of the bacteria? It is unlikely that this
is a major factor because malonate is the most potent inhibitor for reducing
survival times and yet both arsenite and fluoroacetate cause greater accu-
mulations of citrate. The primary cause of the augmented bacterial proli-
feration has not been found but the range of possible mechanisms has been
narrowed. Since there are many other possible substrates for Salmonella
that accumulate during malonate inhibition, it will be necessary to examine
these in mice and their effects on the growth of Salmonella.
Some work on this problem in other laboratories may be briefly mention-
ed. Malonate reduces the antibacterial activity of guinea pig blood toward
224 1. MALONATE
Salmonella enteritidis but this is not due to a decrease in the number of
leucocytes (Yamauchi, 1956). The survival times of chicks infected with
Salmonella pullorum are reduced by 500-800 mg/kg of malonate injected
simultaneously or shortly after the bacterial inoculation (Gilfillan et al.,
1956). On the other hand, the diethyl ester of malonate increases the sur-
vival time of mice infected with Mycobacterium, tvbercvlosis, when admin-
istered daily for 2 weeks at oral doses of 250-500 mg/kg (Davies et al.,
1956). Compounds of this type are thus considered worthy of study as
chemotherapeutic agents in tuberculous infections.
METABOLISM OF MALONATE
Malonate occurs normally in many types of organism and occasionally
at concentrations possibly inhibitory to succinate dehydrogenase. Many
organisms are capable of metabolizing malonate by various pathways and
some are able to utilize it for growth or cell functions. In some cases, in-
deed, it is difficult to demonstrate the inhibitory action of malonate in the
presence of its own oxidation. The metabolism of malonate must always be
considered in studies of the effects of malonate on any type of cellular ac-
tivity. It is often impossible to detect and correct for the metabolism of
malonate without using labeled malonate. It is possible that many studies
of the inhibition of respiration or cycle activity by malonate have been
complicated by the oxidation of the malonate.
Occurrence of Malonate
Malonate has been isolated or demonstrated chromatographically from a
number of microorganisms, plants, and animals, and it is likely, consider-
ing the recent demonstration of its role and the role of malonate derivatives
in fatty acid metabolism that its occurrence is widespread. The accom-
panying incomplete tabulation will serve to illustrate this. Malonate has
also been found in winter wheat, barley, oats, alfalfa, kidney bean leaves,
clover, pea leaves, vetch leaves (Soldatenkov and Mazurova, 1957), sake
(Kawano and Kawabata, 1953), and several products prepared from plants.
Although no thorough studies of animal tissues have befen made, it is
evident that malonate must occur in rat, dog, and human tissues to some
extent if it is found in the urine. Although the name of malonate comes
ultimately from the Latin mains, it has never been identified in apples
or other fruit.
Methylmalonate has been found in Propionibacterium (Stjernholm and
Wood, 1961; Wood and Stjernholm, 1961), pigeon liver (Bressler and Wakil,
1961), pig heart (Flavin et al., 1955), mouse adipose tissue (Feller and Feist,
1957), and rat and human urine (Boyland and Levi, 1936; Barness et al.,
METABOLISM OF MALONATE
225
Source of malonate
Reference
Achromobader guttatum
Nocardia corallina
Penicillium funiculosum
Aspergillus niger
Phaseolus vulgaris (bush bean)
Phaseolus coccineus (runner bean)
Wheat plants
Bunias orientalis (Cruciferae)
Tobacco leaves
Lucerne (green alfalfa)
Hevea brasiliensis
Helianthus annus
Umbelliferae {Anthriscus and Apium)
Leguminosae (18 species)
Rat urine
Dog urine
Human urine
Sguros and Hartsell (1952a)
i.ara (1952)
Igarasi (1939)
Challenger et al. (1927) Walker
et al. (1927), Subramanian
et al. (1929)
Young and Shannon (1959),
Rhoads and Wallace (1957),
Huffaker and Wallace (1961)
Bentley (1952)
Nelson and Hasselbring (1931)
Jermstad and Jensen (1951)
Wada and Kobashi (1953), Bel-
lin and Smeby (1958), Vickery
and Palmer (1956 b, 1957),
Vickery (1959)
Turner and Hartman (1925)
Fournier et al. (1961)
Bentley (1952)
Bentley (1952)
Bentley (1952)
Stalder (1958), Thomas and
Stalder (1958)
Thomas and Kalbe (1953)
Stalder (1958)
1957; Stalder, 1958; Thomas and Stalder, 1958). Ethylmalonate has been
found in rat and human urine (Stalder, 1959). Hydroxymalonate (tartronate)
occurs in Acetohacter (Stafford, 1956) and malonic semialdehyde in Pseudo-
monas (Nakamura and Bernheim, 1961).
The concentrations of malonate in plant tissues are often surprisingly
high. The legumes and umbellifers analyzed by Bentley (1952) contain
0.5-2 mg/g fresh tissue. These values correspond to 7-30 raM malonate if
distributed uniformly throughout the tissue water. The stems of the runner
bean {Phaseolus coccineus) contain 2.1 mg/g and the expressed juice of the
stem is 30 milf in malonate. Since 20 raM malonate at pH 4.5 inhibits the
respiration of these stems 50% and causes accumulation of succinate, one
would expect the metabolism in these plants to be constantly suppressed
by the malonate, unless the malonate is in some manner segregated from
the metabolic systems. Soldatenkov and Mazurova (1957) reported similar
226 1. MALONATE
values in legumes (2-3% of the plant dry weight) and that in kidney-bean
and clover leaves malonate represents 45% of the total di- and tricarbox-
ylates present. Bush-bean {Phaseolus vulgaris) leaves often contain as
much as 10 mg/g dried tissue and malonate is more concentrated than fu-
marate or succinate, although less than malate and citrate (Young and
Shannon, 1959). In man malonate is excreted in the urine at an average
rate of 0.0047 mg/kg/day and in the rat at 10 times this rate (Stalder, 1958).
This amounts to only 0.32 mg/day in man (only two individuals were tested
so these averages are not accurate). Since malonate is apparently metabol-
ized in mammals, the tissue concentration or rate of excretion will reflect a
balance between formation and destruction. In other words, these excretion
values do not necessarily represent the rates of malonate formation.
Relatively little is known about the pathways for the formation of mal-
onate, but the miscellaneous observations make it likely that different reac-
tions are involved in various organisms. Malonate can arise from many
different substrates but in most cases the pathways are complex and the
immediate precursors are not known. Malonate can be formed from pyri-
midines and barbiturates in the mycobacteria (Hayaishi and Kornberg,
1952), from pyrimidines in Nocardia (Lara, 1952), from acetate in Hevea
hrasiliensis (Fournier et al., 1961) and avocado (Mudd and Stumpf, 1961),
from citrate in Aspergillus niger (Challenger et al., 1927), from succinate
in Aspergillus niger (Subramanian et al., 1929), from asparagine in rats
(Thomas and Stalder, 1959), from oxalacetate in pig heart extracts catalyzed
by metmyoglobin and Mn++ (Vennesland and Evans, 1944; Vennesland
et al., 1946), and from malonyl-CoA in Penicillium cyclopium, (Bentley and
Keil, 1961). The high concentrations of malonate in bush-bean plants led
Huffaker and Wallace (1961) to study the mechanism of the accumulation.
They found that the malonate synthesis is related to the dark COg fixation
in the roots, phosphoenolpyruvate being carboxylated to oxalacetate and
this going to malonate with the help of one or more enzymes. The addition
of phosphoenolpyruvate and Mg++ to root homogenate leads to the for-
mation of labeled malonate from 0^*02- It was also found that any other
reactions utilizing oxalacetate decrease the yield of malonate. It is very
interesting that frogs accumulate malonate-C^* from C^Oa, along with other
dicarboxylates (Cohen, 1963). In normal strains the malonate accounts for
only 0.3-0.5% of the total incorporation but in hybrids {R. pipiens X R.
sylvatica) the value is 6-23%. This increased accumulation in the hybrids
was attributed to some defect in the metabolism of malonate.
Methylmalonate can be formed from propionate (Flavin et al., 1955) in a
variety of tissues, and in rat liver the pathway has been shown to go through
succinate (Katz and Chaikoff, 1955). The feeding of isobutyrate and valine
to rats leads to the formation of methylmalonate (Thomas and Stalder,
1958) and the feeding of isoleucine leads to ethylmalonate (Stalder, 1959).
METABOLISM OF MALONATE 227
The first reaction appears to be an oxidative deamination. Since methyl-
malonyl-CoA is a common intermediate in many tissues, methylmalonate
probably arises from any substance forming the coenzyme A derivative.
(This will be discussed in greater detail in the following sections).
General Occurrence and Nature of Malonate Metabolism
The metabolism of malonate by many organisms and tissues has been
conclusively demonstrated by a variety of techniques. The ideal method is
the determination of C^^Og or other labeled products formed from labeled
malonate, but in some instances good evidence is provided by studies of
oxygen uptake, especially when the endogenous respiration is very small,
or by growth in media containing only malonate as a utilizable substrate.
In other cases, the evidence is more indirect. For example, a stimulation of
growth rate or a rise in respiration in the presence of other substrates may
suggest the utilization of malonate but does not prove it. In the tabulation
on page 228 of organisms in which malonate metabolism has been claimed
to occur, those that are probable and based on indirect evidence are des-
ignated by (P). It may be noted in addition that Shannon et al. (1959) found
malonate to be metabolized by the excised leaves of 15 different common
plants (such as fig, peach, eucalyptus, azalea, and lantana) and 30 other
plants representing 27 families, from which it must be concluded that plants
are generally capable of utilizing malonate.
The bacterial oxidation of malonate occasionally shows a lag period, first
observed by Lineweaver (1933). The rate of oxidation by Azotobacter is
very low for 2-3 hr, rises to a maximum around 6-8 hr, and falls off by
10 hr. The malonate is eventually 99% metabolized with an R.Q. of 1.6.
The theoretical R.Q. for complete oxidation:
CHaiCOOH), + 2 O2 -» 3 CO2 + 2 H,0
is 1.5. Lineweaver postulated that two separate reactions are involved: the
decarboxylation of malonate to acetate, and the oxidation of the acetate.
He attributed the lag phase to the slow decarboxylation, which was sup-
ported by the progressive decrease in the R.Q. with time. This interpreta-
tion was criticized by Karlsson (1950) because Lineweaver had not used
cells adapted to malonate. Malonate-grown Azotobacter does not decarbox-
ylate malonate appreciably under anaerobic conditions, so Karlsson con-
cluded that oxygen is required for the initial attack on malonate, either
because malonate must be oxidized before decarboxylation or because
oxygen may be required for some activation of malonate (for example, by
a phosphorylative mechanism). A lag phase was also demonstrated for
Aerobacter by Barron and Ghiretti (1953), the maximal oxidative rate oc-
curring around 2-3 hr after malonate addition. Only 40% of the malonate
228
1. MALONATE
Organisms and tissues
metabolizing malonate
Reference
Pseudomonas aeruginosa
Pseudomonas fluorescens
Escherichia coli (P)
Aerobacter aerogenes
Azotobacter agilis
Salmonella typhimurium (P)
Mycobacterium tuberculosis
Mycobacterium phlei
Aspergillus niger
Aspergillus (6 species) (P)
Penicillium cyclopium (P)
Penicillium (3 species) (P)
Streptomyces olivaceus
Pullularia pullulans (P)
Avena coleoptile (P)
Chlorella pyrenoidosa (P)
Pollen of Camellia, Thea, and Lilium
Tobacco leaves
Bush bean leaves
Peanut mitochondria
Euglena gracilis
Hymenolepis diminuta (cestode) (P)
Locusta migratoria fat body
Chickens (P)
Pigeon liver
Mice
Rats
Rat liver
Rat ventricle
Rabbits
Dogs
Dog heart and muscle
Human placenta
Human prostate
Gray (1952)
Hayaishi (1953, 1954, 1955 a), Wolfe
and Rittenberg (1954), Wolfe et al.
(1954 a, b, 1955)
Grey (1924), Quastel and Whetham
(1925), Cook (1930)
Barron and Ghiretti (1953)
Lineweaver (1933), Karlsson (1950)
Berry and Beuzeville (1960)
Hayaishi and Kornberg (1952),
Bernheim et al. (1953), Horio and
Okunuki (1954), Kusunose et al.
(1960)
Miiller et al. (1960)
Walker et al. (1927), Challenger et al.
(1927)
Berk et al. (1957)
Bentley and Keil (1961)
Berk et al, (1957)
Maitra and Roy (1961)
Clark and Wallace (1958)
Albaum and Eichel (1943)
Eny (1951)
Okunuki (1939)
Vickery (1959), Vickery and Palmer
(1957)
Young and Shannon (1959)
Giovanelli and Stumpf (1957)
Danforth (1953)
Read (1956)
Tietz (1961)
Clementi (1929), Pupilh (1930)
Menon et al. (1960)
Lifson and Stolen (1950)
Lee and Lifson (1951), Busch and
Potter (1952a), Nakada etal. (1957),
Thomas and Stalder (1959)
Menon el at. (1960)
Rice and Berman (1961)
Wick et al (1956)
Pohl (1896)
Menon et al. (1960)
Hosoya and Kawada (1958), Hosoya
et al. (1960)
Andrews and Taylor (1955)
METABOLISM OF MALONATE
229
is oxidized but the R.Q. is 1.47, close to that for complete oxidation to
CO2 and water. Again, no decarboxylation is observed in nitrogen. Horio
and Okunuki (1954) reported a 30-60 min lag period for Mycobacterium and
also found that decarboxylation presumably precedes oxidation, because
CO2 formation always is ahead of Oo uptake, and acetate can be demon-
strated in the culture after 2 hr. The explanation of this behavior is found
in the work of Gray (1952) on Pseudomonas. Unadapted cells show a lag
period of 2 hr whereas cells cultured for 70 hr in 22 mM malonate are able
to oxidize malonate immediately (Fig. 1-21). It was postulated that mal-
Time ( hours)
Fig. 1-21. Oxidation of malonate by normal and
adapted Pseudomonas aeruginosa. (From Gray, 1952.)
onate decarboxylase is an adaptive enzyme, and it was pointed out that
the resistance of certain microorganisms to malonate may be due to such
an enzyme as well as to permeability barriers. Horio and Okunuki showed
that streptomycin does not directly inhibit the decarboxylation of mal-
onate or the oxidation of acetate, but inhibits malonate oxidation in un-
adapted cells, probably by preventing the synthesis of the decarboxylase.
The oxygen requirement may also relate to the adaptive enzyme syn-
thesis.
230
1. MALONATE
Pathways of Malonate Metabolism in Microorganisms
An analysis of the pathways of malonate oxidation was made simultane-
ously in the laboratories of Hayaishi and Rittenberg between 1953 and 1955.
The work was done on Pseudomonas fluorescens, a soil isolate capable of
utilizing malonate as the sole carbon source, and adapted to malonate by
culture in 27-33 nvM malonate media. Hayaishi (1953) observed that the
decarboxylation of malonate requires ATP and CoA and postulated that
malonate must first be activated, probably to malonyl-CoA. Using partially
purified extracts from Pseudomonas, it was shown that malonate is quanti-
tatively converted to COg and acetate, no other products being detectable
chromatographically. The proposed scheme (1-7) may be represented as
follows (Hayaishi, 1954, 1955 a):
Malonate
Acetate
Malonate
(1-7)
Acetyl — CoA
The cyclic process thus involves the transfer of CoA back and forth be-
tween the acetyl and malonyl groups. Wolfe et at. (1954) also ruled out the
direct decarboxylation to acetate by showing a dependence on ATP and
CoA, and in later work (Wolfe et al., 1954 b, 1955; Wolfe and Rittenberg,
1954) proposed the following scheme (1-8) based mainly on chromatographic
analyses of intermediates and products:
Acetate + Malonyl-diCoA-
Malonate
ATP + CoA
t
Malonyl - CoA
Acetyl - CoA
>■
(1-8)
The principal difference between the two schemes is the participation of
malonyl-diCoA. Hayaishi's results do not exlude it but provide no evidence
for it, while Wolfe et al. claim to have detected it chromatographically.
Although the decarboxylation of malonate is characterized by AF = — 7
kcal/mole, the activation energy is presumably so high that the more com-
plex mechanisms above are necessary. The malonate decarboxylase and
METABOLISM OF MALONATE 231
CoA-transferase have been partially purified. The acetyl-CoA formed from
malonate may enter the cycle directly or participate in other reactions, such
as the formation of acetoacetate if the cycle is blocked by malonate, or
transfer its CoA to malonate, or simply be hydrolyzed to acetate. It is very
interesting that a lag period was noticed in the oxidation of malonate by
cell-free extracts (Wolfe et al., 1955). Little oxygen uptake occurs with
malonate until it is all decarboxylated; during the period of the most rapid
CO2 evolution, the respiration is low. Yet acetate is activated and oxidized
rapidly. One possible explanation is a block of the cycle by malonate so
that acetyl-CoA cannot enter until most of the malonate has been me-
tabolized. Another explanation involves a distribution of CoA in favor of
the malonyl derivatives with little acetyl-CoA present during the active
decarboxylation reaction.
Cryptococcus terricolus can grow on malonate as the sole source of carbon
and malonate stimulates the endogenous respiration and CO2 formation after
a lag phase (Pedersen, 1963). It would appear that malonate is completely
oxidized to CO2 and water, since the R.Q. in the presence of malonate is
1.54. In other microorganisms where the oxidation is not complete, mal-
onate may be incorporated into a variety of substances, especially lipids
(Bu'Lock et al., 1962), although the rate of incorporation is seldom very
rapid.
Pathways of Malonate Metabolism In Plants and Animals
Despite the widespread occurrence of malonate metabolism in plant tis-
sues, little is known of the reactions involved, although perhaps they are
not significantly different from those described for microorganisms. Peanut
mitochondria supplemented with ATP, CoA, and other factors are able to
oxidize malonate (Giovanelli and Stumpf, 1957). Incubation with malo-
nate-2-C^* for 2 hr leads to the appearance of labeled citrate, malate, and
succinate, indicating the sequence
Malonate ->■ malonyl-CoA -> acetyl-CoA -> CO2 + HjO
the last step occurring in the cycle. The participation of malonyl-CoA in
the oxidation of propionate by peanut mitochondria is suggested by the
tracing of the label from propionate- 1-C^* (Giovanelli and Stumpf, 1958).
The following sequence involving malonic semialdehyde may be formulated:
Propionate -> propionyl-CoA ->■ acrylyl-CoA -> |3-hydroxypropionyl-CoA ->
^-hydroxypropionate -> malonic semialdehyde -> malonyl-CoA ->
acetyl-CoA -> CO^ + H2O
It is also possible that CoA derivatives are retained throughout, since a
malonic semialdehyde-CoA dehydrogenase which catalyzes the formation
232 1. MALONATE
of malonyl-CoA has been found in Clostridium kluyveri (Vagelos, 1960).
In bush bean {Phaseolus vulgaris) leaves, incubation with malonate-2-C^*
leads to labeled citrate, isocitrate, and malate, indicating a pathway through
acetyl-CoA and the cycle (Young and Shannon, 1959). Malonate is incor-
porated in isolated spinach chloroplasts at about half the rate for acetate,
much of the label appearing in lipids (Mudd and McManus, 1964).
Metabolism of malonate was first described in the dog by Pohl (1896),
who found that only a fraction of the malonate administered to the animals
can be recovered in the urine. This subject was not taken up again until
1950 and since that time much has been learned of how the body deals
with malonate, and of the role of malonate and its derivatives in normal
metabolism. It would appear from the limited data that about 30% of
the administered malonate is metabolized (Lifson and Stolen, 1950; Busch
and Potter, 1952 a). However, the rate of oxidation is relatively slow and
in rabbits represents less than 2% of the total respiration (Wick et al.,
1956). The rate of oxidation may be in part limited by the transfer from
the extracellular space into the tissues, since this is slow.
Various mammalian tissues can decarboxylate malonate and utilize the
acetate formed. This has been investigated most completely in rat tissue
slices by Nakada et al. (1957), who determined the 0^*0, arising from mal-
onate-1-C^^ during 1 hr incubation at pH 7.4, the total concentration of
malonate being 5 xnM (see accompanying tabulation). Kidney, liver, and
Tissue Added C^* as Qi^Oa (%)
Kidney 27.0
Liver 18.0
Heart 15.2
Diaphragm 6 . 8
Spleen 1 . 7
Brain 1 . 5
Lung 1 . 0
Testis 0.6
heart are particularly active, and the tissues show a wide range of decar-
boxylative ability, part of which may be due to different rates of penetra-
tion. The variation of malonate oxidation with concentration is shown in
Fig. 1-22 for rat kidney slices, and an inhibition of its own metabolism is
seen at concentrations above 5 vaM. The inhibition of acetate- 1-C^* oxida-
tion is shown for comparison. The relative rates of activation of malonate,
succinate, and glutarate by several tissues were determined by measuring
the rates of formation of hydroxamic acid from hydroxylamine during the
METABOLISM OF MALONATE
233
incubations (see accompanying tabulation) (Menon et al., 1960). The results
show not only differences between the tissues, but also that the activating
system for malonate is different from that for succinate and glutarate.
Tissue
Hydroxamic
acid formed (m/<moles/mg/30 min)
Malonate
Succinate
Glutarate
Dog heart
10.3
7.4
28.4
Dog muscle
2.6
8.2
16.0
Pigeon liver
4.7
20.5
31.7
Rat liver
11.5
62.9
86.7
The metabolic pathways for malonate in mammalian tissues appear to
be very similar to those in bacteria. The oxidation of malonate requires
ATP, coenzyme A, and Mg++ in extracts or homogenates of rat kidney
(Nakada et al., 1957), human placenta (Hosoya and Kawada, 1958), and,
incidentally, locust fat body (Tietz, 1961). However, the kinases, decarbox-
FiG. 1-22. Effects of malonate concentration on the oxidation
of malonate and acetate by rat kidney slices at pH 7.4 and
37° during 2-hr incubation. (From Nakada et al., 1957.)
ylases, and CoA-transferases for these reactions have not been studied.
An important transfer of coenzyme A between acetoacetyl-CoA and mal-
onate has been shown to occur in pig heart extracts (Beinert and Stansly,
1953). Acetoacetyl-CoA is formed by the condensation of two acetyl-CoA
234 1. MALONATE
molecules, and various carboxylate anions can accept the coenzyme A so
that acetoacetate is formed:
2 Acetyl-CoA :^ acetoacetyl-CoA + CoA
Acetoacetyl-CoA -|- malonate :;^ malonyl-CoA + acetoacetate
succinate and butyrate being normally the most active. In this way mal-
onate can give rise to acetoacetate as well as by its block of the cycle.
Labeled malonate forms labeled acetoacetate in rat liver (Nakada et al.,
1957). This type of reaction has also been shown to occur in yeast, and dog
heart and skeletal muscle (Menon and Stern, 1960). The enzyme, succinyl-
/3-ketoacyl-CoA transferase, was purified from pig heart and catalyzes the
transfer to both malonate and glutarate. The following reaction:
Succinyl-CoA + malonate :^ malonyl-CoA + succinate
also occurs. It is interesting to speculate that some of the succinyl-CoA
formed in the oxidation of a-ketoglutarate transfers its coenzyme A to
malonate when it is present; if the malonyl-CoA is not readily metabolized,
this could deplete the a-ketoglutarate oxidase of coenzyme A and slow down
the reaction. Condensation of malonyl-CoA with coenzyme A derivatives
may be important in fatty acid synthesis. In pigeon liver and carrot roots
there is an enzyme catalyzing the condensation of malonyl-CoA with either
acetyl-CoA or butyryl-CoA; although the product is unknown, it was isolat-
ed chromatographically (Steberl et al., 1960). This product can form pal-
mitate with other enzyme fractions. When malonyl-2-C^^-CoA is incubated
with extracts from various rat tissues, various Cig-Cig acids are formed,
depending on the acyl-CoA acceptor used (Horning et al., 1960). One acyl-
CoA unit is incorporated into long-chain fatty acids and the rest of the
C-chain in supplied from malonyl-CoA. Labeled fatty acids are also formed
from malonate- 1-C^* in particle suspensions of the locust fat body (Tietz,
1961). A highly purified preparation from pigeon liver, which converts
malonyl-CoA and acetyl-CoA to palmitate in the presence of NADPH, has
been reported (Bressler and Wakil, 1961). In the absence of NADPH, mal-
onyl-CoA and acetyl-CoA condense to form an unknown product (which
is not acetoacetate, butyrate, or /^-hydroxybutyrate). The conversion of
malonyl-CoA to fatty acids is perhaps mediated through such condensa-
tions to Cg acids, forming butyryl-CoA, which would again condense with
malonyl-CoA, and so lengthen the chain.
A few words should be said about the pathways of methylmalonate me-
tabolism since it has been recently found to be an important intermediate
in fatty acid |metabolism, and malonate can interfere markedly with at
least one of these reactions. Methylmalonate was shown to be an interme-
diate in the metabolism of propionate in various tissues (Flavin et al., 1955;
Katz and Chaikoff, 1955; Feller and Feist, 1957). In the course of this work
INHIBITORS STRUCTURALLY RELATED TO MALONATE 235
a novel reaction was discovered, namely, the interconversion of methyl-
malonate and succinate. The enzyme, which has been called a methyl-
malonyl-CoA isomerase, catalyzes the reaction:
Methylmalonyl-CoA :^ succinyl-CoA
and has been purified from ox liver (Stern and Freidman, 1960) and Pro-
fionihacterium shermanii (Wood and Stjernhohn, 1961; Stjernholm and
Wood, 1961). At equilibrium the ratio (succinyl-CoA)/(methylmalonyl-CoA)
is 10.5 (pH 7 and 25°). This reaction functions in both the oxidation and
formation of propionate. Another reaction of importance in propionate
metabolism is
Methylmalonyl-CoA + pyruvate :^ propionyl-CoA + oxalacetate
It is catalyzed by methylmalonyl-oxalacetate transcarboxylase, whereby a
carboxylation may be effected without the intervention of COg or the ex-
penditure of energy to activate COg. It may finally be noted that the feed-
ing of malonate to dogs leads to a marked excretion of methylmalonate in
the urine (Thomas and Stalder, 1958). This does not necessarily mean that
the methylmalonate is formed from malonate, since malonate inhibits the
interconversion of methylmalonyl-CoA and succinyl-CoA very strongly
(Flavin et al., 1955).
INHIBITORS STRUCTURALLY RELATED TO MALONATE
The inhibition of succinate dehydrogenase by various dicarboxylate ions
was treated earlier (page 34). Maleate will be dealt with in Chapter III-2
and oxalate will be discussed in Volume IV. Inhibitors related to malonate
will also be found in the following chapter on analogs. It then remains to
take up the esters of malonate, hydroxy malonate, and those compounds
in which the carboxylate groups have been replaced by other anionic groups.
As mentioned previously, there is often confusion in the nomenclature; we
shall adopt the following (using the ethyl derivatives as examples):
COO" Et^ /COO"
Et— Hc:" . /C^ _
coo Et COO
Ethylmalonate Diethylmalonate
/COO /COO— Et
HgC Hz^s,
^COO— Et ^COO — Et
Malonic Malonic
monoethyl ester diethyl ester
236 1. MALONATE
Malonic Esters
Malonic monoethyl and diethyl esters have been found to have some in-
teresting actions. They can increase the survival period of mice infected
with mycobacteria (Davies et al., 1956), are occasionally carcinostatic (Freed-
lander et al., 1956), can inhibit the breakdown of hexobarbital in liver homo-
genates and prolong the narcotic action (Kramer and Arrigoni-Martelli,
1960), and inhibit sporulation of Bacillus cereus (Nakata and Halvorson,
1960). However, the relation of these effects to succinate dehydrogenase
inhibition is obscure. In most cases, malonic esters have been used to cir-
cumvent the permeability barriers to malonate, inasmuch as the esters
should penetrate into cells readily. It has often been assumed that hydrolysis
to malonate occurs within the cells. This hydrolysis must be enzymatic
because the esters are quite stable. A lipase from pig liver hydrolyzes one
ethyl group from malonic diethyl ester but does not remove the other
ethyl group, the product being malonic monoethyl ester (Christman and
Lewis, 1921). I have been able to find no direct evidence for the hydrolysis
to malonate. Malonic esters are neutral at physiological pH's, since they
are very weak acids with pK„ values around 15.75 (Rumpf et al., 1955)
and ionize very slowly with a rate constant of 1.8 X 10~^ min~^ (Pearson
and Mills, 1950).
The effects of the malonic esters on metabolism will now be discussed in
order to determine if there is any indirect evidence for the intracellular
hydrolysis to malonate and the inhibition of succinate dehydrogenase.
When injected into fluoroacetate-poisoned rats, malonate and malonic di-
ethyl ester have approximately the same effects on the accumulation of
citrate in the heart and kidneys (Fawaz and Fawaz, 1954). Furthermore, in
kidney slices, the diethyl ester inhibits succinate oxidation 92% at 20 mM,
and malonate at the same concentrazion inhibits 75%. Less inhibition by
the ester compared to malonate is observed in heart slices. Evidence for
hydrolysis by both tissues was adduced from the decreases in the pH ob-
served. The respiration of the fungus Zygorrhynchus moelleri is not inhibited
readily by malonate although the succinate dehydrogenase from this orga-
nisms is quite sensitive, indicating a failure to penetrate (Moses, 1955).
Malonic diethyl ester was tested and found to inhibit the oxidation of both
glucose and acetate, but at high concentrations (see tabulation). It was felt
Malonic diethyl
ester
% Inhibition of:
{mM)
Glucose oxidation
Acetate oxidation
10
30
100
Stim 49
30
96
10
76
98
INHIBITORS STRUCTURALLY RELATED TO MALONATE 237
that the greater inhibition of acetate oxidation is evidence for the hydro-
lysis of the ester and that the inhibition is exerted by malonate. Malonic
diethyl ester was also used to facilitate penetration into Penicillium chryso-
genum, since even 100 roM malonate does not effect acetate metabolism
(Goldschmidt et al., 1956). The ester at 20 milf inibits the production of
C^*02 from labeled acetate 75-85% and simultaneously decreases the in-
corporation of C^* into cellular materials, the labeling of glutamate being
particularly depressed. The utilization of acetate by Bacillus cereus is also
interfered with by malonic diethyl ester, so that acetate and p>Tuvate ac-
cumulate (Nakata and Halvorson, 1960). Malonate and the diethyl ester
have been compared with respect to their effects on the respiration of
Mycobacterium pJdei (Miiller et al., 1960). Malonate stimulates the endoge-
nous respiration, presumably through its oxidation, and the ester stimulates
even more potently at 1-10 raM, although at 100 mM the ester inhibits
and malonate still stimulates. The respiration with glycerol as the substrate
behaves similarly. Finally, malonic diethyl ester markedly stimulates the
endogenous respiration of Chlorella vulgaris at 4-10 mM, but inhibits the
oxidation of glucose and acetate, the latter more strongly (Merrett and
Syrett, 1960). All of these results show that the diethyl ester is inhibitory
but certainly do not constitute conclusive evidence for a hydrolysis to mal-
onate. This is a subject that should be pursued further and more extensive
tests should be made for enzymes hydrolyzing the esters. Until the intra-
cellular hydrolysis can be established, the results obtained with the malonic
esters cannot be interpreted.
Hydroxy malonate (Tartronate)
The substitution of a methylene hydrogen of malonate by any group
seems to reduce rather strongly the ability to inhibit succinate dehydrogen-
ase. Even the small hydroxyl group almost abolishes the inhibitory activity
and this is evidence that the binding of malonate to the active center of
succinate dehydrogenase must involve severe steric restrictions. Although
tartronate does not inhibit succinate dehydrogenase, it has other actions
of some interest. Quastel and Wooldridge (1928) found that 71.4 roM tar-
tronate does not inhibit succinate oxidation at all whereas it inhibits the
oxidation of lactate 90%, as measured by methylene blue reduction in tol-
uene-treated E. coli. In fact, the marked differences in the effects of mal-
onate and tartronate led Quastel and Wooldridge to postulate the specific
structure of the active centers of enzymes. Tartronate inhibits the respira-
tion of rat liver slices and is strongly ketogenic, acetoacetate being formed
to a greater extent that with equivalent concentrations of malonate (Ed-
son, 1936). Pig heart malate dehydrogenase is inhibited 24% by 60 mM
tartronate (Green, 1936) but in pigeon liver extracts it is about 1000 times
as effective, inhibiting malate oxidation competitively with a ^^ of 0.09 mM
238 1. MALONATE
(Scholefield, 1955). Tartronate is also a competitive inhibitor of the decar-
boxylating malate dehydrogenase (malic enzyme) of pigeon liver with a K^
of 0.1 mM, the inhibition being stronger than with malonate (Stickland,
1959 b). The carboxylation of pyruvate, catalyzed by the same enzyme,
is also strongly inhibited, but there is a large noncompetitive element
(Stickland, 1959 a). The inhibition of lactate and malate oxidations and
decarboxylations is not surprising since tartronate is structurally similar.
Although tartronate occurs in plant and animal tissues to the extent of
8-15 //g/g wet weight of tissue, which would correspond to about 0.1 xnM
in the tissue water (Veitch and Brierley, 1962), it is doubtful if it could
exert a regulatory action on the metabolism. Tartronate is not metabolized
in the rat and consequently is near 8 raM in the urine.
The respiration of guinea pig brain slices is depressed by tartronate at
concentrations of 67-75 rcvM (Jowett and Quastel, 1937). The degree of inhi-
bition depends on the substrate provided and is maximal with lactate and
minimal with pyruvate. Since lactate is probably metabolized through
pyruvate, the inhibition here may be mainly on lactate dehydrogenase.
However, the anaerobic breakdown of pyruvate and anaerobic glycolysis
are also well inhibited. The respiration of Mycohacterium. phlei with lactate
as substrate is inhibited 65% with 66 mM tartronate, although the endo-
genous respiration is stimulated and the oxidation of glucose unaffected
(Edson and Hunter, 1947). This relatively specific effect on lactate dehydro-
genase was used by Fiume (1960) to inhibit aerobic glycolysis in tumor cells,
inasmuch as lactate dehydrogenase is involved. It was postulated that
aerobic glycolysis might be inhibited more strongly in the tumor than in
normal tissues, depleting the ATP supply more severely. It was found that
tartronate inhibits aerobic glycolysis of the Yoshida ascites hepatoma —
26% at 10 mM, 34% at 20 mM, and 58% at 50 mM - but comparisons
with normal tissue were not made.
The inhibition of phosphatases should perhaps also be considered in
work with tartronate, since prostatic acid phosphatase is inhibited com-
petitively with a K^ of around 50 mM. The inhibition by tartronate is
much greater than by malonate and about twice as potent as by ketomal-
onate (Kilsheimer and Axelrod, 1957).
Aminomalonate
This substance was considered to be a possible substrate for the synthesis
of S-aminolevulinate but was found upon examination to be a potent inhib-
itor of S-aminolevulinate synthetase (Matthew and Neuberger, 1963). The
inhibition of the enzyme from Rhodopseiidomonas spheroides and chicken
erythrocytes is competitive; the K, for the bacterial enzyme is 0.0225 mM.
Pyridoxal-P is a coenzyme in these systems and the inhibition depends on
its concentration. Aminomalonate may be considered to be an analog of
INHIBITORS STRUCTURALLY RELATED TO MALONATE 239
glycine (it is carboxyglycine) and presumably inhibits 8-aminolevulinate
synthetase by binding to the glycine site and complexing with pyridoxal-P
as well. Aminomalonate condenses with aldehydes nonenzymatically in
the presence of pyridoxal-P. Furthermore, other pyridoxal-P-dependent
enzymes are inhibited, e.g. serine hydroxy methyltransferase, whereas en-
zymes involved in glycine metabolism but not requiring pyridoxal-P are not
inhibited. Aminomalonate can be formed in the tissues by transamination
between ketomalonate and glutamate, and can be decarboxylated by an
enzyme found in silkworm larvae and rat heart and liver to glycine. This
derivative of malonate is a good illustration of how a simple change in the
structure can create an inhibitor with quite different properties and inhib-
itory spectrum.
Substituted Malonates
Although the alkylmalonates are not particularly interesting as inhibi-
tors, there are two malonate derivatives that may warrant further investi-
gation. Fluoromalonate was studied by Chari-Bitron (1961) on the principle
that if malonate is metabolized through acetyl-CoA, fluoromalonate might
follow the same pathway and enter the cycle as fluoroacetyl-CoA, produc-
ing the same effects as fluoroacetate, namely, a block of the cycle at the
aconitase step. The toxicity of fluoromalonate is a good deal less than
fluoroacetate but the ester is as toxic in mice (see accompanying tabulation).
LD50 (mg/kg)
Animal
Fluoromalonate Fluoromalonic diethyl ester Fluoracetate
Mouse
80
Rat
60
Guinea pig
2
15 15
70 5
— 0.25
Death is associated with a marked accumulation of citrate in the tissues
and it differs strongly from malonate in this respect. Accumulation of cit-
rate also occurs in kidney mitochondria with fluoromalonate at concentra-
tions around 1 mM. It was further established that decarboxylation of
fluoromalonate occurs in kidney preparations. Finally, fluoromalonate is
only about one tenth as effective as malonate in the inhibition of succinate
dehydrogenase. The results thus conform quite well to the predicted mech-
anism. Difluoromalonate and its amide inhibit quite readily the oxida-
tions of succinate and fumarate by Pseudomoyias (around 70% inhibition
at 0.7 mM), but there is no inhibition of succinate dehydrogenase in soni-
cates; the mechanism is unknown (Bernheim, 1963).
240 1. MALONATE
Inasmuch as the active center of succinate dehydrogenase possesses a
sulfhydryl group close to the cationic binding sites, this being the basis
for the inhibition by mercurials and other sulfhydryl agents, the possibility
of combining a sulfhydryl inhibition with malonate presents itself. Mercuri-
malonamide and mercurimalonic diethyl ester have been prepared (Naik
and Patel, 1932) and these compounds, or particularly the hydrolyzed
^COO-Et
^C(X>-Et
forms if they are stable, might be interesting to examine as inhibitors of
succinate dehydrogenase.
Acetylene-dicarboxylate and Propane-tricarboxylate
Succinate dehydrogenase is inhibited competitively by acetylene-dicar-
boxylate (Dietrich et at., 1952). The order of addition of the succinate and
acetylene-dicarboxylate is important; for example, if acetylene-dicarboxy-
late is added 20 min after the succinate, the rate of oxygen uptake de-
creases slowly and does not become equal to that observed when the suc-
cinate is added after the inhibitor until 5 hr (Thomson, 1959). Acetylene-
dicarboxylate is about as effective as malonate on long incubation with the
enzyme. The constants obtained on rat kidney succinate dehydrogenase
are ir,„ = 4.12 mM and K^ = 0.81 roM when substrate and inhibitor are
added together; after 18 hr incubation with the inhibitor, iC, =^ 0.171 vaM.
It is difficult to understand why the rate of inhibition is so slow. Succinate
dehydrogenase from pig heart is less readily inhibited, K^ being 1.4 mM
with A'^-methylphenazine as electron acceptor and 16.5 when ferricyanide is
the acceptor (Hellerman et al., 1960). Possibly insufficient time for equilib-
rium was allowed. The inhibitions of succinate and pyruvate oxidations
by acetylene-dicarboxylate in suspensions of rate heart mitochondria are
shown in Fig. 1-23 (Montgomery and Webb, 1956 b). Acetylene-dicarboxy-
late is less potent than malonate against succinate oxidation and more po-
tent against pyruvate oxidation, indicating that an inhibition is exerted
at some other point in the cycle.
Propane-tricarboxylate is a rather weak inhibitor of succinate dehydro-
genase from rat heart, 5 mM inhibiting around 30% when the succinate is
also 5 mM. The oxidation of a-ketoglutarate is inhibited 60% under the
same conditions, suggesting some effect on the a-ketoglutarate oxidase.
The oxidation of pyruvate is inhibited even more strongly (Fig. 1-24). It
might be thought that propane-tricarboxylate would inhibit aconitase or
isocitrate dehydrogenase, but very little inhibition is noted, either of cit-
rate oxidation or the ability of citrate to function as a source of oxalac-
etate for the oxidation of pyruvate.
INHIBITORS STRUCTURALLY RELATED TO MALONATE
241
Traws-l,2-Cyclopentane-dicarboxylate inhibits the succinate dehydroge-
nase of Tetrahymena geleii homogenates, around 50% inhibition being ob-
served when the inhibitor/substrate ratio is 0.5 (Seaman and Houlihan,
1950). On the other hand, in intact cells this substance increases the utili-
zation of pyruvate, acetate, and succinate. The uptake of acetate is in-
80
-
Pyruvote + Malate
^^
%60
INH
-
/
40
■
/
20
-
1 1 i
Succinate
I 5 10 50
(Acetylene - Dicarboxylate)
Fig. 1-23. Effects of acetylene-dicarboxylate (in milf)
on the oxidations of p>Tuvate -f malate and succinate
by rat heart mitochondria. (From Montgomery and
Webb, 1956 b.)
creased as much as 50% by ^ra??5-l,2-cyclopentane-dicarboxylate and in
its presence succinate is taken up and oxidized whereas succinate does not
enter the cells normally. It was postulated that trans-1, 2 cyclopex\':ine-
dicarboxylate increases the permeability of the membrane to these sub-
strates, possibly by an action on the metabolic systems involved in the
)5 I 2 5
( Propone - Tncarboxylote)
Fig. 1-24. Effect of propane -tricarboxylate (in
inM) on the oxidation of pyruvate + malate by
rat heart mitochondria. (From Montgomery and
Webb, 1956 b.)
242 1. MALONATE
inward transport. It is not known if such an effect is observed with other
dicarboxylate anions.
Sulfonate, Phosphonate, and Arsenate Analogs of Succinate
Succinate and malonate are bound to the active center of succinate de-
hydrogenase in part through electrostatic forces involving the negatively
charged carboxylate groups. It might be expected that substances in which
the carboxylate groups are replaced by other anionic groups would also be
inhibitory. The structures for some of the compounds and the intercharge
distances are given in Table 1-1. Klotz and Tietze (1947) first demonstrated
the inhibition of succinate dehydrogenase (rat liver) by 1,2-ethanedisulfo-
na'te and /5-sulfopropionate, 50% inhibition being observed in both cases
when the inhibitor is 13 mM and succinate is 20 mM, these inhibitions
being approximately equivalent to those of malonate. Intact E. coli cells
are not affected by 1,2-ethanedisuIfonate but the oxidation of succinate
by cell-free preparations is well inhibited, corresponding to the results with
malonate (Ajl and Werkman, 1948). Some inhibitor constants for these
substances are shown in Table 1-29.
There appears to be definite differences in the susceptibility of the suc-
cinate dehydrogenases from different sources. Seaman (1952) noted that
/?-phosphonopropionate inhibits the Tetrahymena enzyme more strongly
than does malonate, and yet no inhibition is observed on the enzymes
from rat heart, liver, brain, or muscle. The inhibitions are competitive
wherever they have been tested and there is no evidence that the mecha-
nism is in any way different from that of malonate. Since these groups are
larger than the carboxylate group, the interchange distances are greater
than in malonate or succinate, and this must play some role in determining
their binding to the enzyme. However, the distances for the malonate ana-
logs, methanedisuffonate and arsonoacetate, are between those for malon-
ate and succinate, so that binding should be as tight as for these latter
substances if this were the only factor. Another factor of importance is the
total net charge on these inhibitors, inasmuch as each of the sulfonate,
phosphonate, and arsonate groups can ionize more than once. In other
words, a disulfonate can exist in five different forms with charges 0,-1,
— 2, —3, and —4. Furthermore, the third and fourth ionization constants
are in the physiological range of pH (see accompanying tabulation). The
vK„ vK„
Arsonoacetate
7.7
—
1 ,2-Ethanediphosphonate
6.84
8.17
1 ,4-Butanediphosphonate
7.28
9.05
Pyrophosphate
5.69
7.76
INHIBITORS STRUCTURALLY RELATED TO MALONATE
243
Table 1-29
Inhibitor Constants for Sulfonate, Phosphonate, and Arsonate Analogs of
Substrates "
Inhibitor
Preparation
K,
Reference
Malonate
Tetrahymena succinate
6.67
Seaman (1952)
/S-Phosphonopropionate
dehydrogenase
1.51
Arsonoacetate
9.25
1,2-Ethanedisulfonate
Rat liver succinate
2.73
Klotz and Tietze
/?- Sulfopropionate
dehydrogenase
3.69
(1947)
Malonate
Mouse liver succinate
0.19
Tietze and Klotz
1 ,2-Ethanedisulfonate
dehydrogenase
26.5
(19.52)
o-Sulfobenzoate
11.7
/5-Phosphonopropionate
No inh
Arsonoacetate
No inh
Methionate
20.6
Malonate
Beef heart succinate
0.014
Rosen and Klotz
Methanediphosphonate
dehydrogenase
0.5
(1957)
Phosphonoacetate
0.15
Pyrophosphate
0.0011
Hs^pophosphate
0.14
1,2-Ethanediphosphonate
9.6
1,4-Butanediphosphonate
8.9
Arsonoacetate
15.3
Malonate
Fumarase
40
Massey (1953b)
a-Hydroxy-/3-sulfo-
propionate
16.5
" The A', values for rat and mouse liver succinate dehydrogenases were cal-
culated assuming A'^ = 6 X 10~^ M, and for beef heart succinate dehydrogenase
K^ = 4 X 10"* M. Although these values are undoubtedly inaccurate, the Aj values
so calculated are useful for comparisons since within each experiment the relative
values are reliable.
rather poor inhibition produced by the arsono and phosphono derivatives
was postulated by Tietze and Klotz (1952) as possibly due to the extra
negative charge carried bj^ these substances. In the more recent work of
Rosen and Klotz (1957), the ionization was taken into account and the
inhibitor constants calculated on the basis of the concentration of doubly
charged anion present. Evidence that too high a charge reduces the inhib-
itory potency is provided by the falling off of the inhibition as the pH is
raised. On the other hand, it is difficult to understand why there should
244 1- MALONATE
be less affinity between cationic groups on the enzyme and doubly charged
anions, compared to singly charged anions, and one cannot help but wonder
if the alterations in pH in the experiments of Kosen and Klotz were not
affecting the ionizable groups on the enzyme. These workers suggested that
the binding of these substances involves an iron ion on the enzyme and
correlated affinities with the ability to chelate iron. This is certainly a type
of binding that should be kept in mind, but it is difficult to reconcile with
the fact that iron-chelating agents such as 1,10-phenanthroline and 2,2'-bi-
pyridine do not interfere with the binding of succinate to the dehydrogenase
even though their attachment to the enzyme can be demonstrated spectro-
scopically.
An interesting succinate dehydrogenase inhibitor, not fundamentally re-
lated to the compounds previously discussed, is 3-nitropropionate (hiptage-
nate), found to be the toxic principle of Indigofera endecaphylla (Morris
et al., 1954). It inhibits succinate dehydrogenase competitively with K^ ==
0.19 n\M (Hylin and Matsumoto, 1964,) although no inhibition of the
enzyme is found after administration of the substance to animals, so that
it is not possible to correlate the toxicity with an effect on this enzyme. It
is rather surprising that 3-nitropropionate binds so well to succinate de-
hydrogenase, lacking two anionic groups, and it would be interesting to
know more of the nature of the interaction of the nitro group with the
enzyme.
With respect to inhibitors related to malonate in one way or another,
it must be concluded that none possesses particular advantages over mal-
onate for the specific inhibition of succinate dehydrogenase, although cer-
tain derivatives have interesting properties and metabolic actions. Progress
could be made by the finding of forms of malonate, or related inhibitors,
that are uncharged, reasonably stable outside the cell, and easily split
to the active inhibitor intracellularly. It would also be valuable to have an
inhibitor which would initially bind to the succinate dehydrogenase at the
substrate site, because of its complementary configuration, and then react
chemically with an adjacent group, so that the inhibition of this enzyme
would be not only specific but slowly reversible.
CHAPTER 2
ANALOGS OF ENZYME REACTION
COMPONENTS
An enzyme-catalyzed reaction involves the combination of the com-
ponents with specific sites on the apoenzyme protein surface, these areas
possessing the particular molecular configuration and the electrical field
distribution required for the attachment and the electronic displacements
characterizing the activated complex. If one of these components is modified
in any way, its behavior in the system will usually be altered, due primarily
to the new pattern of interaction between the modified component and
the enzyme. The development and study of such analogs of substrates and
coenzymes have been very active fields during the past few years for several
reasons. First, the determination of the relative aflfinities of analogs that
are substrates or inhibitors for enzymes is one of the most effective means
for analyzing the topography of active centers and establishing the types
of interaction involved in the catalysis. Second, it is hoped that inhibitors
more specific for blocking certain enzymes than the inhibitors previously
available will be found and this has been justified to a certain extent.
Third, it has been realized that analog inhibition has direct bearing on the
important phenomenon of feedback control of metabolic sequences and on
the general regulation of cellular metabolism. Last, it is anticipated that
the use of proper analogs may be useful in the specific correction of certain
abnormal metabolic patterns and growth processes, such as occur in here-
ditary enzyme defects or neoplastic changes.
The previous chapter is concerned with malonate, a classic analog inhib-
itor, and in the present chapter it is proposed to extend this principle to
a variety of enzymes in order to establish some basic concepts. There are
a number of inhibitors which act, at least occasionally, because they are
structurally related to some enzyme reaction component, but which for
various reasons will be discussed in separate chapters. Such are carbon
monoxide, fluoroacetate and fluorocitrate, parapyruvate, arsenate, pyro-
phosphate, monoamine oxidase inhibitors, certain inhibitors of cholinester-
ase, and various drugs. Furthermore, it is necessary to point out that no
attempt will be made to review the vast literature on the depression of
245
246 2. ANALOGS OF ENZYME REACTION COMPONENTS
growth and proliferation exerted by many analogs, inasmuch as my pur-
pose here is to restrict the discussion to enzymic and metabolic levels.
TERMINOLOGY
The term analog is defined very broadly as any substance that is in some
way structurally related to a substrate, coenzyme, or cofactor.* It may
either participate in the enzyme reaction to a greater or lesser extent than
the normal components, or inhibit the reaction by interfering with the
functioning of these normal components. The commonly used term anti-
metabolite generally implies that the substance is a biologically abnormal
compound synthesized in the laboratory and capable of interfering with the
reactions of some cellular metabolite. We shall in this chapter frequently
be concerned with inhibitions produced by substances such as carbohy-
drates, amino acids, purines, and nucleotides, which naturally occur in most
cells, and thus the more general term analog is preferred. The use of the
term isostere has generally been restricted to a substance produced by the
substitution of an atom or group in the normal compound by another atom
or group with similar electronic or steric properties. A homolofj is a member
of a series in which some part or property of a basic chemical type is progres-
sively varied, as is the case when an aliphatic chain is lengthened by adding
successive methylene groups. Most of the analogs to be discussed therefore
fall in to one or more of these latter categories, but it is felt that there is
little benefit to be derived from using these more specific terms.
POSSIBLE SITES AND MECHANISMS OF INHIBITION
The most common mechanism of inhibition is a competition between
the analog and the normal reactant for a specific site on the enzyme surface.
However, the frequently made assumption that this is the only mechanism
involved is often unjustified. Other mechanisms which should be borne in
mind will be mentioned here; they will be illustrated and discussed in greater
detail later in the chapter. Let us first consider the mechanisms which may
apply particularly to the inhibition of pure enzymes.
(A) Binding of the analog to the enzyme sites for substrate, coenzyme,
or activator by interactions which are at least in part those involved in
the binding of the normal reactant and which allow reversibility.
(B) An irreversible, or practically irreversible, reaction with the enzyme
* The definition of analog must be imprecise because it is impossible to limit accu-
rately how much structural deviation can occur before the derivative can no longer
be thought of as related to the parent compound.
POSSIBLE SITES AND MECHANISMS OF INHIBITION 247
site subsequent to binding. Inasmuch as substrates occasionally form a
temporary chemical bond with the enzyme, an analog may do likewise but
fail to complete the reaction, remaining chemically attached to the site.
(C) Binding of the analog to an enzyme site other than that with which
the normal reactant interacts. Such binding may be simply fortuitous or
the site may be specifically for the purpose of allowing feedback inhibition
by a product formed in the sequence in which the enzyme participates.
Regions outside the catalytic areas with which inhibitors can react are of-
ten called allosteric sites.
(D) The analog may be a substrate of the enzyme and will inhibit the
reaction of the normal substrate to a degree dependent on the relative
binding affinities and reaction rates.
{E) Binding of the analog to a complex of the enzyme with the normal
substrate, coenzyme, or activator.
(F) The formation of a molecular complex of the analog with the normal
reactant as a result of their structural complementarity. Although such
complexes are probably uncommon and have seldom been considered in
work with analogs, we shall see that examples of this mechanism are known.
(G) Inhibition by a mechanism only indirectly related, or completely
unrelated, to the structural similarity of the analog to the normal substra,te.
An analog may, for example, possess chelating properties not exhibited
by the substrate, or it may react with SH or carbonyl groups.
When one is investigating more complex systems, particularly cellular
preparations, a number of other mechanisms for analog inhibition may be
proposed, and these should be added to the above list.
(H) The analog may interfere with the transport mechanism by which the
normal substance is taken through the cell membrane, since the two sub-
stances may both combine with some membrane carrier or enzyme system
required for efficient transport or accumulation.
(/) The analog may not be the actual inhibitor, but may be transformed
through a metabolic sequence into a substance which blocks a later reaction,
a process frequently termed lethal synthesis. In certain instances the analog
may complete a long and complex metabolic journey to terminate as a
component in some important cellular product. The incorporation of pyrim-
idine and purine analogs (e.g., the 5-halouracils, 2-thiouracil, 8-azagua-
nine, and 8-azathymine into RNA and DNA) and amino acid analogs
(e.g., tryptazan, 7-azatryptophan, ethionine, p-fluorophenylalanine, and
/5-2-thienylalanine into proteins) has been frequently demonstrated. The
products containing the analogs may be so abnormal as to fail to function
properly in the cells, thereby producing far-reaching and complex dis-
turbances.
248 2. ANALOGS OF ENZYME REACTION COMPONENTS
(J) The analog may act not on the enzyme attacking the normal sub-
strate but on an enzyme involved in the formation of this substrate, since
the precursors of the substrate will usually be structurally similar to it.
In the linear sequence:
E, Ej
X -^S-> P
an analog of S may have been designed to inhibit Eg but actually in cellular
metabolism acts primarily on Ej to reduce the rate of formation of P.
KINETICS OF ANALOG INHIBITION
The kinetics of competitive inhibition have been presented in Chapter
1-3, and the graphical analyses for the proof of competition and the deter-
mination of the constants discussed in Chapter 1-5. Type A plots of l/v
against 1/(S) have almost invariably been used to demonstrate the types
of inhibition produced by analogs, but other types of plotting may be more
satisfactory in certain situations, especially when the inhibition is not
clearly and completely competitive. It is my opinion that many kinetic
analyses of inhibition would be improved if several types of plotting proce-
dure were used, allowing comparison of the results and more accurate cal-
culations of the constants.
It is quite often the case that the inhibition by an analog is not, by the
usual methods of analysis, competitive with the substrate or coenzyme
to which the analog is structurally related, and such results have puzzled
many workers. The plotting may indicate noncompetitive, coupling, mixed,
or indeterminable inhibition mechanisms. It has been pointed out (Chap-
ter 1-3) that true noncompetitive inhibition must be rather rare and this
is particularly true for inhibition by analogs. Coupling or uncompetitive
inhibition is perhaps more common among analogs than with other in-
hibitors, especially with regard to coenzymes or cofactors, inasmuch as
the substrate combines with enzyme-coenzyme or enzyme-cofactor com-
plexes in many reactions. It is important to determine in any case if the
inhibition is really competitive and the kinetics modified to obscure this,
or whether the mechanism is actually other than competitive, in those
instances in which the graphical analysis does not demonstrate the typical
and expected competitive picture.
There are several reasons why an analog inhibition may not turn out to
be competitive by the usual plotting procedures, and it may be useful to
list them at this point.
(A) The analog may be acting by some mechanism other than specific
attachment to an active site on the enzyme, that is, by one of the mecha-
nisms listed in the previous section.
KINETICS OF ANALOG INHIBITION 249
(B) The analog may be bound very tightly to the enzyme, in which case
the order of addition of the substrate and the analog may be of great im-
portance. If the analog is added to the enzyme and the mixture is incubated
before the substrate is added, the inhibition may be very marked and the
substrate may be unable to displace the inhibitor from the enzyme in a
reasonable time. On the other hand, if both substrate and analog are added
together, the inhibition may be very low initially but progress slowly, due
to the relatively small fraction of the active centers available for reaction
with the inhibitor. In either case secondary changes in the enzyme may
occur and complicate the kinetics (Chapter 1-12). The basic problem in
the interpretation of such inhibitions is the inability experimentally to
achieve satisfactory equilibrium conditions, and it must be stressed that
the usual inhibition equations and plotting procedures apply only to equi-
librium conditions. There has been confusion between noncompetitive and
irreversible inhibitions. The arsenicals, the mercurials, and iodoacetate,
for example, are often thought to inhibit succinate dehydrogenase non-
competitively, and yet succinate and these inhibitors react at the same site
on the enzyme, as shown by the protection afforded by the presence of
succinate when it is added with the inhibitors. These inhibitions are indeed
competitive under certain conditions and during specific time intervals,
but once the inhibition has been established it is difficult to demonstrate
a competitive effect. Several examples of this situation using analog inhi-
bitors will be encountered in this chapter.
(C) The concentration of free inhibitor may be depleted due to its com-
bination with the enzyme or other materials and one is then dealing with
a mutual depletion system (Chapter 1-3). The quantitative aspects of com-
petition may be modified quite markedly in such cases. This behavior must
be looked for particularly when one is working with very potent analog
inhibitors.
(D) An irreversible or semi-irreversible change in the configuration or
properties of the active center may occur following reaction with the inhibi-
tor, so that even after dissociation of the inhibitor from the enzyme the
affinity of the enzyme for the substrate is altered. The active centers of
certain enzymes appear to be flexible and adapt in some way to the in-
teracting molecules, and it is possible that such a change would not be readily
reversible. The active center, or at least the immediately adjacent region,
of the penicillinase of Bacilltis cerevs is altered by combination with the
competitive analog of benzylpenicillin, in that there is a marked increase
in the sensitivity to iodination, and this is prevented by the presence of
substrate (Citri and Garber, 1961). Although this alteration is presumably
reversible, since the ability to hydrolyze benzylpenicillin after removal of
the inhibitor is unimpaired, it is easy to imagine changes only slowly re-
versible.
250 2. ANALOGS OF ENZYME KE ACTION COMPONENTS
(E) 111 cellular systems, or possibly in subcellular preparations of some
structural complexity, the failure of the analog to penetrate to the site of
inhibition as readily as the substrate would obviously distort the kinetics,
although the inhibition itself is truly competitive.
{F) If the substrate possesses several groups through which it is bound
to the enzyme, an analog which has only one of these groups (perhaps an
analog comprising only a part of the normal substrate molecule) may block
off one enzyme attachment point but allow the substrate to bind through
the remaining groups. In many cases this will prevent catalysis, but in
others it could only reduce the rate at which the substrate is reacted.
Simultaneously there will be a reduction in the affinity of the enzyme for
the substrate. The plots will give evidence of a mixed inhibition, which is
the actual situation, but nevertheless competition of a type is occurring.
Analogs of coenzymes often present a special problem in this connection
since the coenzyme may be bound quite tightly to the apoenzyme. The
addition of analog to the complete enzyme may not result in significant
displacement of the coenzyme and little inhibition will be observed. How-
ever, if the enzyme is resolved into its components by dissociating the coen-
zyme in some manner, competition between the coenzyme and analog may
be demonstrated in recombination experiments in which the analog reduces
the ability of added coenzyme to reactivate the enzyme.
The type of inhibition may depend on the concentration of the analog.
It has been noted several times that an inhibition may be competitive at
low analog concentrations and partially or completely noncompetitive at
higher concentrations. The noncompetitive elements of the inhibition
probably reflect the increasing unselectivity of action that is a common
property of all inhibitors when the concentration is raised beyond a cer-
tain level.
An analog of a substrate will occasionally undergo reaction in the presence
of the enzyme and, since both substrate and analog bind at the same site
on the enzyme, competition will occur. The behavior in such a situation
was discussed in Chapter 1-3 (page 96). The inhibition observed will depend
on what is determined. If we designate the substrate by S and its analog
by S':
E + S ^ES— >E + P (2-1)
E + S' ;?t ES' ^ E + P' (2-2)
and the individual rate expressions may be written as:
F„.(S)
{S)+K, 1 + Vv
A,
KINETICS OF ANALOG INHIBITION 251
F.'(S')
(S') + K,
1+^
(2-4)
If the disappearance of S or the formation of only P is measured, the inhi-
bition by S' will be typically competitive, but the inhibitor constant de-
termined by the plotting procedures, K^,, wiU not necessarily be a true dis-
sociation constant. The result wiU depend on the ratio kg.jkg, which in
most cases will be considerably less than unity. If P and P' are identical
and determined together, the inhibition wiU be less than in the previous
case. It is important in work with analogs to establish whether they are
catalytically reacted and, if so, to plan the inhibition experiments accord-
ingly. It is also necessary in many instances to determine initial rates, since
the concentration of the analog may be reduced significantly because of
its conversion to the product.
The product, or one of the products, of the enzyme-catalyzed reaction
may generally be considered as an analog of the substrate, or of part of
the substrate molecule. Inhibition by products was taken up in Chapter
1-4 (page 140) and it was pointed out there that the inhibition is not
necessarily competitive, since the product can react with the enzyme or
other components of the reaction in a variety of ways. Several instances
of product inhibition will be encountered in this chapter and in none of
these is the inhibition due to a simple reversal of the forward reaction,
but to actual combination with the enzyme at or near the active center.
A somewhat more complex situation, in which the analog is transformed
into a product that inhibits a subsequent reaction of the substrate, is fairly
common and warrants some discussion although a complete kinetic analysis
is difficult. The simplest system may be represented by:
A --^ B A C (2-5)
E. t
A' — ^ B' (2-6)
The substrate, A, and its analog, A', both are reacted by E^ and the analog
product, B', inhibits E,. There are several ways in which the rate can vary
with time and the behavior of the system will depend on many factors.
In the first place, the presence of A' may slow down reaction 1 whereby
A is transformed to B, this being a case of competing substrates. The for-
mation of C may thus be initially slowed for this reason. The concentra-
tion of B' will progressively rise and the inhibition on Ej increase. However,
the concentration of A' will decrease and the competition w^th A be reduced,
leading to a relative acceleration of reaction 1. The change in the rate of
formation of C will depend on the balance between these two types of
inhibition. For example, if A' is reacted fairly rapidly but B' is not a very
252 2. ANALOGS OF ENZYME REACTION COMPONENTS
potent inhibitor of Eg, the rate may first rise as the inhibition on E^ is
released, and then fall later when the concentration of B' rises sufficiently.
On the other hand, if A' is not readily depleted and B' inhibits well, the in-
hibition will steadily increase. It is clear that the kinetics may not indicate
competitive inhibition when the rate of formation of C is measured. The
concentration of the intermediate B may rise and fall in a complex manner,
as discussed in Chapter 1-9 (page 438). This is actually the simplest case
of lethal synthesis complicated by competition in the initial reaction.
MEANS OF EXPRESSING RESULTS
The results in the study of analogs have usually been given in terms of
inhibitions at different concentrations of substrate and analog.* Such
results are often valuable but rather difficult to interpret, especially when
different analogs at different concentrations must be compared. It would
be more valuable if inhibitor constants were calculated and presented,
along with Michaelis or substrate constants, and fortunately this practice
is becoming more common. With values of K,„ and K^ it is possible to de-
termine the inhibition expected at any combination of substrate and analog
concentrations. Furthermore, it is possible to calculate relative interaction
energies from a series of K-q obtained for a group of analogs, and thereby
put the inhibition on a more molecularly interpretable basis.
There are fundamentally two types of /i",. The actual dissociation constant
for the EI complex may be called the true inJiihitor constant and refers to
a particular free and active form of the inhibitor. The experimentally
determined K^, on the other hand, often differs from the true Ki and may
be termed the apparent inhibitor constant. This apparent K^ may depend
on a number of factors, the most important of which is usually the pH
since many inhibitors are weak acids or bases. This problem has been
discussed in some detail in Chapter 1-14. If the inhibitor is a weak acid
only one form may be the inhibitor, say the ionized I~ form, in which case
the apparent K^ will vary with pH in the range around the p^^ of the
inhibitor. The true K^, which refers to the equilibrium
E + I- ^ EI-
will not vary with the pH because of changes in the ionization of the inhi-
bitor, although it may for other reasons. The apparent K^, in fact, will
depend on any type of equilibrium between active and inactive forms of
the inhibitor, for example an enol-keto isomerism, or on the binding of
the inhibitor to nonenzyme components of the preparation. The Kj's given
* The substrate concentration has been omitted in some reports and this vitiates
the results and makes them quantitatively meaningless.
MEANS OF EXPRESSING RESULTS 253
in this chapter will in almost all cases be apparent inhibitor constants be-
cause the true K/s have generally not been calculated in the reports,
although in many instances they must be very close to the true K/s because
of the nature of the analog or the conditions of the experiments. I have
taken the liberty in certain cases where the data are adequate of calculating
the K/s from the inhibitions reported. In every instance where plotting
procedures have been used to determine the type of inhibition, it would
have been possible to determine the appropriate constants, but these con-
stants have been seldom reported.
An especially unsatisfactory means of expressing the results is to give the
inhibition for a certain ratio of inhibitor to substrate concentrations,
(I)/(S), which has often been called the inhibition index. If the actual con-
centrations are not given, it is not possible to visualize the inhibition quan-
titatively, because (I)/(S) is not constant for a certain degree of inhibition
even when the inhibition is competitive. This has been clearly pointed out
in Chapter 1-3 (page 106) but it is important to re-emphasize it here. The
ratio, (I)/(S), is constant and meaningful only at high substrate concen-
trations which saturate the enzyme. This may be seen from the following
expression, obtained by rearranging the equation for competitive inhibition:
1 1
~k7 ^'Wi
1
(2-7)
It has sometimes been assumed tliat for 50% inhibition, (I)/(S) = KJKg,
but this is not necessarily true, which can be easily seen by rewriting Eq.
2-7 for i = 0.5 and (S) = nK,:
(I)
l:[v + ^l
, -. (2-8)
(S) Ks ' '
(I)/(S) = KJK^ only when the substrate concentration is high relative to
Kf. (i.e., when n is much greater than unity). This can also be seen in another
way: If K, = 1, K^ = 0.3, and (I)/(S) is kept constant at 0.3, the inhibition
will vary with the absolute magnitudes of the concentrations as shown in
the following tabulation:
(S)
(I)
i
0.1
0.03
0.083
0.3
0.09
0.19
0.5
0.15
0.25
1
0.3
0.33
2
0.6
0.40
5
1.5
0.45
10
3
0.48
20
6
0.49
254 2. ANALOGS OF ENZYME REACTION COMPONENTS
It is obvious that a statement that the inhibition was a certain value at
(I)/(S) = 0.3 would be of little significance.
When a series of analogs is tested, quantitative expression of the relative
affinities of the enzyme for the various analogs is desirable when possible.
The term affinity implies no units and has a vague meaning, having been
used in a variety of ways. A common method of expressing the affinity
is to equate it tolIK^ in the case of an inhibitor, and to HKg for a substrate.
The ratio of the affinities of an inhibitor and a substrate is thus often ex-
pressed as KJKi. If K, = 1 mM, and K, = 0.001 mM, it would be stated
that the affinity of the enzyme for the inhibitor is 1000 times that for the
substrate. This may sound dramatic but is misleading in a way. It would
seem that affinity might be better expressed in terms of binding energies.
The ratio of the free energies of binding of inhibitor and substrate is
given by:
AF, vKs
(2-9)
In the hypothetical case above, AFJAF^ = 2, and the inhibitor is bound
twice as tightly as the substrate, which is a more reasonable way of desig-
nating the relative affinities.
One might consider three types of AF for the binding of an inhibitor to
an enzyme. There is the true AF corresponding to the true K^^ and an ap-
parent AF corresponding to the apparent experimental K^. In addition
there is the theoretical AF corresponding to the interaction energy of the
enzyme and inhibitor in a vacuum, uncomplicated by solvent and ions. In
order to compare different analogs with respect to their affinities for the
enzyme, it is actually this last AF one would wish in most cases, but it is
impossible to obtain. Lacking this, one must use the true AF values, but,
as pointed out above, true K/s are not often available. It may be quite
misleading to compare calculated AF values for a series of analogs if these
analogs have different p-ff^'s and the experimental pH is in the region of
these p-fi'./s. The differences in the AF's may reflect mainly the different
degrees of ionization rather than the differences in binding energies of the
inhibitory forms. In several instances in this chapter, I have calculated the
AF values for such series of analogs and it must be remembered that the
validity of comparing these values is sometimes questionable. It is some-
times impossible to calculate even an apparent AF and one must be satisfied
with values that are relative to some chosen compound; these will be called
relative AF values. Although the AF itself may not be meaningful, occa-
sionally the difference of the AF values for two inhibitors will be significant
in attributing interaction energies to certain groups, as discussed in Chapter
1-6 (page 268), since all of the other factors involving the solvent and ionic
atmosphere may remain relatively constant for the inhibitors. The relative
binding energies may be calculated in some cases even though the K^'a
IMPORTANT TYPES OF MOLECULAR ALTERATION 255
are not known, since for two inhibitors:
AF, — AF, = 1.422 log [' il ~ \'l (2-10)
ii (1 —I2)
if(Ij) = (l2)-
It is easy to derive an expression for the relationship between the true
AF difference and the apparent JF difference for two inhibitors when the
sole factor involved is the ionization of the inhibitors. If we designate the
true free energy of binding as AF and the apparent free energy of binding
as AF', the difference in the true binding energies for the two inhibitors
will be AFi — AF^, and the difference in the apparent binding energies
will be AF^ — AF^'. The relationship between these for analogs that are
singly ionizing weak acids is:
(AF, - AF,) = {AF,' - AF,') - 1.422 log \ + [|h^)/^?] ^^'^^^
It is therefore possible to correct for the pH effect if the p/C^'s of the inhibi-
tors are known and the true interaction energy difference may be obtained.
IMPORTANT TYPES OF MOLECULAR ALTERATION
PRODUCING INHIBITING ANALOGS
A substrate or coenzyme might generally be considered to have three
different types of molecular region: (1) groups involved primarily in the
binding to the enzyme, (2) groups involved in the catalytic reaction, and
(3) groups or regions not directly involved in either binding or reaction.
There is overlap between these in some cases, of course, because the groups
undergoing chemical change usually participate to a certain extent in the
binding. Furthermore, some substrates, such as succinate, do not possess
the third type of group, all of the molecule being directly involved in bind-
ing and reaction. The properties of a substance produced by altering a single
group or region of a substrate wiU depend on the type of group or region
that is modified, that is, its function in the reaction of the substrate with
the enzyme. A change in a binding group will usually alter the interaction
energy in the combination of the substance with the enzyme and may or
may not affect the susceptibility to chemical reaction, whereas a change in
a group directly involved in the catalysis will generally reduce the reactivity
without necessarily modifying the binding to the enzyme. It is also evident
that a change in the third type of neutral group will not be so likely to
alter the behavior of the substrate, unless such a change in some way
secondarily modifies the interactions of the other groups. The aim in the
design of analogs for enzyme inhibition is to produce a compound which
will bind reasonably tightly to the enzyme (preferably more tightly than
256 2. ANALOGS OF ENZYME REACTION COMPONENTS
the substrate), but which is resistant to chemical reaction in its complex
with the enzyme. It would appear that the most effective inhibitors would
result from modifications of the reactive groups, or of groups adjacent
to the reactive region, rather than changes in binding groups. Malonate
illustrates this in a simple way because here the — CH2CH2 — group of
succinate has been altered to the nonoxidizable — CHg — group, while
the binding — COO" groups remain. Amidation of the — C00~ groups of
either succinate or malonate, thereby eliminating the negative charges,
produces a substance that is neither a substrate nor an inhibitor because
the affinity for the enzyme has been lost.
The total interaction energy between a substrate or an inhibitor and the
enzyme active center is the result of all the forces of attraction and repulsion
summed over all the participating groups. Every atom or group of a sub-
strate or its analog contributes to some degree to the interaction energy
but, practically, the binding may be attributed usually to two or three
groups that serve to orient the molecules on the enzyme surface. The sub-
traction, addition, or alteration of substrate groups may change the binding
energy in various ways. Modifying a region vicinal to a binding group may
sterically interfere with the normal approach of this group to the enzyme
group with which it interacts, or it may by inductive or resonance effects
alter the properties of the binding group, as discussed in Chapter 1-6
(page 304). It is worthwhile emphasizing again that a change in a particular
region of a molecule may produce variations in the electronic configurations
throughout the entire molecule, and that a change in the interaction energy
cannot generally be attributed solely to this altered region. Furthermore,
the volume and configuration of a substrate and its analog may involve
water of hydration, so that a group change can secondarily affect the bind-
ing by modifying the disposition of the bound water molecules. The in-
troduction of so-called neutral groups, such as hydrocarbon chains, can
bring about an increase in the binding energy through nonspecific van der
Waals' interactions, providing these groups do not interfere sterically with
the approach of the important binding groups to the enzyme surface.
Most analogs of substrates have less affinity for the enzymes than do the
natural substrates, which is reasonable in view of the enzyme active center
conformation to the substrate configuration. However, occasionally an
analog will exhibit a much tighter binding than the substrate, the extra
binding energy being more than could be attributable simply to a new group
introduced into the molecule. In such cases it is likely that a qualitative
change in the binding is involved. A substrate frequently forms a covalent
bond with the enzyme during the catalytic reaction, and normally this
constitutes only an intermediate state in the sequence of changes. Certain
analogs may be able to form this type of bond but are unable to complete
the sequence, so that the analogs remain firmly attached to the enzyme.
IMPORTANT TYPES OF MOLECULAR ALTERATION 257
This is known to occur in the case of diisopropylfluorophosphate and re-
lated cholinesterase inhibitors, as well as with monoamine oxidase inhibi-
tors such as iproniazid.
An analog may be related to the corresponding substrate in one of two
general ways: it may be either an isomer of tne substrate, or a substance
obtained by the replacement of one or more groups on the substrate.
An isomeric analog may be a geometric isomer (e.g., one of a cis and trans
pair), optical isomer, or any stereoisomer of the substrate. It might be
thought that such analogs would often be specific and useful inhibitors but
actually, except for certain optical isomers (see page 268), this is seldom
the case, the reason being that the configuration of the analog is more
important than a simple equivalence of all the atoms and groups. A sub-
stitution analog can result from a variety of molecular changes in the sub-
strate. What is often called addition of groups is usually only a substitution
of the new group for a H atom (e.g., the replacement of a H atom with a
F atom or a CH3 group), and what is called deletion of groups is usually
a substitution of a H atom for the group that is removed. On the other hand,
an important type of analog is derived by the substitution of one functional
group with another group (e.g., the replacement of an OH group with a
SH group, or of a CH3 group with a CI atom). Some commonly interchangea-
ble groups might be put in the following families:
(a) — NH2 —OH — SH — CH3 —CI — F — H
(b) —COO- — SO3- — ASO3H- — PO3-
(c) — CONH2 — SO2NH2
{d) — S— —0— — NH— — CH=CH— — CH2—
(e) — Phenyl — benzyl — pyridyl — pyrimidyl — cyclohexyl
A good deal has been written about isosteric and isomorphic groups in the
production of analogs (for an excellent review see Schatz, 1960), especially
with respect to the development of new drugs, but this has limited bearing
on the elaboration of enzyme inhibitors. The replacement of substrate
groups with isosteric and approximately isomorphic groups usually leads
to substances that are also substrates. It is generally necessary to alter
the proper region of the substrate molecule significantly in order to produce
an effective inhibitor. There are actually at the present time no general
rules for the most efficient procedures to be used for the modification of
substrates to produce inhibitors. Various enzymes exhibit quite different
reaction mechanisms and an effective transformation of the substrate in
one case will not work for other enzymes. For this reason the most important
thing to establish initially is the nature of the particular enzyme mechanism,
if an attempt is to be made to design analogs rationally. The binding groups,
reactive groups, and relatively neutral groups in the substrate must be
258 2. ANALOGS OF ENZYME REACTION COMPONENTS
determined so that modifications in the structure may be made in the proper
regions. The examjjles discussed in this chapter will clearly demonstrate
that many analog inhibitors are not isosteric or isomorphic with the sub-
strate and that, indeed, many of the most useful inhibitors appear to dif-
fer quite markedly from the substrate. In this connection it is necessary
to call attention to the danger of visualizing molecules on the basis of their
classic two-dimensional formulas. One must also realize that usually not
all of a substrate molecule is involved in the binding and reaction with the
enzyme; the side of the molecule more distant from the enzyme may be
relatively less important than the rest of the molecule and modifications
on this side may give the appearance of producing radically different sub-
stances, whereas from the standpoint of the enzyme surface these sub-
stances may be very similar to the substrate. Conversely, just because two
substances look alike when written in the usual structural formulas is not
enough to ensure that they will exhibit comparable interactions with the
active center of an enzyme. Ideally, one should learn to conceive enzyme
reactions on an electronic and molecular level and to visualize analogs
with three-dimensional molecular imagination, in other words, to approach
the problem of analog design from the point of view of a rational active
center.
Some analogs are not directly inhibitory but are metabolically trans-
formed into inhibitory substances that block a sequence at a later step.
Such analogs are often very interesting and useful, being in many cases
specific and potent. The design of an analog to be an inhibitor precursor
presents a slightly different problem than in the general case. If the analog
is to enter into the metabolic sequence it must be a substrate of the initial
enzymes and, hence, structurally similar to the natural substrate in the
region of the reactive groups. Isosteric and isomorphic substitution is often
useful in this situation. This probably accounts for the popularity of fluo-
rine as a replacement for a H atom in the design of this type of analog.
Many F-substituted analogs enter into metabolic sequences and interfere
at a more distal region, for example, fluoroacetate, 5-fluorouracil, 2>-fluoro-
phenylalanine, and 6-deoxy-6-fluoro-D-glucose. The F atom is the smallest
of the common atoms that may be substituted for a H atom, and approaches
the H atom in size (van der Waals' radii for H and F atoms being 1.2 and
1.35 A, respectively) (see Table 1-6-8). The F atom is also relatively un-
reactive and forms a stable bond to carbon. However, it is strongly electro-
negative and alters the electronic configuration in comparison to the parent
compound. The dipole moment of the C — F bond will be quite different
from the C — H bond, this altering neighboring bonds as well as inducing
an ability to form hydrogen bonds with the enzyme. Thus the F analog
not only may be much like the substrate in over all size and configuration,
allowing it to be metabolically reactive, but eventually may be transformed
THE CONCEPT OF INHIBITION BY ANALOGS 259
into a substance in which the F atom, because of its electronegative char-
acter, interferes in some way with the catalytic reaction. Another consid-
eration arises when phosphorylation of the substrate is an initial step in
the metabolic sequence. Here one must be careful not to alter the groups
involved in the phosphorylation, but to modify groups that are reactive in
a later enzymic step.
DEVELOPMENT OF THE CONCEPT OF INHIBITION
BY ANALOGS
This important concept, which today plays a major role in the fields
of enzymology and biochemistry and is becoming more and more important
in pharmacology, chemotherapeutics, and pathology, has an interesting
history illustrating a typical growth pattern of a scientific idea. The de-
velopment of the general concept has been traced by Martin (1951), Wool-
ley (1952), and Albert (1960), so that here it is necessary only to present a
cursory exposition related particularly to enzyme inhibition. However,
it must be realized that many fields of study — including immunology,
drug antagonism, and microbial growth inhibition among others — con-
tributed in one way or another to this concept.
Despite the fact that numerous examples of competitive inhibition by
analogs had been demonstrated since 1910, and that the analog concept
had been quite clearly stated around 1930, general recognition of the basic
principles did not occur until after 1940. Wohl and Glimm (1910) reported
the inhibition of amylase by glucose and galactose, as well as by the reac-
tion product, maltose, and shortly Michaelis described the inhibition of
/5-fructofuranosidase by fructose and a-methylglucoside (Michaelis and
Pechstein, 1914), and the inhibition of a-glucosidase by glucose (Michaelis
and Rona, 1914). Isolated, unpremeditated, and unrecognized discoveries,
such as the inhibition of arginase by ornithine (Gross, 1921) and the inhi-
bition of /?-fructosidase by /^-glucose but not by cf-glucose (Kuhn, 1923) un-
fortunately gave no impetus to the formulation of a general concept. Even
the classic work of Quastel on malonate inhibition, described in the previous
chapter, wherein competitive inhibition by structural analogs was considered
in a modern fashion, apparently did not activate anyone outside Cambridge,
where Bernheim (1928) studied the aconitate inhibition of citrate oxidation,
Murray (1929) and Murray and King (1930) applied the principles of
analogs (although in a somewhat naive way) to the inhibition of lipase by
various ketones and alcohols, Richter (1934) proved the inhibition of ca-
techol oxidase by resorcinol to be competitive, Keilin and Hartree (1936)
reported potent competitive inhibition of uricase by the methylurates, and
Green (1936) extended the malonate inhibition of succinate dehydrogenase
to the inhibition of malate dehydrogenase by several dicarboxylates.
260 2. ANALOGS OF ENZYME REACTION COMPONENTS
Several accidental observations were made of the toxicity of metabolite
analogs while they were being tested for activity in animals, for example,
the discovery that ethionine is toxic to rats (Dyer, 1938) and that pyridine-
3-sulfonate is lethal to dogs suffering from nicotinic acid deficiency (WooUey
et al., 1938), but the importance was not immediately recognized. Appar-
ently the stimulus for the formulation of the general theory had to come
from a discovery of clinical importance, and this was provided by the find-
ing of the antagonism by p-aminobenzoate of the action of the sulfona-
mides by Woods (1940). Between 1940 and 1943 many examples of analog
antagonism were reported, and from 1943 to 1946 it was shown that vi-
tamin deficiency symptoms could be produced in animals by the administra-
tion of the appropriate analogs of all the known vitamins. It was then
possible to return to the enzyme level and apply the principles formulated
long before to the new results. Since 1946 there has been a steadily increas-
ing interest in analog inhibitors, which is reflected in the large numbers of
publications on this subject each year (Fig. 2-1). At the present time a paper
on analog enzyme inhibition as covered in this chapter appears every other
day; this does not include all of the investigations on analog inhibitors
that are to be treated separately (such as fluoroacetate, arsenate, carbon
monoxide, parapyruvate, and others), or analogs commonly used clinically
Fig. 2-1. Curve showing the annual number of publications on analog inhibition
of enzymes.
ANALOG INHIBITION OF MEMBRANE TRANSPORT 261
(sucli as the cholinesterase and monoamine oxidase inhibitors), or antime-
tabolites used in microbial growth suppression or tumoristasis.
A few general references treating aspects of the subject not included
in the present work are suggested for additional information: Welch (1945),
Work and Work (1948), Woolley (1950 b, 1952), Martin (1951), Rhoads
(1955), Matthews (1958), Albert (1960), Schueler (1960), Schatz (1960),
Kaiser (1960), and the Symposium on Antimetabolites sponsored by the
National Vitamin Foundation (1955). A great deal of information on the
biological actions of many types of analog will be found in Volume I of
"Metabolic Inhibitors" edited by Hochster and Quastel (1963), and this
aspect of the subject will be mainly omitted in the present work so that
the enzymic effects may be discussed in sufficient detail.
ANALOG INHIBITION OF MEMBRANE TRANSPORT
Before discussing specific enzyme inhibitions by analogs, we must turn
our attention to the possibility that the depression of the utilization of
some substrate or metabolite by an analog is not due to an action on any
enzyme involved in the metabolism, but is the result of a specific inter-
ference with the transport of the substrate or metabolite into the cell.
It is quite probable that some of the actions of analogs on metabolism, at-
tributed to competition at the enzyme level, are actually exerted at the
cell membrane; indeed, certain instances will be discussed. It is becoming
more and more evident that many substrates and metabolic precursors
are taken up by cells by processes other than simple diffusion, and this
applies particularly to certain carbohydrates, amino acids, coenzymes,
or coenzyme precursors. It is not necessary that the transport be active
to be influenced by analogs; any movement of a substance through a mem-
brane which involves a carrier, a special size or configuration of pores, or a
specific type of mechanism, active or passive, can be slowed in the presence
of an analog of this substance. It is possible, o^ course, that the membrane
trasport is mediated through an enzyme reaction, such as a phosphoryla-
tion, and the competition by the analog is then truly an enzymatic one.
It is sometimes difficult to determine if there is inhibition of a transport
process. The demonstration of a reduced uptake of a substrate from the
medium is generally not sufficient evidence, inasmuch as a decreased in-
tracellular utilization might also be responsible. The best procedure is to
determine directly the concentration of the substrate within the cells in
the absence and presence of the analog, but sometimes the concentration
is too low to measure accurately, especially when the substrate is rapidly
metabolized. The demonstration of typical competitive inhibition with
respect to the action of an analog on some metabolic process is also not
sufficient evidence for an enzymic site of action, because the inhibition of
262 2. ANALOGS OF ENZYME REACTION COMPONENTS
membrane transport may exhibit competitive kinetics. The kinetics will
often depend on whether the transport is limiting the metabolic utilization
of the substrate or not. Membrane transport involving a carrier (C) molecule
can often be represented by:
S + C ^ SCo -> SC, -► C + S (2-12)
where the subscripts refer to the outside and inside of the membrane. The
later reactions may be written in one direction only because of the utili-
zation of the substrate as it enters the cell. If an analog also combines
with the carrier:
S' + C ^ S'C (2-13)
whether it is transported into the cell or not, typical competitive behavior
will be observed since the forms of Eqs. 2-12 and 2-13 are the same as
those for competitive enzyme inhibition.
Plots of transport rates against the external concentrations of the trans-
ported substance usually yield hyperbolic curves, and double reciprocal
plots are often linear, allowing the calculation of a constant which corres-
ponds to the Michaelis-Menten constant in enzyme kinetics. It is frequently
assumed that this is the dissociation constant for the complex of the sub-
stance with the carrier, but it is not necessarily true for the same reasons
that K^,^ is not always K,. The kinetics of transport inhibition are likewise
commonly similar to those observed with enzymes and values of K^ may
be determined by appropriate plotting, this constant representing the dis-
sociation constant of the carrier-analog complex. The kinetics of carrier
transport and its inhibition have been elaborated by Wilbrandt and Rosen-
berg (1961) and Rosenberg and Wilbrandt (1962) for facilitated diffusion
and certain restricted types of active transport. It is interesting in connec-
tion with certain types of inhibition work to note that the accumulation
ratio, (X),/(X)^ = VJkK„j, where F^„ is the maximal transport rate, K,„
is the Michaelis-Menten constant for transport, and k is the passive diffu-
sion constant for the membrane. Thus the cell/medium ratio may be altered
by the inhibitor as a result of changes in any of these three parameters.
Carbohydrate Transport
Competition between sugars for entrance into cells has been observed
in many tissues but has seldom been studied quantitatively, so that in most
cases it is impossible to know if true competition kinetics are followed.
Occasionally a reduction in the inhibition with increase in the concentration
of the transported substrate has been noted; for example, the active ac-
cumulation of D-galactose by rabbit kidney cortex slices is inhibited 61%
by 5.6 rciM glucose when D-galactose is 0.1 mM and only 28% when d-
galactose is 0.2 mM (Krane and Crane, 1959), but these results do not fit
ANALOG INHIBITION OF MEMBRANE TRANSPORT 263
a simple competitive formulation. Indeed, the exact site of inhibition has
not been established in any case. The entrance of L-arabinose into rat
heart cells is inhibited 92% by glucose at equimolar concentration and,
since L-arabinose is not metabolized, the inhibition is presumably on some
phase of membrane transport (Morgan and Park, 1958). In the same tissue
the competition between glucose and 3-methylglucose at the outer but
not the inner surface of the cell membrane leads to a net outward transport
of 3-methylglucose against a concentration gradient, and it was concluded
that there are stereospecific combining sites at both surfaces. A compari-
son between cellular and subcellular preparations can occasionally indicate
a membrane site for an inhibition. Galactose markedly inhibits the utili-
zation of fructose by intact ascites tumor cells but not in homogenates,
pointing to a competition before the hexokinase step and probably in
transport (Nirenberg and Hogg, 1957). The effects of 2-deoxy-D-glucose
on the fermentation of glucose, fructose, and mannose by yeast (to be
discussed in more detail on page 391) led Scharff (1961) to assume a trans-
port system which is stable, since it still functions in acetone-dried cells,
and perhaps bound to a complex or "bundle" of fermentation enzymes
somewhere in the outer regions of the yeast cells.
Although the nature of the transport mechanisms and the site of inhi-
bition are generally not known, it is clear that the interference is quite
specific and dependent on the molecular configurations of the sugars. The
transport of D-galactose (5 mM) by hamster jejunum is inhibited to varying
degrees by other sugars and derivatives at 25 nxM (see tabulation) (Wilson
Sugar % Inhibition
D-Mannose Stim 4
D- Xylose 7
3-0-Methyl-D-glucose 59
a-Methyl-D-glucoside 95
et al, 1960), and the uptake of glucose by lymph node cells is likewise
inhibited differently by several sugars at 9 mM (see tabulation) (Helm-
reich and Eisen, 1959), both observations pointing to stereospecific trans-
Sugar % Inhibition
D-Arabinose 7
D-Galactose 9
D-Fructose 40
D-Mannose 61
264 2. ANALOGS OF ENZYME REACTION COMPONENTS
port systems presumably involving carriers in the membranes. The pat-
terns of competition between various sugars have often led to the assumption
of two or more different transport mechanisms for carbohydrates and that
these systems can operate simultaneously and independently. Competition
for transport can be demonstrated in rat diaphragm muscle for members
of the group including D-glucose, D-mannose, D-xylose, D-arabinose, l-
arabinose, D-lyxose, and 3-0-methyl-D-glucose, but not between these
and D-galactose, D-fructose, maltose, a-methyl-D-glucoside, and /?-methyl-
D-glucoside (Battaglia and Randle, 1960), so that different sites for entry
were postulated. A number of sugars are transported into erythrocytes:
all the aldoses penetrate by a transport system characterized for glucose,
while the ketoses penetrate according to a pattern of passive diffusion
(LeFevre and Davies, 1951). The aldoses compete with each other, e.g.,
the uptake of glucose is strongly inhibited by mannose, but any aldose
delays the entrance of a ketose, e.g., the uptake of fructose is prevented
by glucose and galactose, while the ketoses do not perceptibly alter the
transport of the aldoses. One carrier system in the erythrocyte has been
characterized as reacting only with those monosaccharides in which the
pyranose ring tends to assume the "chair" configuration, which illustrates
a unique type of stereospecificity (LeFevre and Marshall, 1958). On the
other hand, in hamster intestine only a single transport mechanism or car-
rier seems to be present, mutual inhibition occurring between D-glucose
and D-galactose, D-glucose and 1,5-anhydro-D-glucitol, D-galactose and 1,5-
anhydro-D-glucitol, D-glucose and 6-deoxy-D-glucose, and 6-deoxy-D-glucose
and 1,5-anhydro-D-glucitol, whereas sugars that are not transported do not
interfere with those that are (R. K. Crane, 1960).
Amino Acid Transport
The situation here is very much the same as in sugar transport and there
is good evidence for stereospecific systems and multiple pathways. There
is competition between L-leucine and DL-isoleucine for renal tubular resorp-
tion in the dog, these being well resorbed amino acids, and there is also
competition between the poorly resorbed L-arginine and L-lysine (Beyer
et at., 1947). However, no interference is observed between L-leucine and
L-arginine. It is possible to classify the amino acids into groups with respect
to their mutual interference in resorption. In the rat kidney, dibasic amino
acids (arginine, lysine, and cystine) are actively accumulated and there
is mutual inhibition of the transport (Rosenberg et al., 1962). Monobasic
amino acids (alanine, phenylalanine, and histidine) do not interfere with
the uptake of the dibasic amino acids, nor does arginine depress the uptake
of the monobasic amino acids. It seems likely that separate transport sys-
tems are present. The transport system for basic amino acids in the hamster
intestine is distinct from that for other amino acids and similar to the renal
ANALOG INHIBITION OF MEMBRANE TRANSPORT
265
system (see accompanying tabulation) (Hagihira et al., 1961). Arginine
and cystine interfere more with lysine transport than with glycine trans-
port, whereas methionine behaves in the reverse manner, and there is
/-I J. A.- % Inhibition of transport of :
, , ., . Concentration
inniDitor
{raM)
Glycine
L-Lysine
L-Arginine
2
16
89
L-Cystine
0.8
0
45
L-Methionine
1
73
32
L-Lysine
1
14
—
Glycine
1
—
0
little or no interference between glycine and lysine. Proline, histidine, and
glycine are actively transported across the intestinal wall and methionine
at eqiiimolar concentration completely inhibits this (Wiseman, 1954). This
intestinal transport carrier is limited to monoamine-monocarboxylates and
they compete with each other. In addition to a common carrier, there may
be an additional carrier for glycine and proline (Newey and Smyth, 1964),
and it was pointed out that although each carrier system would conform
to Michaelis-Menten kinetics, the total transport with two or more carriers
involved would not necessarily.
Intestinal transport systems may react with only the l- or the D-form
of an optically isomeric pair. D-Methionine is accumulated and transported
against a concentration gradient by the rat intestine and this is blocked
completely by equimolar concentrations of L-methionine ( Jervis and Smyth,
1960). Similarly, the transport of L-I^^^-monoiodotyrosine, which involves
active accumulation of the amino acid in the gut wall, is inhibited by
many L-amino acids (the most effective being L-tryptophan, L-methionine,
L-leucine, and L-isoleucine) but scarcely at all by any of the four D-amino
acids tested (Nathans et al., 1960). L-Tryptophan, for example, at 10 mM
reduces the tissue/medium ratio from 8.85 to 1.55 and the inhibition is
apparently competitive.
The active cumulative uptake of amino acids by ascites carcinoma cells
comprises several transport systems, each with a specific range of substrates.
The uptake of glycine-1-C^* {K,,, = 6.4 mM) is most strongly inhibited by
1-aminocyclopentanecarboxylate {K, = 1.47 mM), and this is competitive,
while the uptake of DL-methionine-S^^ {K„, = 1.7 mM) is inhibited best
by allylglycine {K^ = 0.86 mM). Glycine transport is moderately inhib-
ited by aUylglycine and less readily by furylglycine and thienylglycine
(Scholefield, 1961). On the other hand, the uptake of DL-leucine-1-C^^
is stimulated by most of these inhibitors. Transport of L-tryptophan in
266 2. ANALOGS OF ENZYME EEACTION COMPONENTS
ascites cells occurs by both diffusion and active transport (Jacquez, 1961).
Certain amino acids accelerate this at lower concentrations (1 mM) and
competitively inhibit at higher (5 mM), while other amino acids (such
as L-alanine, L-lysine, and L-arginine) only inhibit. Oxender and Christen-
sen (1963) thoroughly studied the effects of many amino acids on the up-
take of neutral amino acids by ascites cells and found they fall into two
overlapping clusters, the transport systems apparently not being very
specific.
The penetration of amino acids into the brain is probably important
for the metabolism and function of that tissue, and there appear to be
several transport systems available. Tyrosine enters the brain readily in
vivo and a specific transport is probably involved, since L-tyrosine pene-
trates more rapidly than D-tyrosine and the entry is potently inhibited by
certain other amino acids, particularly L-tryptophan, L-leucine, L-valine,
/5-fluorophenylalanine, and L-histidine (Chirigos et al., 1960). In phenyl-
ketonuria the blood levels of phenylalanine are high due to the inability
of the tissues to metabolize it to tyrosine. It is possible that these high
concentrations can interfere with the entry of other amino acids into the
brain and partially account for the central nervous system disturbances.
The uptake of five amino acids by rat brain slices is indeed inhibited by
L-phenylalanine (see accompanying tabulation) and it was felt that such
Amino acid (2 raM) % Decrease of concentration gradient
L-Proline 9
L-Histidine 42
L-Arginine 46
L-Ornithine 47
L-Tvrosine 70
could occur in vivo (Neame, 1961). The transport of L-histidine is inhib-
ited by neutral aliphatic amino acids and short-chain diamino acids to a
degree dependent on the length of the carbon chain (Neame, 1964). Inhi-
bition by the dicarboxylic amino acids is not dependent on the chain length.
In general, the L-isomers inhibit more potently than the D-isomers. It was
suggested that histidine is transported by a system which transports most
other amino acids, but with different affinities, since the inhibitions are all
competitive. The synthetic amino acid, 1-aminocyclopentanecarboxylate,
is not metabolized but is actively transported in brain slices and ascites
cells, and the system involved must be the same as for methionine since
it is affected similarly by the same competitive amino acids (Ahmed and
Scholefield, 1962). Such transported but nonmetabolized amino acids may
well be of use in studying transport inhibition, since effects can be clearly
ANALOG INHIBITION OF MEMBRANE TRANSPORT 267
distinguished from possible inhibitions of amino acid incorporation in the
cell. Reference should be made to the very complete investigation of amino
acid transport in brain slices by Abadom and Scholefield (1962), in which the
many competitive inhibitions established point to several separate amino
acid transport systems.
The entrance of valine, proline, and hydroxyproline into the human
erythrocyte is not mutually competitive, but is inhibited markedly by
certain sugars, such as glucose, galactose, and xylose, although not by
fructose (Rieser, 1961). These results would indicate that some amino acids
and sugars follow the same transport pathway. If this is a general phenome-
non, one must consider in the use of analogs of these substances the possi-
bility that an amino acid analog might depress glucose uptake, and thereby
secondarily interfere with the transport by suppressing energy generation.
The accumulation of L-histidine by the parasitic fungus Botnjtis fabae
is inhibited by most other amino acids, and one transport system seems to
be available to all the amino acids (Jones, 1963). Substitution at the NH2
or COOH groups lessens or abolishes the inhibitory activity, indicating
that the binding to the carrier is at least partly electrostatic.
Miscellaneous Transport Systems
The oxidation of protocatechuate by a Flavohacterium is competitively
inhibited by ^-aminosalicylate and one might conclude that mutual in-
teraction with some enzyme is responsible. However, 59-aminosalicylate
does not affect the rate or extent of the oxidation in extracts, measured in
different ways (Hubbard and Durham, 1961). These results thus point to
competition for a transport system in the membrane, rather than the more
usual explanation. The active transport of biotin across the hamster in-
testine is inhibited by various analogs (e.g., biocytin, desthiobiotin, di-
aminobiotin, and biotin methyl ester), but the nature of the inhibition was
not investigated so it may not be competitive (it is not with lipoate) (Spen-
cer and Brody, 1964). The active influx of urate into erythrocytes is com-
petitively inhibited by hypoxanthine with K^ = 0.1 mM, whereas the
efilux consists of two components, one sensitive to hypoxanthine (Lassen
and Overgaard-Hansen, 1962).
There are several instances in which the transport of inorganic ions is
inhibited by other related ions. The transfer and exchange of phosphate
across the membrane of S. aureus are inhibited by chlorate, borate, and ar-
senate (Mitchell, 1954), although only arsenate is able to substitute com-
pletely for phosphate in the exchange. The uptake of sulfate by yeast is
competitively inhibited by thiosulfate (Kleinzeller et al., 1959). Nitrate
inhibits quite well the accumulation of iodide in the rabbit ciliary body,
50% reduction of the tissue/medium ratio occurring at 3 vaM, but it is
not known if this is truly competitive (Becker, 1961).
268 2. ANALOGS OF ENZYME REACTION COMPONENTS
The examples chosen to ilhistrate transport inhibition do not always
involve analogs, except in the most general sense, but clearly demonstrate
the importance of considering such a type of interference whenever analogs
are used in cellular preparations.
ANALOGS WHICH ARE ISOMERS OF SUBSTRATES
The behavior of the isomers of normal metabolites, particularly optical
isomers, constitutes, in a way, a special field and therefore some of the more
interesting results will be discussed in this section rather than under the
specific enzymes that are involved. The concept that a proper fit of a sub-
strate to the enzyme surface is necessary for reaction implies that enzymes
will usually be stereospecific, and this has been demonstrated many times.
Since an enzyme commonly attacks only one form of an isomeric pair,
the unreactive form may be either an inhibitor or completely inert. In most
cases the unreactive form does not bind to the enzyme at all, as might be
expected from the different spatial configurations of most isomeric pairs,
and is not inhibitory.
Enantiomeric Analogs
These analogs are related to the corresponding substrates on the basis
of molecular asymmetry and the most common examples are optically
active due to an asymmetric carbon atom. Unnatural enantiomers often
exhibit no affinity for the enzymes. Indeed, it has been generally found
that D-amino acids do not interfere with the microbial growth-promoting
activity of L-amino acids, although there are exceptions. Some examples
of a lack of inhibition by optical isomers may be mentioned. The oxidation
of L-phenylalanine by the L-amino acid oxidase of Neurospora is not in-
hibited by D-phenylalanine, even when the latter is present at 500 times
the concentration of the substrate (Burton, 1951 b), and D-tryptophan
does not inhibit E. coli L-tryptophanase even though it interferes with
growth (Gooder and Happold, 1954). D-Malate is not oxidized by the malate
dehydrogenase of Mycobacterium tuberculosis nor does it inhibit the oxida-
tion of L-malate (Goldman, 1956 b). A rather unusual case is presented by
potato tyrosinase in that both l- and D-tyrosine are attacked at the same
rate, but there is no evidence of mutual inhibition, perhaps because of the
limited range of concentrations used (Spencer et al., 1956). Occasionally
a slight inhibition is noted but one which would not be of any practical
significance, as in the just detectable inhibition by D-leucine of the oxida-
tion of L-leucine by the L-amino acid oxidase of the hepatopancreas of
Cardium tuberculatum, an inhibition actually much less than exerted by
other amino acids and hence probably not specific (Roche et al., 1959).
ANALOGS WHICH ARE ISOMERS OF SUBSTRATES 269
However, the primary purpose of this section is to discuss instances in
which significant inhibition by optical isomers is observed.
The asparaginase of Mycobacterium phlei attacks only L-asparagine and
this deamidation is quite well inhibited by D-asparagine (Grossowicz and
Halpern, 1956 a). It is a particularly clear and straightforward instance
of stereomeric inhibition and a 1/v — 1/(S) plot shows it to be completely
competitive. Ai'ound 75% inhibition is produced by 40 mM D-asparagine
when L-asparagine is 10 mM. On the other hand, the asparaginases of
Bacillus coagulans and B. stearothermophilus are inhibited by D-asparagine
but not competitively (Manning and Campbell, 1957). The type of inhi-
bition is difficult to designate since the double reciprocal plots intersect
to the right of the ordinate, that is, the inhibition does not tend toward
noncompetitive kinetics. A yet more complex situation is presented in the
depression of the formation of a-amylase by D-aspartate in Pseudomonas
saccharophila, 0.2 mM blocking the protein synthesis completely (Eisen-
stadt et a]., 1959). The inhibition is readily reversed by L-aspartate and is
characterized by a fairly long lag period before inhibition is observed.
The inhibition was assvimed to be on the reaction:
L- Aspartate + IMP + GTP -> fumarate + AMP + GDP + P
thus producing an impairment in AMP synthesis, this secondarily disturbing
the formation of ATP and the activation of amino acids for protein synthesis.
Examination of this reaction in cell-free extracts showed that D-aspartate
does indeed inhibit competitively. Another type of inhibition is exhibited
by pea glutamine synthetase with L-glutamate, ammonia, and ATP as
substrates (Varner, 1960). D-Glutamate inhibits the formation of L-gluta-
mine. However, D-glutamate is also a substrate and actually has a lower
K„^ although the reaction rate is slower than with L-glutamate (^,„ for
L-glutamate is 10 mM and for D-glutamate is 2 mM). This is then an exam-
ple of a competitive inhibition between isomeric substrates. It may also
be mentioned that D-glutamate inhibits, although quite weakly, the de-
carboxylation of L-glutamate by bacterial glutamate decarboxylase (Ro-
berts, 1953).
The splitting of L-histidine by rat liver histidase is inhibited by D-histi-
dine, 11% inhibition being given by 2 mM, 36% by 12 mM, 55% by 24
mM, and 85% by 48 mM when the L-histidine is 12 mM (Edlbacher et al.,
1940). The formation of L-histidine from L-histidinol occurs in two steps:
L-Histidinol -^ L-histidinal -> L-liistidine
The enzymes catalyzing both these reactions are inhibited by D-histidinol
and D-histidinal competitively (Adams, 1955). The Ki for D-histidinol is
0.05 mM for both oxidations by a yeast preparation and it is possible that
only a single enzyme is involved.
270 2. ANALOGS OF ENZYME REACTION COMPONENTS
Germination of Bacillus cereus spores is induced by certain amino acids,
such as L-alanine, and it would appear in this case that L-alanine dehydro-
genase is essential for the activation process (O'Connor and Halvorson,
1961 b). D- Alanine and some other analogs, such as D-a-amino-w-butyrate,
inhibit the germination when it is induced by L-alanine, and also inhibit
the oxidative deamination of L-alanine and to a lesser extent other amino
acids, there being a good correlation between these two inhibitory actions
in the series of analogs used (see accompanying tabulation). The inhibition
% Inhibition of deamination
Substrate ^^ D-alanine (100 mi/)
L-Alanine 61
L-a-Amino-?i-butyrate 59
L-Norvaline 48
L-Serine 28
L-Valine 24
L-Cysteine 15
L-Isoleucine 12
L-Leucine 0
L-Phenylalanine 0
by D-alanine is, however, by no means specific for L-alanine. It is possible
that several enzymes, with different susceptibilities to D-alanine, are in-
volved in the deamination of the various amino acids.
An interesting illustration of optical specificity is provided by the 0-
phosphoserine phosphatase from chicken liver (Neuhaus and Byrne, 1960),
Both L-phosphoserine and D-phosphoserine are substrates and both l-
serine and D-serine inhibit. The L-serine is a much more potent inhibitor
(see accompanying tabulation), l- Alanine also inhibits, being between
K,
(railf)
Substrate
L-Serine
D-Serine
L-Phosphoserine
D-Phosphoserine
0.68
0.70
27
29
l- and D-serine in potency, but D-alanine does not inhibit. The inhibition
by L-serine seems to be uncompetitive from a double reciprocal plot, but
ANALOGS WHICH ARE ISOMERS OF SUBSTRATES 271
this is apparent only and the type of inhibition does not fit into the usual
classical categories. The following scheme was proposed:
E + PS ^ EPS :^ EP + S
E + P
where PS is phosphoserine. It was shown to fit the data kinetically if
k_Jki is small. This example shows well the danger of uncritically accepting
the usual interpretation of a plotting procedure since, as emphasized
previously, there are types of inhibition different from those included
in the classic formulations.
a-Chymotrypsin hydrolyzes the L-isomers of various tryptophanamides
and tyrosinamides, and these reactions are usually inhibited by the D-iso-
mers (Huang and Niemann, 1952; Manning and Niemann, 1958). When
the substrate is nicotinyl-L-tryptophanamide (Kg = 2.7 mM), the reaction
is inhibited by nicotinyl-D-tryptophanamide (K^ =1.4 mM) and a number
of other derivatives of D-tryptophanamide. The hydrolysis of several de-
rivatives of L-tyrosinamide is similarly inhibited by the D-isomers (see ac-
companying tabulation). It is to be noted that in every case the D-isomer
is bound more tightly than the L-isomer, assuming that K^ does indeed
represent a dissociation constant. The possible forces binding these sub-
stances to the enzyme will be discussed in a later section (pages 370-375).
Tyrosinamide Kg for L-isomer K^ for D-isomer
Nicotinyl-
12
9
Chloroacetyl-
27
6.5
Trifluoroacetyl-
26
20
Acetyl-
32
12
Anomeric Analogs
Michaelis and Pechstein (1914), in their early work on /5-fructofuranosi-
dase, and Michaelis and Rona (1914), studying yeast maltase inhibition,
concluded that the configuration around carbon 1 of carbohydrates (i.e.,
a- and /5-anomers) is of importance in determining the affinity of these
substances for the enzymes, since a-methylglucoside inhibits both enzymes
quite potently whereas /?-methylglucoside inhibits very little or not at
aU. The splitting of phenol-/?-glucosides by taka-/?-glucosidase is also in-
hibited by phenol-a-glucoside (Ezaki, 1940). The synthesis of polysaccha-
ride from a-D-glucose-1 -phosphate by muscle phosphorylase is not inhib-
ited by /?-D-glucose-l-phosphate; however, it is interesting that a-methyl-
272 2. ANALOGS OF ENZYME REACTION COMPONENTS
glucoside inhibits while /5-methylglucoside does not, indicating the impor-
tance of the carbon 1 configuration (Campbell et al., 1952). In general, the
/5-anomers cannot act as either substrates or inhibitors of phosphorylase.
The /^-glucuronidase of mouse liver is also stereospecific, since menthyl-
/?-glucoronide is a substrate but menthyl-a-glucuronide only a very weak
inhibitor (Levvy and Marsh, 1952).
Positional Analogs
Isomers in which a ring group is moved from one position on the ring
to another are generally not inhibitory due to the fairly marked structural
changes involved. There are exceptions, however, and one of the most
striking is the inhibition of the oxidation of p-hydroxyphenylpyruvate to
homogentisate by m-hydroxyphenylpyruvate in preparations from dog li-
ver (La Du and Zannoni, 1955). A depression of 50% is seen with 0.2 mM
and over 90% with 0.5 mM m-hydroxyphenylpyruvate when the substrate
concentration is presumably 1.2 raM, indicating a tighter binding to the
enzyme of the m-isomer. Other examples of positional isomers will be
encountered in later sections.
Geometric Isomeric Analogs
One would not expect that potent inhibitors would be found in cis
and trans pairs because of the different molecular configurations. We have
already seen that fumarate and maleate differ markedly in their reactions
with succinate dehydrogenase (page 34). The outstanding exception to
this rule is the well-known inhibition of aconitase by fraws-aconitate. This
enzyme catalyzes the interconversion of the tricarboxylates:
Citrate ^ cts-aconitate ;fi isocitrate
although perhaps cis-aconitate is not an obligatory intermediate between
citrate and isocitrate. Bernheim (1928), impressed by the results obtained
by Quastel with malonate, tested the effect of frans-aconitate on liver
"citric dehydrogenase" (the reduction of the methylene blue used in this
system was actually due to the oxidation of isocitrate formed from
citrate via aconitase) and found definite inhibition. He believed the inhi-
bition to be related to the structural similarity between citrate and trans-
aconitate, stating, "The curve obtained seems to indicate that the aconitic
acid is adsorbed on the enzyme so that part of the surface is unavailable
for citric acid." Twenty years later a thorough study of this inhibition was
made by Saffran and Prado (1949), using aconitase from pigeon breast
muscle. Both the conversion of m-aconitate to citrate and the disappearance
of citrate are inhibited by irans-aconitate. However, trans-acomta.te is
ANALOGS WHICH ARE ISOMERS OF SUBSTRATES 273
not bound as tightly to the enzyme as are the substrates. When citrate is
3.3 mM, 50% inhibition is found with 16 xnM trans-a.comta.te; since K^^
for citrate is roughly 1 mM, K, is approximately 4 mM. The inhibition is
competitive although the Ijv — 1/(S) plots are not ideal, perhaps because
of some enzyme inactivation or failure to achieve equilibrium. The inhibi-
tion of rat mammary gland aconitase seems to be somewhat more potent,
since equimolar concentrations of citrate and ^raws-aconitate lead to around
50% inhibition (Abraham et al., 1960). Studies on the stereospecificity and
deuterium transfer during reactions catalyzed by aconitase (Speyer and
Dickman, 1956; Englard and Colowick, 1957) point to a three-point at-
tachment of the tricarboxylates (and perhaps an intermediate carbonium
ion) to the apoenzyme and Fe++. The carboxyl groups in trans-aconitate
would not appear to be in such a favorable position as in m-aconitate for
the formation of this complex and this might explain the relatively weaker
binding. The effects of trans-acomtate on other enzymes have been little
investigated but it has been found to be a fairly potent inhibitor of fu-
marase, Z^ being 0.63 mM at pH 6.35 (Massey, 1953 b). The K, increases
with rise in the pH (Fig. 1-14-11) and, as with malonate, the formation of
the EI complex is exothermic at low temperatures and endothermic at
high temperatures.
It is somewhat surprising that tro ns-aconitate is a reasonably effective
inhibitor of the respiration of intact cells, inasmuch as penetration into the
cells should be difficult. The following inhibitions have been observed:
22-36% of endogenous respiration of various tumor shces and 28% of
endogenous respiration of liver slices at unspecified concentration (Wein-
house et al, 1951), 25% of Paramecium respiration at 10 mM (Holland and
Humphrey, 1953), and 40% of the ion-linked respiration of barley roots
at 20 mM (Ordin and Jacobson, 1955). However, no inhibition of the
respiration of Australorbis mince at 10 mM was reported (Weinbach,
1953). The oxygen uptake resulting from the addition of citrate or cis-
aconitate to rat liver slices is strongly inhibited by 30 mM ^raws-aconitate
(Sherman and Corley, 1952). The most complete study of respiratory in-
hibition is by Saffran and Prado (1949) with rat liver and kidney slices.
In the latter the inhibition is 27% at 2 mM and 73% at 20 mM, which is
quite comparable to malonate. The inhibition by 2 mM ^rans-aconitate
is not altered by adding malate or fumarate, but is reversed with 5 mM
citrate or cis-aconitate. In other experiments the sensitivity to trans-
aconitate is unexplainably less. The inhibition of aconitase in liver and
mammary gland homogenates leads to a fairly marked depression of the
conversion of citrate to CO2 and of acetate to fatty acids by fmws-aconitate
(Abraham et al., 1960). The synthesis of mammary fatty acids is inhibited
45% by 7.1 mM and 75% by 21.4 mM. Accumulation of citrate accompa-
nies the inhibition in kidney, liver, and tumor slices (Weinhouse et al.,
274 2. ANALOGS OF ENZYME REACTION COMPONENTS
1951; Saffran and Prado, 1949), and this is augmented by the addition of
cycle intermediates such as pyruvate or malate. These results indicate
clearly that aconitase is being inhibited intracellular ly.
Essentially nothing is known of the possible effects of ^rans-aconitate on
cellular functions. No depression of Paramecium motility is seen at 10 mM
(Holland and Humphrey, 1953). However, the active transport of ions by
barley roots is markedly reduced (Ordin and Jacobson, 1955). K+ and Br"
uptakes are inhibited 32% and 33%, respectively, by 10 milf ^raws-aco-
nitate, and 63% and 47%, respectively, by 20 mM. These inhibitions are
probably not specific but the result of the depression of respiration.
^rans- Aconitate is known to occur naturally in many plant tissues and
is abundant in sugar cane juice. It is formed from acetate-C^^ in corn tis-
sues and 95% of the aconitate which accumulates is in the trans form, it
being out of equilibrium with the cycle acids; further evidence for its com-
partmentalization during endogenous formation is provided by the fact
that it is metabolized quite readily when it is added to corn roots (Mac-
Lennan and Beevers, 1964). It was suggested by Rao and Altekar (1961)
that it may arise from ci'.s-aconitate through the mediation of an aconitate
isomerase, which they isolated from soil organisms. Some pseudomonads
are capable of metabolizing fraw^^-aconitate without previous exposure to
it and other strains can adapt to utilizing it (Altekar and Rao, 1963).
FUMARASE
Fumarase has been studied more intensively than most enzymes with
regard to interactions with competitive inhibitors, the effects of pH on
these interactions, and the nature of the active center. A generalized
representation of the bindings of fumarate and L-malate to the enzyme is
shown in Fig. 1-6-2, the pH effects are discussed in Chapter 1-14 (page
691), the apparent p^,'s for various competitive inhibitors are given in
Table 1-14-2, and the P-K'^'s of the two catalytically active sites for
fumarase and its substrate complexes are given in Table 1-14-3.
Emphasis in this section will be directed to a more accurate delineation
of the active center configuration and to a more quantitative expression
of the ways in which competitive inhibitors interact with the active center.
Fumarase possesses four important groups: two cationic groups for binding
the COO" groups of the substrate in the trans position, and two ionizable
groups interacting with the groups on the a- and /5-carbon atoms and in-
volved in the addition or removal of water. The latter enzyme groups wiU
be designated as Rl and R^ in conformity with Wigler and Alberty (1960);
each may exist in the protonated form, RlH or Rj^H. The p^^'s of these
groups, which are 6.3 and 6.9 in the free enzyme, point to their phenolic
or imidazole nature; indeed, it is possible that these two groups are identical,
rUMARASE
275
but evidence against this comes from the study of inhibition by the tar-
trates. The catalytically active form of fumarase may be represented by
EH, in which one of these groups is protonated, although both E and EHg
are capable of binding both substrates and inhibitors.
The values of K^ for several competitive inhibitors at pH 6.35 and 23°
(Table 2-1) may be used as a rough and provisional means of evaluating the
relative energies of ])inding, bearing in mind that these are apparent K-&,
that the enzyme exists in three different ionized states for each of which
the binding is different (so that the K^s are in a sense composite), and
that the various inhibitors alter the pA^^'s of the enzyme groups in different
ways in the EI complexes. However, some reasonable conclusions may
be drawn from these AF values.
Table 2-1
Inhibitor Constants" and Relative Binding Energies
FOR Competitive Inhibitors of Fumarase
Inhibitor
Apparent K^
(mM)
Relative —AF
(kcal/mole)
Adipate
Succinate
Glutarate
Malonate
D -Tartrate
Mesaconate
Maleate
L-a-Hydroxy-/5-sulfopropionate *
D-Malate
Citrate
<rans-Aconitate
100
1.35
52
1.73
46
1.80
40
1.90
25
2.16
25
2.16
11
2.64
10
2.70
6.3
2.97
3.5
3.32
0.63
4.32
° Values of Ki determined at pH 6.35 and 23°.
* The Ki for L-a-hydroxy-^-sulfopropionate was changed from 16.5 mM as given
in the table (Massey, 1953 b) to correspond to the value in the curve presented (16.5
is probably a misprint for 10.5.)
[A) Since all monocarboxylates and the methyl ester of fumarate are
without inhibitory activity, it must be assumed that at least two negatively
charged groups are necessary for binding. However, it is evident that for
the more tightly bound substances other attraction forces are involved. If
we assume that these additional forces arise from hydrogen bonding be-
276 2. ANALOGS OF ENZYME REACTION COMPONENTS
tween hydroxyl groups and the Rl and R^ enzyme groups, polarization
of double bonds, and interactions of a third C00~ group (in the tricar-
boxylates), the tentative values shown in the following tabulation may be
assigned for the contributions made by the various interactions to the
total binding:
Two C00~ groups 1.75 kcal/mole
— OH group hydrogen bonding 0.5-1.5 kcal/mole
— C=C — polarization 1.9 kcal/mole
Additional CHgCOO- group 0.5 kcal/mole
The hydrogen bonding and polarization values are minimal since, in part,
they were derived from the K„'s of the substrates (for fumarate K„^ =
= 1.78 mM and — JF = 3.71 kcal/mole, and for L-malate Z,„ == 4.0 mM
and — AF = 3.24 kcal/mole). The ^„/s may not represent dissociation
constants but in any case the true K^'s would be equivalent to or smaller
than the K„'s, so that the binding energies for the substrates may be
somewhat higher. The cis configuration of maleate reduces the attraction,
but the value for maleate in the table should be corrected since the pK^
— 5.9 (at 23° and around 0.1 ionic strength) and only 74% of the total
maleic acid would be in the form of maleate=: this increases — JF to 2.82
kcal/mole. The difference in binding between fumarate and maleate is
thus at least 0.9 kcal/mole. It is also interesting that the introduction of
a methyl group into fumarate to form mesaconate brings about a 1.55
kcal/mole or greater reduction in the binding energy, resulting possibly
from a steric displacement and lowered polarization interaction. The 1.9
kcal/mole estimated for electrical polarization of the double bond is not
unreasonable and actually corresponds fairly closely to that calculated,
using appropriate molar refractions and an interaction distance of 4 A.
A factor of unknown importance is the possible deformation of the dicar-
boxylates to fit the active site and the energies that would be involved
with the different inhibitors.
(B) If these conclusions are valid, the interaction energy for fumarate is
approximately half due to coulombic ion-ion forces and half due to the
inductive polarization by a strong dipole. It is possible that one of the R
groups on the enzyme is positively charged and the other negatively
charged on the active enzyme, as suggested by Massey (1953 b). The ionic
interactions serve to orient the fumarate at the active center, and the
polarization not only stabilizes the complex but initiates the addition of
water.
(C) The third COO" group of citrate and trans-aconitate seems to be
able to interact with an adjacent positive group on the enzyme, but rela-
FUMARASE 277
tively weakly (0.3-0.6 kcal/mole). In fact, the low interaction energies of the
terminal COO" groups (0.87 kcal/mole for each group, which may be com-
pared with the 3.3-3.6 kcal/mole binding per COO" group of malonate or
succinate on succinate dehydrogenase) might indicate that the distance
between them and the enzyme cationic groups is relatively great (perhaps
12-15 A), or could even point to a type of interaction other than ion-ion
attraction.
(D) L-a-Hydroxy-/?-sulfopropionate is bound at least 0.54 kcal/mole less
tightly than L-malate, much of the affinity of this analog resulting from
the OH group interaction. This would indicate that the sulfonate group
is not a very good substitute for a COO" group in this case. It would be
interesting, in this connection, to have inhibition data on L-/5-sulfopropio-
nate.
(E) When one turns to the effects of pH on the binding of these inhi-
bitors, it is evident that the situation is more complex than assumed from
the data at a single pH (see Fig. 1-14-11). Several inhibitors exhibit a pro-
gressive decline in binding with increase in the pH, but in the case of fu-
marate the pZ,„ rises between pH 7 and 8, and the pZ,„-pH curve for l-
malate shows several changes of slope. The inhibitor D-malate also shows
an increase in binding between pH 7 and 8. Massey (1953 b) suggested that
binding to different sites might be involved. However, since deviant be-
havior is noted with substances containing a double bond or OH group,
it is possible that the affects of pH on the polarization and hydrogen bonding
interactions may be involved. Succinate exhibits a linear decrease in jiK^
with pH whereas fumarate behaves quite differently. Mesaconate and ma-
leate have succinate-type pH dependences and it is possible that the pola-
rization interaction is sterically prevented in these substances, as postulated
above. Apparently some change in the active center ionization occurs
between pH 7 and 8 which alters these interactions, and we shall return to
this problem later when more recent inhibition data have been presented.
(If the crude approach to these problems in the preceding paragraphs
serves either to irritate or activate others to further theoretical or experi-
mental study, a purpose will have been accomplished.)
One would expect the most potent competitive inhibitors of fumarase
to have either a polarizable group (such as — C=C — ) or a group capable
of forming hydrogen bonds (such as OH). Substitution of groups at the
double bond of fumarate seems to reduce the binding, and acetylene-di-
carboxylate has not been studied. Thus one is left with the tartrates as
possibly interesting inhibitors, and they were investigated by Wigler and
Alberty (1960) in an excellent study designed to establish the more intimate
nature of the catalysis. The variation of the inhibitions with pH allowed the
determination of the p^^'s of the enzyme groups, the changes in these pro-
duced by complex ing with the inhibitors, and the dissociation constants
278 2. ANALOGS OF ENZYME REACTION COMPONENTS
COO
H^i .H
^C^
1
COO'
1
COO
H. i OH
1
COO
H^ i .OH
^C^
1
1
H-^^^H
coo-
1
H-^ ; OH
coo-
1
HO^ i ^H
coo-
1
Q
U^ 1 ^OH
coo-
Succinate
L-Tartrate
D -Tartrate
wieso- Tartrate
of the inhibitors with the variously protonated forms of the enzyme. It
was found that meso-tartrate is the most potent inhibitor of this group
and it was concluded that this configuration of OH groups allows the
formation of two hydrogen bonds with the Rl ^^^ ^d enzyme groups
(Fig. 2-2). The singly protonated form of the enzyme, EH, binds the
C-—C FUMARATE
/ \
Rp-H COO 0
©
ooc
Rjj-H
L-MALATE
MESO- TARTRATE
Fig. 2-2. Simplified scheme of the fumarase active site described by Wigler and
Alberty (1960). The cationic, R^, and R^ groups occur on different levels and are so
located they can interact with certain isomers of substrates and inhibitors.
meso-tartrate most tightly. The binding energies contributed by the hy-
drogen bonds can be estimated from the relative interactions of succinate
and the tartrates. Weak hydrogen bonds are indicated for the doubly pro-
tonated form of the enzyme, EHgl, with D- and L-tartrates, while stronger
bonds (— 2.8 to — 3.3 kcal/mole) are formed with meso-tartrate and the
less protonated enzyme (see tabulation).
INHIBITION OF XANTHINE OXIDASE 279
AF for displacement of succinate
(kcal/mole)
EH2I EHI EI
D-Tartrate —0.5 0 +0.4
L-Tartrate —0.5 +0.8 +0.7
meso-Tartrate — 1.6 — 2.8 — 3.3
The fumarase from, liver is inhibited differently from the heart enzyme,
in that mesaconate (methylfumarate) is inactive whereas citraconate
H3C ^COO' H COO" V*^
C C C=CH2
II II I '
/C^ . ^C^ CH3
H COO H3C H i,^-
Citraconate Crotonate Itaconate
(methylmaleate) inhibits well (Jacobsohn, 1953). Itaconate inhibits less
potently and crotonate even less potently (about the same as succinate).
The fact that crotonate inhibits at all is interesting, in that it suggests
that one C00~ is sufficient if a double bond is present. czs-Aconitate and
^raws-aconitate inhibit equally and weakly. DL-/5-Fluoromalate inhibits com-
petitively the conversion of malate to fumarate by fumarase (Krasna,
1961). Assuming that both optical isomers inhibit equally (which may
not be true), K^ = 28 mM, and K„^ for malate is 3.5 niM. The inhibition
of malate dehydrogenase is much stronger, if ^ being 0.16 mM, which may
be compared to a K,,^ of 11 mM for malate.
INHIBITION OF XANTHINE OXIDASE
BY PURINE ANALOGS AND PTERIDINES
Some potent and specific inhibitors of xanthine oxidase have been dis-
covered and have proved to be interesting not only on the enzyme level
but because of the disturbances in purine metabolism produced in whole
animals. Xanthine oxidase catalyzes the oxidation of hypoxanthine and
H3C ^coo
^c
II
OOC H
Mesaconate
0
1 1
H
1
H
Xanthine
N N'
Hypoxanthine Xanthine Uric acid
280
2. ANALOGS OF ENZYME REACTION COMPONENTS
xanthine to uric acid, each step essentially involving the addition of water
and the removal of two hydrogen atoms which are transferred to oxygen
along a typical electron transport sequence. Hydroxypurines exhibit keto-
enol tautomerism, and it is only recently that spectroscopic evidence point-
ing to the predominance of the keto form has been obtained (Mason,
1957). The structures of the purines and pteridines to be discussed will
be written as far as possible in conformity to these results. However, in
some cases it is difficult to assign the most important structure, especially
in multiply substituted compounds. Ionization may also be a complicating
factor. Most purines appear to be predominantly neutral at physiological
pH (piiC^ between 1 and 4; pK^, between 8 and 12), but some, such as uric
acid, lose a proton in the acid pH range and physiologically exist as anions
(e.g., pK^ for uric acid is 5.4). The site of loss of the proton is not known
and it is quite possible that the anions should be considered as being
equilibrium mixtures of several structures. The form in which they are
written, hence, does not imply that this structure is the only one present,
or even that it is necessarily the most important structure. Such considera-
tions become important in treating the forces between these compounds
and the enzyme. A final factor must be borne in mind: the most stable
form in solution is possibly not the form in which the purine is bound to
the enzyme surface, inasmuch as the interaction may modify the structure
appreciably.
Purine Analogs
Dixon and Tliurlow (1924) reported that xanthine oxidase is inhibited
by various purines, such as adenine and uric acid, but no quantitative data
were given. There was no further investigation until the introduction of
the azapurines. 2-Azaadenine and 2-azahypoxanthine, like many purine
Purine
Guanine
8-Azaguanine
NH.
NH,
NH.
Pyrazoloisoguanine
INHIBITION OF XANTHINE OXIDASE 281
analogs, are oxidized by the enzyme in the 8-position (Shaw and Woolley,
1952). Eqiiimolar concentrations of 2-azaadenine prolong the formation of
urate from xanthine 2-fold and from hypoxanthine 4-fold, the inhibition
being competitive. The kinetics in such situations may be complicated
by two factors: (1) the disappearance of the inhibitor (Shaw and Woolley
found, for example, that the azaadenine essentially all disappeared before
much urate was formed), and (2) the inhibition produced by the product
of the inhibitor oxidation. 8-Azapurine and all of its monohydroxyl and
monoamino derivatives are oxidized by xanthine oxidase and the products
are frequently inhibitory not only to xanthine oxidase but to other enzymes.
2-Amino-8-azapurine is converted to 8-azaguanine and hence can be used
as a precursor of this inhibitor (Bergmann et al., 1959).
Certain inhibitions of xanthine oxidase by purine compounds are sum-
marized in Table 2.2 The inhibitions are not always competitive despite
the close similarity of substrate and inhibitor structures. Some of the simple
analogs are bound more tightly to the enzyme than are the normal sub-
strates. 6-Chloropurine and pyrazoloisoguanine are bound particularly well
and this brings up questions regarding the forces involved. Very little is
known about these forces. Ionic forces must be unimportant and it is pos-
sible that hydrogen bonds, coupled with an appropriate fit of the bonding
groups, play a major role. PjTazoloisoguanine is the 4-amino-6-hydroxy
derivative of pyrazolopyrimidine, and it is interesting to note that the
4-amino derivative is a very weak inhibitor relatively, as are the 4-methyl-
amino and l-methyl-4-amino derivatives (Feigelson et al., 1957). It has
been stated that there is some correlation between the potency of the xan-
thine oxidase inhibition and the carcinostatic activity of these and related
compounds.
When inhibitory purine analogs are administered to animals it is often
difficult to determine the toxic mechanisms because of the multiple possible
sites for interference. The biological effects of 6-mercaptopurine seem to
be related to its conversion to the ribonucleotide, which inhibits inosinic
acid metabolism, rather than to any direct enzyme inhibition (Silberman
and Wyngaarden, 1961). On the other hand, it has been postulated that
8-azaguanine induces a guanine deficiency by inhibiting xanthine oxidase,
which operates in one guanine biosjoithetic pathway (i.e., hypoxanthine
-^ xanthine -^ guanine) (Feigelson and Davidson, 1956 a). It has been
shown in one instance that purine metabolism can be inhibited in vivo.
6-Chloropurine given to rats at 80 mg kg inhibits the formation of C^^Og
from xanthine-6-C^* about 40% when administered 20 min before the xan-
thine (Duggan et al., 1961). This is probably not due to a direct action
on xanthine oxidase but to the formation of 6-chlorourate and the resulting
inhibition of uricase. 6-Chloropurine also depresses the conversion of ace-
tate to lipid, of glycine to protein, and nucleic acid synthesis.
282
2. ANALOGS OF ENZYME REACTION COMPONENTS
P5
O ^
6 6
^ ^
o §
+
CO '^
tH c :r cr
112 1^
■^ ^ o
c3 c3 ^ 'i)
e c g c c s
O TO Co CO o f
C U) M M c ^
§ 5 5 5 § -2
^ C C
™ Oi (D
o3 c«
cS cS
bC bD
C C
^pq hJQQPh-1S^^
S fc — I SO s S ;::!
-ti -ti +i
!^ ^ ^
,ifO -< o
I I
+ + i
.:- .- (D
.S ^ . " — ^
773 O S) ■:3 773
^ c3 ce
§ 05 P^
O S
m
2 5
(U
:a ^
111
cS
CK pJH PQ
"-'do
t< t<
+
f=^
Q
s i^
o J4 M M
13 ^ ^ " ^
?? fer* ^T-f w P-
O
§
.g
§
^
^
s
s
s£
1^ -^
-«
"^
;2 ^
f^ T^
772
o
Si rs
" g
S
O
" §
u ^ tl,
o
o §
'S 'S >> ^
c c X o
cs ce o ,i2
5 5 ^ >. •£
<D
- -3 -^
0) cS c
■d 3 cS
c3 bC X
N N SI
>> K 3 oi <; <c
'-* & . I I
C-] GO 00
:s a
^ .2
"5 .-:<
INHIBITION OF XANTHINE OXIDASE 283
A new xanthine oxidase competitive inhibitor, 4-hydroxypyTazolo(3,4-d)
pyrimidine (allopurinol, Zyloprim), is now being clinically tested in hyper-
uricemia and for the potentiation of the antitumor activity of the 6-substi-
tuted purines (e.g. 6-mercaptopurine) (Elion et al., 1963; Information for
Investigators report from the Burroughs Wellcome Company). It is a
N
I
H
4-Hydroxypyrazolo(3, 4-d)pyrimidine
very potent inhibitor, with K^ = 0.000032 mM, being bound around 100
times more tightly to the enzyme than is xanthine, but it is also a substrate
for xanthine oxidase and its oxidation product is likewise a potent inhib-
itor, with Ki = 0.000054 mM. Mice and dogs given 100 mg/kg intraperi-
toneally show an increased urinary excretion of xanthine and hypoxanthine,
with a decrease in aUantoin, and in man a similar action has been demon-
strated, serum and urinary urate being depressed. It is well tolerated by
man at 200-1000 mg/day orally up to several weeks. It apparently has no
antitumor activity itself but is able to potentiate the action of 6-mercapto-
purine by interfering with its metabolism. Mice given 20 mg/kg intraperi-
toneally along with 6-mercaptopurine exhibit a reduced urinary excretion
of thiourate. The value of this analog in neoplastic disease and gout is
not yet known.
Inhibition of Uricase by Purine Analogs
It is appropriate at this time to refer to certain studies on uricase (urate
oxidase) before proceeding with the inhibition of xanthine oxidase by the
pteridines. Uricase catalyzes the oxidative opening of the pyrimidine ring
to form aUantoin. Many methyl and ethyl derivatives of urate were tested
by Keilin and Hartree (1936) on an enzyme from pig liver; they found that
none is a substrate but that several inhibit quite potently. They considered
the mechanism to be competitive and stated, "The fact that the methyl
compounds of uric acid, although not oxidizable by the enzyme, inhibit
the oxidation of uric acid shows that these methyl compounds react with
the same active grouping of the enzyme molecule as uric acid itself." It
is very interesting that the 1,3,7-derivative is so much more potent than
the 1,3,9-derivative, particularly in view of the greater potency of the
monomethyl compounds compared to the latter derivative (the urate was
10 mM in all cases) (see tabulation). 2,6,8-Trisubstituted purines were
284 2. ANALOGS OF ENZYME REACTION COMPONENTS
Inhibitor
Concentration
(mif)
%
Inhibition
1,3,9-Trimethylurate
7.7
16
7-Methylurate
9
41
3-methylurate
9
49
1-methylurate
9
58
1,3,7-Trimethylurate
7.7
68
studied by Mahler et at. (1956) (see tabulation) and the results give some
information on the nature of the binding to the active site. Three binding
sites were recognized: (1) a cationic group binding the 2-substituent,
Substituent in position:
A', {mM)
2
6
8
CI
CI
CI
0.0008
CI
CI
OH
0.0013
OH
NH,
NH2
0.0018
OH
OH
H
0.012
OH
OH
OH
0.025 (A', for urate)
CI
NHa
OH
0.04
OH
NHa
OH
0.15
NH2
NHa
OH
0.5
NH2
OH
NH2
No inhibition
NH2
NH,
XH,
\o inhibition
(2) a neutral group binding the 8-substituent, and (3) the copper which
chelates with the 6- substituent and the 7-N atom. It is difficult to under-
stand on this basis the high affinity of the trichlorourate for the enzyme,
since one would predict that substitution of chlorine in the 2-position would
reduce binding to the cationic group and substitution in the 6-position would
interfere with the chelation. The increased binding produced by chlorine
substitution might be in part the result of a stabilization of conjugative
resonance and an increased hydrogen bonding (and perhaps an increased
chelation of two N atoms with the copper). Also resonance with structures
in which one or more of the chlorine atoms are in the form
C1+
— N-— C—
would produce strong dipoles. The most potent inhibitor apparently is
6-chloro-2,8-dihydroxy purine (usually misnamed 6-chlorourate), which is
INHIBITION OF XANTHINE OXIDASE 285
formed from 6-chloropurine by the action of xanthine oxidase, K^ being
0.00006 mif (Duggan and Titus, 1959). It inhibits urate degradation in
the rat 75% at a dose of 20 mg/kg (Duggan et al., 1961). The over all AF
for the binding to uricase is approximately 10 kcal/mole and this would
indicate the formation of some type of stable bond. It was pointed out that
this analog is very stable and might be useful in studying the metabolism
and disposition of urate. It should induce urate accumulation in the tissues
and the effects of this might have some bearing on the manifestations of
gout. The inhibition of uricase by xanthine (2,6-dihydroxypurine) in the
tabulation above is interesting because it illustrates a novel effect in a multi-
enzyme system. In the sequence:
xanthine
oxidase uricase
Xanthine > urate > allantoin
the initial substrate inhibits the second enzyme in the series, causing ac-
cumulation of urate in increasing amounts as the xanthine concentration
rises (e.g., in rat liver homogenates). At high concentrations the urate con-
centration falls due to the substrate inhibition of xanthine oxidase (Van
Pilsun, 1953).
Further studies on uricase have been reported by Bergmann et al. (1963 a)
and some of the results, including calculations of the approximate apparent
relative binding energies, are presented in the following tabulation. The
xV-methylpurines are relatively weak inhibitors and are not included in
the tabulation. It is clear that the best inhibitors contain a 2-OH group
(designated as OH for convenience but the keto form is probably dominant),
and this position is the most important in the binding; substitution of the
2-0II with a 2-SH group lowers the inhibitory activity markedly. The
weakening of the binding by A^-substitution points to the importance of
the imino group for attachment. The inhibitions are generally" sensitive to
the pH and for most analogs increase with a rise in pH, although for 6-
SH-8-OH-purine there is a decrease, and with 8-OH-purine there is no
effect. The pH probably influences the tautomerism, which is quite impor-
tant since the binding depends on the states of the N and OH groups. The
K^ for 2,8-diOH-6-SH-purine is 0.0026 niM and for 2,6-diOH-8-SH-purine
is 0.00039 mM (Bergmann et al, 1963 b). The 2-OH-6,8-diSH-purine (which
is 6,8-dithiourate) is a much more potent inhibitor and, although its K^
was not given, it must be at least around 0.00004 mil/.
Pteridines
The inhibition of xanthine oxidase by synthetic folate, reported by
Kalckar and Klenow in 1948, was soon found to be due to some impurity,
and the simultaneous observation, by Lowry and Bessey, of the very po-
2. ANALOGS OF ENZYME REACTION COMPONENTS
Substituent in position:
(1)50 (mif) Relative —AF (kcal/mole)
2
6
8
OH
CH3S
OH
0.00013
6.80
OH
SH
SH
0.0004
6.12
OH
H
Aza
0.0016
5.25
OH
SH
H
0.0027
4.94
OH
OH
SH
0.0050
4.55
OH
H
OH
0.0052
4.52
OH
OH
Aza
0.0059
4.45
OH
CH3S
H
0.0060
4.44
OH
H
H
0.012
4.01
OH
H
OH
0.012
4.01
OH
SH
OH
0.014
3.91
OH
CH3
H
0.017
3.78
OH
OH
H
0.018
3.76
H
CH3S
OH
0.032
3.40
OH
OH
CH3S
0.038
3.30
H
OH
OH
0.066
2.95
H
SH
OH
0.070
2.91
SH
OH
SH
0.080
2.84
H
H
OH
0.11
2.65
SH
SH
OH
0.15
2.44
SH
OH
H
0.19
2.30
H
OH
H
0.22
2.21
SH
OH
OH
0.25
2.13
SH
H
OH
0.50
1.70
H
SH
OH
0.50
1.70
H
SH
H
0.70
1.19
tent inhibition produced by 2-amino-4-hydroxy-6-pteridyl aldehyde, a
photolytic product of folate, led to the conclusion, subsequently verified,
that this was the contaminant (Kalckar et al., 1948; Lowry et al., 1949 a,
b). For convenience we shall follow Hofstee (1949) in designating 2-amino-
4-hydroxypteridine as pterin and the inhibitor thus as pterin-6 -aldehyde*
Pterin and many of its substituted derivatives are oxidized to varying
degrees by xanthine oxidase while other derivatives are only inhibitory.
* This substance has been variably called 2-amino-4-hydroxy-6-pteridyl aldehyde,
2-amino-4-hydroxy-6-formylpteridine, 6-formylpteridine, pteridylaldehyde, 2-amino-
4-hydroxy-6-pteridine carboxaldehyde, and 2-amino-4-hydroxy-6-formylpterine, in
most cases without either justification or accuracy.
INHIBITION OF XANTHINE OXIDASE
O
H,N
287
H,N
H,N
«^N^
1 "^
HsN-^N-^
^N^
Pterin-6-
-aldehyde
O
H
1
H-nA
^Ny-o
H2N^^^N-^
1
1
H
Xanthopterin Isoxanthopterin Leucopterin
Actually all the photolytic oxidation products of folate are inhibitory
but pterin-6-aldehyde, the primary product:
Folate -> pterin-6-aldehyde -> pterin-6-carboxylate -> pterin -> isoxanthopterin
is by far the most potent; indeed, it is one of the most potent inhibitors
known.
Inhibition of xanthine oxidase by pterin-6-aldehyde is observed at a
concentration of 2 X 10^* //g/ml or roughly 10"^ M (Lowry et at., 1949 a).
Competitive inhibition with respect to both xanthine and pterin has been
demonstrated, and a /iC, of 6 x 10"' mM calculated for the milk enzyme
(Lowry ef al., 1949b). It was shown that 35% inhibition occurs when enzyme-
FAD = 9.3 X 10-6 jnM, the substrate pterin = 78 x 10-« mM, and pte-
rin-6-aldehyde = 2.26 X 10-« mM, this indicating that 2.26 X 10-« mM
inhibitor completely blocks 3.3 X 10-^ mM enzyme (on the basis of FAD
content). It may have been that all the FAD was not catalytically active
or that more than one FAD molecule was associated with one enzyme
molecule. However, there is no doubt that this is a mutual depletion system
and that pterin-6-aldehyde titrates the enzyme. A few other reports will
be mentioned to illustrate the potency. Both the milk and rat liver enzymes
are inhibited, 40-50% inhibition occurring at 5-8 X 10"^ mM when xan-
thine is 0.07 mM (Kalckar et al, 1950). The xanthine oxidase from Clostri-
dium cylindrosporum is potently inhibited by 0.0002 mM (Bradshaw and
Barker, 1960). Milk xanthine oxidase is completely inhibited by pterin-6-
aldehyde at 0.0033 mM when hypoxanthine is 3.33 mM (Petering and
Schmitt, 1950), and the rat intestine enzyme is inhibited completely by
0.067 mM when hypoxanthine is 6.6 mM (Westerfeld and Richert, 1952).
In many experiments the substrate concentrations have been unnecessarily
high since maximal rates are usually obtained at concentrations well below
288 2. ANALOGS OF ENZYME REACTION COMPONENTS
0.1 niM, and hence the true potency of the inhibition has been somewhat
obscured. Pterin-6-aldehyde is actually oxidized very slowly by xanthine
oxidase and reversal of the inhibition develops gradually. The oxidations
of aldehydes (Kalckar et al., 1950) and sulfite (Fridovich and Handler,
1957) by xanthine oxidase are also inhibited by pterin-6-aldehyde, but the
oxidation of NADH is not affected (Lowry et al., 1949 b).
Although it has generally been stated that the inhibition by pterin-6-
aldehyde is competitive, the Ijv — 1/(S) plots are not linear (Bradshaw
and Barker, 1960), and others (Hofstee, 1949) have presented only quali-
tative evidence for competition. Deviations from the classic kinetics might
be expected because of the mutual depletion of free enzyme and inhibitor
concentrations, the difficulty in achieving true equilibrium, and the oxida-
tive removal of the inhibitor. There is no evidence against a competitive
mechanism, however, and competition has been more clearly demonstrated
for some other pteridines.
The inhibition by pterin-6-aldehyde is quite specific although it must
be admitted that not many enzymes have been tested. Uricase, glucose
oxidase, and 3-phosphoglyceraldehyde dehydrogenase are not affected, but
the quinine oxidase of liver is inhibited (Kalckar et al. 1950; Villela, 1963).
Mouse liver guanase, using 8-azaguanine as a substrate, is inhibited but
not potently when the substrate is 11 mM and 30-min preincubation is
allowed (see accompanying tabulation) (Shapiro et al., 1952). In fact,
Pterin-b-aldehyde n, n ■ i i x-
/n Guanase inhibition
(mi/)
1 32
6 62
11 80
16 88
xanthopterin is a more potent and more rapidly acting inhibitor (Dietrich
and Shapiro, 1953 b). Pterin-6-aldehyde is not carcinostatic itself, but
potentiates the action of 8-azaguanine and a suppression of 8-azaguanine
destruction was claimed as the mechanism, although the relatively low
inhibitory potency coupled with the low doses (20 mg/kg) necessary makes
it difficult to accept this explanation. Byers (1952) investigated the effects
of pterin-6-aldehyde injections in rats (200 mg/kg intraperitoneally) on
the tissue urate levels and found no significant changes, which might in-
dicate a rapid destruction of the inhibitor in the animal. Daily injections
of 30 //g pterin-6-aldehyde in chicks also does not alter liver xanthine oxi-
dase activity despite the potent inhibition in vitro (Dietrich et al., 1952).
Other pteridines, although less potent than pterin-6-aldehyde, are nev-
INHIBITION OF XANTHINE OXIDASE 289
ertheless effective inhibitors of xanthine oxidase. The following tabulation
shows the inhibitions after 20 min incubation at pH 8.5 when the substrate
Inhibitor
Concentration
(mM)
% Inhibition
Pterin-6-aldehyde
0.031
100
Pteroate
0.033
100
Xanthopterin
0.033
72
Isoxanthopterin
0.066
95
Leucopterin
0.032
34
7-methylxanthopterin
0.077
91
6-MethyUsoxanthopteiin
0.077
89
Xanthopterin-7-carboxylate
0.050
62
Isoxanthopterin-6-carboxylate
0.075
23
Pterin-6-carboxylate
0.055
15
2,4-Diamino-6,7-dihydroxypteridine
0.040
76
concentration is 0.063 mM (Hofstee, 1949). The potency relative to pterin-
6-aldehyde is not seen here since 0.001 mM inhibits 82% under these con-
ditions. From the K„, for xanthine of 0.02 mM (Hofstee, 1955), a K^ for
pterin-6-aldehyde of 5.1 X 10"^ mM may be calculated, which is higher
than was obtained by Lowry et al. (1949 b). The inhibition by xanthopterin
appears to be completely competitive (i^,„ for xanthine 0.0053 milf , and K,
= 0.0016 mM), and xanthopterin-7-carboxylate is of comparable potency
(Krebs and Norris, 1949). Bovine serum xanthine oxidase is inhibited 94%
by 0.052 mM pterin-6-aldehyde but only 3% by xanthopterin at a com-
parable concentration (Villela et al., 1956). There is no doubt that the al-
dehyde group at the 6-position confers a strong affinity for the enzyme,
since when it is reduced to a CHgOH group, oxidized to a COO" group,
or altered to a CHg group, the inhibitory activity is markedly reduced (Pe-
tering and Schmitt, 1950). Pterin-6-aldehyde is bound to xanthine oxidase
approximately 4.9 kcal/mole more tightly than xanthopterin, and it would
appear that pterin itself is bound somewhat less tightly than xanthopterin.
It is tempting to relate the augmenting action of the aldehyde group to
the fact that xanthine oxidase oxidizes simple aldehydes. There must be
a site on the enzyme capable of reacting with aldehyde groups, and it is
possible that the pterin-6-aldehyde is bound in a configuration such that
the aldehyde group interacts in this manner. Support for this interpretation
comes from the observation by Lowry et al. (1949 b) that pterin-6-aldehyde
is slowly oxidized to pterin-6-carboxylate by xanthine oxidase.
290
2. ANALOGS OF ENZYME REACTION COMPONENTS
CHOLINE OXIDASE
The inhibitions of liver choline oxidase reported by Wells (1954) (Table
2-3) recall the interferences exerted by these and similar analogs on cho-
linesterase and tissue acetylcholine receptor groups. It would appear that
a certain critical distance between the N+ group and the terminal CH2OH
Table 2-3
Inhibition of Rat Liver Choline Oxidase by Choline Analogs"
Inhibitor
Relative
rate of
oxidation
Equimolar
%
inhibition
Relative
AF of
Series
Ri
R2
R3
binding
(kcal/mole)
Ethanolamine
H
H
H
0
23
0
H
Me
Me
0
53
- 0.81
R
H
Et
Et
0
25
- 0.06
R-N^^CHoCHgOH
R^
Me
Me
Me
Et
Et
Et
79
24
-
-
Et
Et
Et
0
11
+ 0.55
3- Aminopropanol- 1
H
H
H
0
25
- 0.06
R
Me
Me
Me
5
15
+ 0.33
R— N^!— CH2CH2CH2OH
R
H
Me
Et
Et
Et
Et
0
8
18
7
+ 0.19
+ 0.88
Et
Et
Et
0
14
+ 0.38
1- Aminopropanol -2
Me
Me
Me
66
-
-
R CH3
R— N^— CH.— CH- OH
R
H
Me
Et
Et
Et
Et
Et
Et
Et
0
14
4
13
6
16
+ 0.43
+ 0.96
+ 0.29
2-Amino-2-methylpropanol-l
R CH,
R— N^— C-CH2OH
/ 1
R CH3
H
Me
Et
H
Me
Et
H
Me
Et
4
105
8
64
65
- 1.09
- 1.12
2 - Amino - 2 - methy Ipropanediol
R CH,
R— N^-C-CHjOH
R CH.OH
H
Me
Et
H
Me
Et
H
Me
Et
3
41
5
41
50
- 0.51
- 0.74
"Experiments done with rat liver homogenates. Choline and all analogs at 37.5 mM. The
rates of oxidation are given relative to that for choline (100). (From Wells, 1954.)
INHIBITION OF NITROGEN FIXATION BY OTHER GASES 291
group is necessary for substrate activity. The most reactive substrates
are dimethyl or trimethyl compounds, indicating that an exact fit of the
cationic head is necessary to place the hydroxyl group in position for
oxidation. Substitution of groups on the C-1 atom reduces the binding
whereas substitution on the C-2 atom increases the affinity even though
the groups are fairly bulky. The simplest interpretation is that the cationic
head anchors the molecules in position so that the CH2OH group can react
with an enzyme group on the opposite side of a hole or slit in the protein.
When the analogs are too long they do not readily fit into this region,
whereas groups protruding from C-2 interact by van der Waals' forces
with the walls of the cavity. A three-point attachment of the cationic head
is suggested by the reduction in affinity brought about by altering only
one of the R groups, this perhaps tilting the molecules so that the hydro-
carbon chain is not in the normal direction. The differences in binding
energies between these analogs are rather small and this might indicate
that dispersion forces are mainly involved, but it may also be that changes
in the electrostatic interactions (resulting from the different volumes of
the R groups, for example) are offset by opposite changes in the dispersion
energy. Since choline must be oxidized to betaine before it can serve as
a methyl donor, it is interesting that Wells demonstrated the inhibition
of methionine synthesis in liver homogenates by 2-amino-2-methyl-l-
propanol and its triethyl derivative. Niemer and Kohler (1957) studied
analogs of choline in which one of the methyl groups is substituted by var-
ious radicals (e.g., — CHoCH.^OH, — CHaCH^Br, — CH2CH=CH2, — CH2=
=CH2, and —CH2COO-) and found 10-20% inhibition of liver choline
oxidase at concentrations approximately equimolar with choline (11.5 mM).
None of these analogs is a potent inhibitor, confirming the importance of
fit at the cationic head.
INHIBITION OF NITROGEN FIXATION BY OTHER GASES
Some simple instances of competitive interference between gases in
nitrogen fixation and hydrogen evolution have been observed, and are
reminiscent of the suppression of hemoglobin oxygenation by carbon mon-
oxide, nitric oxide, and other gases. The primary product of nitrogen
fixation in microorganisms is probably ammonia, and the enzyme system
responsible for this is generally termed nitrogenase. A few examples of
inhibition are given in Table 2-4. Most of these have been shown to be
strictly competitive. CO and NO are the most potent inhibitors while H2
and NgO are relatively weak. O2 is a special case in that as pOg is increased
from zero the nitrogen fixation accelerates, but above a certain value,
depending on the organism, the rate falls off (Burris, 1956). Ethane, neon,
argon, and helium have no significant effects (Molnar et al., 1948).
292
2. ANALOGS OF ENZYME REACTION COMPONENTS
03
O
Zl-
05
05
3
-ti
cS
Tin
oo
3
C
C5
05
40
^
o
03
o
^
05
cS
lO
73
lO
"e
'e
05
c
T3
Oi
^
o
^
c«
:^
«u
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INHIBITION OF NITROGEN FIXATION BY OTHER GASES 293
Inhibition of Azotohacter growth by N2O occurs if N2 is the sole source
of nitrogen, but does not if ammonia (Repaske and Wilson, 1952) or nitrate
(Mozen et al., 1955) is present, indicating that the site of the block is prev-
ious to these substances. Likewise, in Clostridium H^ inhibits uptake of
N2 but not of ammonia (Hiai et al., 1957). These inhibiting gases are oc-
casionally utilized. For example, NgO is assimilated by Azotohacter, although
slowly, and this is inhibited by N, and Hg (Burris, 1956). It is quite likely
that the competition in all these cases, with the possible exception of Og,
is at the nitrogenase active site binding Ng. Nitrogenase is a metalloflavo-
protein containing molybdenum and it is reasonable that these gases are
bound to the metal, the catalysis of Ng reduction being of similar type to
those mediated by various inorganic metal preparations (which are also
inhibited by other gases).
It will be necessary before discussing mechanisms of inhibition in greater
detail to consider the enzyme hydrogenase, which has recently been closely
linked to nitrogen fixation, and its inhibitions. This enzyme catalyzes the
reduction of some unknown primary acceptor by molecular Ho and the
reduced acceptor then transfers the hydrogen atoms to other acceptors,
such as dyes, NAD, or eventually oxygen. It has been postulated that it
may in some instances participate in the reduction of Ng. The inhibition
by O2 is primarily due to oxygenation of the enzyme:
E + nO, ^ E(02)„
and this inhibition is reversible upon removal of the O2 by dialysis (Krasna
and Rittenberg, 1954; Fisher et al., 1954). Prolonged exposures to Og
lead to progressive inactivation of bacterial hydrogenase (Shug et al.,
1956). Although n has generally been assumed to be 1, Atkinson (1956)
obtained rather complex data possibly indicating a value of 2 for the
Hydrogenomonas facilis enzyme. The hydrogenase-catalyzed evolution of
H2 in Rhodospirillum rubrum (Lindstrom et al, 1949) and soybean root
nodules (Hoch et al, 1960) is inhibited by Ng. Although this might be at-
tributed in part to a diversion of the flow of hydrogen atoms to the reduction
of N2, Bregoff and Kamen (1952) observed that 1 mole of N2 prevents
the release of several moles of Hg. One of the difficulties in assuming a direct
competition between Hg and Ng for the hydrogenase active site is the
fact that, despite the inhibition of Hg evolution by N2, the exchange reac-
tion whereby HD is formed from D2 and a hydrogen donor is actually accel-
erated by N2 (Hoch et al., 1960). The Hj inhibition of nitrogen fixation
was previously claimed to be competitive, but Parker and Dilworth (1963)
found that Hg causes a lag in the N2 uptake at low pN2, whereas in cells
of Azotohacter vinelandii adapted to Hg the lag is abolished. Taking the
lag phase into account, the inhibition is not competitive; from the reciprocal
294 2. ANALOGS OF ENZYME REACTION COMPONENTS
plot presented, although the point scatter is marked, the inhibition might
be uncompetitive. The inhibition of nitrogen fixation in soybean root nod-
ules by O2 is complex, due to both plant and bacterial components of the
respiration, but is competitive when the O2 is above 80% (Bergersen,
1962). NgO is a somewhat more potent inhibitor of H2 evolution than is
Ng, but the most potent is NO, complete and irreversible inhibition being
produced by concentrations of 1% or greater (Shug et al., 1956). The hy-
drogenase in cell-free extracts of Proteus vulgaris is inhibited 87% by
0.002% or 0.00004 m.M NO, and the inhibition at these low concentrations
is partially reversible (Krasna and Rittenberg, 1954). The NO is neither
oxidized nor reduced by the enzyme.
Although the configurations, electronic structures, and physical prop-
erties of these simple gases must be important in determining the inter-
action with nitrogenase and hydrogenase, it is difficult to establish correla-
tions. Some structural and physical properties that might relate to the
interactions of these molecules are given in Table 2-5. Comparing the effects
of H2, N2O, ethane, and the rare gases on Azotobacter nitrogen fixation,
Molnar et al. (1948) concluded there is no correlation with the van der
Waals constants and doubted if any mechanism could be based on physical
properties alone. Wilson and Roberts (1954) postulated that N2O is inhi-
bitory because the N — N distance is close to that in Ng', since NgO is linear,
the oxygen might neither interfere nor be involved in the binding. If the
binding is to metal groups on the enzymes, the degree of interaction would
depend more on the types of bond possible and hence on the electronic
structures of both the gases and the metal, as it is in the interactions of
O2, CO, and NO with hemoglobin and cytochrome oxidase.
The inhibition of nitrogen fixation by O2 has been explained as a compe-
tition between N2 and 0, as terminal acceptors for electrons originating
in the oxidation of substrates by various dehydrogenases, nitrogen fixation
SH, »- XH,
being considered as a form of respiration (Parker, 1954; Parker and Scutt,
1958, 1960). It is likely that the interrelationships between N, and Hg
metabolism, and the inhibitions on these systems, must be considered in
the light of a hydrogen or electron pool with all the possible pathways for
formation and utilization of hydrogen atoms. The scheme below, modified
from Gest et al. (1956), may serve as a means of visualizing some of these
pathways. Some of the inhibitions observed are due to competition for
INHIBITION OF NITROGEN FIXATION BY OTHER GASES
295
^
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+ ^ +;?;
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fi S- -^ O)
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296
2. ANALOGS OF ENZYME REACTION COMPONENTS
Photolysis of water
SH,
(nitrogenase)
N2 (forming NH3)
available hydrogen atoms, while others are based on direct competition
at the enzyme active sites.
PHENOL OXIDASES
These enzymes usually hydroxylate monophenols in the oriho position
and further oxidize these to o-quinones. They have often been named ac-
cording to the particular substrate chosen: e.g., catechol oxidase (cate-
cholase), cresolase, tyrosinase, phenolase, o-diphenol oxidase, or polyphenol
oxidase. Different specificities are observed with enzymes from various
sources. The role such enzymes play in tyrosine metabolism will be dis-
cussed in the following section.
The competitive inhibition of potato catechol oxidase by resorcinol was
first observed by Richter (1934). He noticed that the enzymes from various
sources exhibited quite different susceptibilities to resorcinol, those from
elder (Sambucus) and lilac {Syringa) being more sensitive than the potato
enzyme, and those from mushroom (Polyporus) and mealworm {Tenebrio)
less sensitive. The respiration of apple skin is inhibited about 35% by
50 mM resorcinol and thus Hackney (1948) studied an extracted catechol
OH OH OH
"OH H3C ^-^ OH
Resorcinol Orcinol
COOH
C^
CH-CH— COOH
Nicotinic acid
Cinnamic acid
PHENOL OXIDASES 297
oxidase. Although the inhibition of this enzyme is reduced by increasing
catechol concentration, her data do not correspond to pure competitive
inhibition and the derived K^ seems to be in error, as pointed out by War-
ner (1951). A potato enzyme oxidizing p-cresol is inhibited by resorcinol,
phloroglucinol, and orcinol (see tabulation), the inhibition being completely
Inhibitor (10 mM) % Inhibition (p-cresol = 10 mif)
Resorcinol 78
Phloroglucinol 43
Orcinol 20
reversible by dialysis and apparently competitive (Schneider and Schmidt,
1959). 4-Chlororesorcinol is a much more potent inhibitor of the potato
enzyme, oxidizing either catechol or p-cresol, and it has been claimed that
the initial inhibition is competitive, although progressive inactivation oc-
curs (Heymann et al., 1954). A K^ of 0.024 tqM was calculated. However,
the double reciprocal plots seem to me to be of the perfectly noncompetitive
type. Bonner and Wildman (1946) postulated that the bulk of spinach leaf
respiration passes through a polyphenol oxidase. p-Nitrophenol is a rather
potent inhibitor of this respiration, 1 voM inhibiting 94%, and of the po-
lyphenol oxidase, whether it is oxidizing catechol or p-cresol. On the other
hand, o-nitrophenol is only a weak inhibitor of both. It was felt that p-nitro-
phenol might well be an analog of naturally occurring substrates, and o-
nitrophenol an analog of o-phenols against which the enzyme is inactive.
It is interesting, finally, to note that dihydroxymaleate is an inhibitor of
catechol oxidase (Florkin and Duchateau-Bosson, 1939), and it was sug-
gested that the
OH OH
— C = C—
grouping complexes with the enzyme in a manner similar to the analogous
catechol grouping. Most of this work on inhibiting phenolic compounds
is unsatisfactory from the quantitative standpoint and clear-cut proofs
of competitive inhibition are lacking. Nevertheless, effective analog in-
hibition for this class of enzymes has heen demonstrated.
We shall now turn to a more potent, more interesting, and more
thoroughly studied type of inhibitor, namely, the benzoates. The inhibition
of mushroom catechol oxidase by benzoate itself was attributed to a com-
petition with the substrate for the active center, although no direct evidence
for this was adduced (Ludwig and Nelson, 1939; Gregg and Nelson, 1940),
but good competitive kinetics for the inhibition by m-hydroxybenzoate
298
2. ANALOGS OF ENZYME REACTION COMPONENTS
were observed later (Warner, 1951), with K^ 2.5 mM for the potato en-
zyme and 0.6 mM for the mushroom enzyme. An excellent investigation
by Kuttner and Wagreich (1953) of the inhibition of a catechol oxidase
from Psalliota campestris provides data from which some ideas of the me-
chanism may be obtained. Some of the inhibition data are presented in
Table 2-6, and have been used to calculate the apparent relative binding
energies. However, these inhibitors are mostly weak acids and the degree
Table 2-6
Inhibition of a Phenol Oxidase from Psalliota campestris with Catechol ( 1 .82 mM)
AS Substrate (at pH 5.2 and 25°)
Inhibitor
Concentration
(mM)
% Inhibition
Relative
— zli^ of binding"
(kcal/mole)
Benzoate
0.012
50
6.96
p-Chlorobenzoate
0.023
50
6.56
p-Methylbenzoate
0.04
50
6.23
o-Chlorobenzoate
0.21
50
5.21
p-Methoxybenzoate
0.26
51
5.10
Nicotinate
0.34
48
4.88
o-Hydroxybenzoate
0.38
50
4.84
p-Nitrophenol
0.48
50
4.70
p-Hydroxybenzoate
0.49
50
4.69
irans-Cinnamate
0.81
62
4.43
Phenylacetate
0.77
50
4.40
o-Methylbenzoate
1.0
50
4.25
o-Nitrophenol
1.1
50
4.19
Benzoate methyl ester
0.65
30
3.94
Hydroquinone
2.2
50
3.76
p-Nitrobenzoate
0.72
19
3.55
Orcinol
3.2
50
3.53
Resorcinol
4.5
50
3.32
o-Nitrobenzoate
0.72
8
2.95
o-Methoxybenzoate
8.0
49
2.94
" The relative energies of binding to the enzyme were calculated assuming competi-
tive inhibition and without taking into account the state of ionization. (Data from
Kuttner and Wagreich, 1953.)
PHENOL OXIDASES
299
of ionization at the experimental pH of 5.2 varies. It was found that the
inhibition by benzoate decreases with rise in the pH (see tabulation) and
pH
% Inhibition by benzoate (0.0123 mil/)
% Un-ionized
5.2
5.8
6.4
7.0
56
28
9.0
2.5
0.6
0.2
similar results were obtained with some substituted benzoates. This was
interpreted to mean that the un-ionized form of the inhibitor is the active
one, for example that benzoic acid is the inhibitor and not the benzoate
ion. The relative binding energies have been recalculated (Table 2-7) on
this basis and a somewhat different order of potency is obtained. This il-
Table 2-7
Inhibition of Mushroom Phenol Oxidase Corrected for the State of Ionization
OF THE Inhibitors "
Corrected relative
Inhibitor
P^a
(HA)/(A,)
— AF of binding
(kcal/mole)
Benzoate
4.203
0.0914
8.44
o-Chlorobenzoate
2.943
0.0055
8.39
p-Chlorobenzoate
3.978
0.0565
8.32
o-Hydroxybenzoate
3.001
0.0063
7.95
p-Methylbenzoate
4.371
0.129
7.47
o-Nitrobenzoate
2.173
0.00094
7.24
p-Methoxybenzoate
4.470
0.157
6.24
o-Methylbenzoate
3.909
0.0487
6.10
p-Nitrobenzoate
3.425
0.0165
6.07
p-Hydroxybenzoate
4.559
0.186
5.71
Nicotinate
4.854
0.311
5.59
o-Methoxybenzoate
4.092
0.0724
4.56
° Relative binding energy corrected on the basis that the un-ionized forms of the
inhibitors are active. The ionization constants were obtained from Dippy (1939);
inhibition data are given in Table 2-6 (Kuttner and Wagreich, 1953.)
300
2. ANALOGS OF ENZYME REACTION COMPONENTS
lustrates the importance of taking ionization into account in the comparison
of inhibitors, as discussed earlier in the chapter. It must be emphasized
that the absolute values of the binding energy are meaningless; it is only
the differences between the — JF values that are significant.
Before discussing the implications of these results we will examine the
data reported by Krueger (1955) on the inhibition of a mushroom enzyme
oxidizing p-cresol (Table 2-8). Confirming the work of Kuttner and Wag-
Table 2-8
Inhibition of p-Cresol Oxidation by Mushroom Tyrosinase '
Inhibitor
Concentration
(mif)
% Inhibition
Relative
— AF oi binding
pH 5.3
pH7.0
(kcal/mole)
Benzoate
4*
90
27
5.20
Oxalate
4*
83
—
4.78
Cyclohexanecarboxylate
16
84
—
3.57
Phenylacetate
4
54
0
3.51
Fluoride
40*
72
14
2.99
Butyrate
40
73
10
2.60
Bromide
40*
56
5
2.56
Iodide
40*
54
8
2.51
Benzamide
4*
8
—
2.33
Lactate
20
40
0
2.16
Terephthalate
4
11
—
2.12
Phthalate
4
10
—
2.05
Chloride
40
45
5
1.85
Formate
4
6
—
1.71
20
24
—
40*
10
0
Acetate
20
21
—
1.68
40
42
6
Trimethylacetate
40
30
4
1.46
Chloroacetate
20
16
0
1.39
Benzenesulfonate
40
20
0
1.13
Trichloroacetate
20
8
—
0.91
" Concentration of p-cresol was 4.63 mM except where indicated by asterisks, in
which cases it was 9.26 n\M. The relative binding energies were calculated for pH 5.3
and were not corrected for ionization. (Data from Krueger, 1955.)
PHENOL OXIDASES 301
reich, a marked decrease in the inhibition with an elevation of the pH
from 5.3 to 7 is observed, but Krueger interpreted this as indicating an
ionizing group on the enzyme with a pK^ around 6 and possibly an imida-
zole group. One reason for assuming the ionizing group to be on the enzyme
is the pH effect on inhibitions by the inorganic anions; however, the in-
hibition by these ions may be through a different mechanism than the
benzoates, and indeed Krueger found noncompetitive kinetics for chloride.
The substrates for the enzyme are, of course, un-ionized and this might
favor the concept of the acid form of the inhibitors being active and the
important ionizing group on the inhibitors. It is impossible at the present
time to decide which is the correct interpretation and hence the signifi-
cance of the pH effects. Ionizing groups on both enzyme and inhibitors
might also be considered. It should be added that the following were found
to be without effect: sulfate (40 mM), nitrate (40 mM), pyrophosphate (40
mM), pyruvate (20 mM), succinate (20 mM), maleate (20 mM), fumarate
(20 mM), and ethyl benzoate (4 mM).
The lack of information on the exact catalytic mechanism involved in
these enzymes, and particularly our ignorance of the state and role of
copper, make it difficult to understand the binding of the inhibitors. There
are two copper ions at the active center but we do not know if they com-
plex with oxygen, or the substrates, or both, or whether one copper com-
plexes with oxygen and the other with the substrates. Some of the ionic
inhibitions might be due to the formation of complexes with the copper;
such complexes might be more difficult to form at higher pH's because of
competition with hydroxyl ions. The strong inhibition by oxalate might
be due to chelation of the copper, but it is odd that pyrophosphate does
not inhibit since it also chelates copper weU. It is also surprising that sul-
fate and nitrate do not inhibit at all, unless the ionic size is as critical as
Krueger believes.
It would appear that inhibitory activity is related to the presence of
a carboxyl group (excepting the inorganic ions). Benzoate ester and benza-
mide are bound much less tightly than benzoate (around 3 kcal/mole
difference). The weak action of benzenesuLfonate might be explained in
three ways; (1) if the un-ionized form of the acid is necessary for activity,
there would be less in the case of benzenesulfonic acid than with benzoic
acid, (2) the sulfonate group might be too bulky, according to Krueger
(although it is certainly not much larger than the carboxyl group), and
(3) the sulfonate group does not have the ability to form bonds with the
copper or hydrogen bonds with an enzyme group. Separation of the car-
boxyl from the benzene ring reduces the binding, as in phenylacetate or
cinnamate. The second requirement for potent inhibitory activity is a
benzene ring. This may be seen by comparing acetate and phenylacetate,
benzoate and nicotinate, and benzoate and cyclohexanecarboxylate; in
302 2. ANALOGS OF ENZYME REACTION COMPONENTS
the last example the increased binding of benzoate may be due to its great-
er polarizability. In general the substitution of groups on the benzene
ring reduces the affinity for the enzyme. This may be due to steric inter-
ference with the approach of the ring or to inductive effects on the carboxyl
group's interaction with the enzyme. It is rather odd that an oriho chlorine
does not disturb binding much while an ortho methoxy group reduces the
binding some 4 kcal/mole.
The forces binding the substrates and inhibitors to these enzymes are
thus vague at the present time. It is possible that hydrogen bonds between
the OH or COOH groups and the enzyme are important, and it is equally
possible that bonds to the copper ions are involved. One might conceive
of the inhibitor's COOH group reacting with either the two copper ions,
or with a copper ion and a vicinal — NH — group. Copper ions are able to
catalyze the oxidation and hydroxylation of phenols nonenzymically, and
it might be interesting to study the inhibition of such reactions by some
of the compounds active in the enzymic reaction.
TYROSINE METABOLISM
Many interesting and practically important examples of analog inhibi-
tion are to be found in the general field of amino acid metabolism, and we
shall begin the discussion of this subject by considering the inhibitions of
the various pathways of tyrosine metabolism. Tyrosine may be hydroxyl-
ated to form dihydroxyphenylalanine (dopa), oxidatively deaminated or
transaminated to form p-hydroxyphenylpyruvate, decarboxylated to form
tyramine, or activated prior to incorporation into proteins; inhibition
of all of these reactions by analogs has been reported. The scheme on page
303 indicates the major pathways of tyrosine metabolism. Interference
with these reactions might be expected to bring about physiological changes
due to the acceleration or suppression of active amine synthesis, and also
to affect melanin formation.
Tyrosinase (Phenol Oxidase)
These enzymes hydroxylate tyrosine in the oriho position to form dopa
and further oxidize dopa to dopa-quinone. The enzymes discussed in
the previous section generally possess this activity. However, mammalian
tyrosinases are much more specific than the plant enzymes and oxidize
tyrosine and dopa more rapidly than other phenols. Inhibitions of the en-
zymes with tyrosine as the substrate will be considered in this section,
but the results with p-cresol or catechol as substrate are probably generally
applicable to tyrosinase activity. The high inhibitory potency of 4-chloro-
resorcinol on potato tyrosinase, as determined by the rate of formation of
TYROSINE METABOLISM
303
n ^ -2 — ■
2 rt tn D M G
•5 ^ O -rl O 5
oS
2 >> e
a
304 2. ANALOGS OF ENZYME REACTION COMPONENTS
melanin from tyrosine, was noted by KuU et al. (1954), for example (see
following tabulation). Other substitutions in the 4-position usually reduce
Lowest
Inhibitor
inhibitory concentration
(mil/)
4-Chlororesorcinol
0.000069
Resorcinol monobenzoate
0.0047
m-Aminophenol
0.0092
4-«.-Hexylresorcinol
0.103
Naphthoresorcinol
0.125
Orcinol
0.141
j)-Aminophenol
0.183
Resorcinol
0.183
Phloroglucino!
6.17
2-Nitroresorcinol
65
or abolish the inhibitory activity. The formation of melanin involves several
steps and the inhibitions observed are not necessarily entirely on tyrosinase.
Hydroquinone was found to be a weak inhibitor but the monobenzyl
ether of hydroquinone is as potent as resorcinol. This latter substance,
"°A\ /r°-'''^-A\ //
Monobenzone (Benoquin)
known also as monobenzone (Benoquin), is an inhibitor of melanin for-
mation in the skin when applied topically, can produce leucoderma in
Negroes, and is used in various conditions of melanosis. Some have thought
that it releases hydroquinone after penetration into the skin but this is
questionable in view of its own inhibitory activity.
An active tyrosinase occurs in the Hardin-Passey mouse melanoma and
is probably responsible for pigment formation. It is competitively inhib-
ited by various tyrosine analogs (Lerner et al., 1951). The values of K^
shown in the tabulation were calculated on the basis of a K„, of 0.60 mM
Inhibitor J^, (mM)
iV-Acetyl-L-tyrosine 0.140
iV-Formyl-L-tyrosine 0.177
3-Amino-L-tyrosine 0.314
3-Fluoro-L-tyrosine 1.25
TYROSINE METABOLISM 305
obtained from the reciprocal plots. 3-Nitro-L-tyrosine and O-methyl-L-
tyrosine are not inhibitory. The effects of 3-substitution may be mediated
through inductive effects on the 4-OH group and its interaction with the
enzyme, whereas iV-substitution must lead to an altered position of binding
to prevent oxidation. L-Phenylalanine and phenylpjTuvate inhibit com-
petitively the tyrosinase from melanoma, inhibit the incorporation of ty-
rosine-C^* into melanin, and depress the respiration of tumor tissue with
tyrosine as the substrate (Boylen and Quastel, 1962). The high concentra-
tions of these inhibitors in phenylketonuria might be responsible for the
reduced pigment formation in these individuals.
Tyrosine : a-Ketoglutarate Transaminase
The effects of numerous analogs on the formation of p-hydroxyphenyl-
pyruvate from tyrosine and a-ketoglutarate by a dog liver transaminase
were reported by Canellakis and Cohen (1956 b); some of the results are
given in Table 2-9. Certain of these analogs are transaminated (e.g., the
3-substituted tyrosines) and the inhibitory activity varies inversely with
their abilities to act as substrates. The rapid transamination of 3-fluoro-
tyrosine may partly explain its toxic effects and inhibition of growth,
since fluorofumarate or fluoroacetoacetate may be formed. Comparing the
hydroxyl-substituted phenylalanines, it is seen that a m- or p-hydroxyl
is necessary for tight binding, the contribution to the binding energy
being over 2 kcal/mole. The if,,; for L-tyrosine is 0.71 mM, so that its rel-
ative binding energy is at least — 4.47 kcal/mole, and that of m-hydroxy-
DL-phenylalanine is at least — 4.75 kcal/mole (— 5.18 kcal/mole if only
the L-isomer is active), which may be compared with the — 2.55 kcal/
mole for L-phenylalanine. Hydroxyl groups in the o-positions, on the other
hand, do not augment binding very much. The tighter the binding between
a basic hydroxyl and an acidic enzyme group, the greater the inhibitory
activity; ring substituents modify the electronic character or basicity of
the phenolic group. A carboxylate group is necessary for strong binding,
as may be seen by comparing tjTosine with tyramine, and p-hydroxyben-
zoate with p-cresol. The a-amino group may also be involved in the binding
(— AF for p-hydroxyphenylacetate is 2.87 kcal/mole), but lacking data on
p-hydroxyphenylproprionate it is not possible to evaluate this accurately.
The strong inhibition by epinephrine is rather surprising in the light of
the absence of a carboxylate group, but the /?-hydroxyl or iV-methyl group
may contribute to make up for this deficiency. A similar study on the rat
liver enzyme has been reported by Jacoby and La Du (1964).
p-Hydroxyphenylpyruvate Oxidase
The further oxidation of the product of tyrosine transamination is ca-
talyzed by an enzyme from dog liver and is inhibited markedly by phenyl-
306
2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-9
Inhibition of Tyrosine : a-KETOGLUTARATE Transaminase from Dog Liver "
Relative
Inhibitor
(I)/(S)
% Inhibition
— zJi^ of binding
(kcal/mole)
3-Araino-DL-tyTosine
1
85
6.00
3,4-Dihydroxy-L-phenylalanine
2
70
5.03
Epinephrine
2
70
5.03
m-Hydroxy-DL-phenylalanine
2
60
4.75
2,5-Dihydroxy-L-phenylalanine
2
35
4.13
3-Fluoro-DL-tyrosine
2
20
3.65
3,5-Dibromo-L-tyrosine
2
10
3.15
o-Hydroxy-DL-phenylalanine
2
7
2.91
p-Hydroxybenzoate
10
26
2.87
p-Hydroxyphenylacetate
10
26
2.87
p-Aminobenzoate
10
21
2.70
L-Tryptophan
10
21
2.70
p-Nitro-DL-phenylalanine
2
5
2.69
L-Phenylalanine
2
4
2.55
o-Hydroxybenzoate (saHcylate)
10
17
2.53
p-Nitrobenzoate
10
15
2.44
3,5-Diiodo-L-tyrosine
2
3
2.36
D-Tyrosine
2
3
2.36
j)-Cresol
10
13
2.34
Tjrramine
10
9
2.09
" Relative binding energies were calculated on the basis of competitive inhibition
with K^ = 0.71 mM for L-tyrosine. No correction for ionization was made. It should
be noted that the different isomers of the DL-compounds may have different activities,
so that the binding energy of the active form should be increased. (Data from Canel-
lakis and Cohen, 1956 b.)
pyruvate, essentially complete inhibition occurring with 0.4 mM in the
presence of 2 mM substrate (Zannoni and La Du, 1959). The inhibition is
negligible for the first 5 min but then increases to become complete at
around 20 min. This might be due to protection by the substrate and pre-
incubation experiments would have been informative. An equally good
inhibitor is m-hydroxyphenylpyruvate but phenylacetate, 2,5-dihydrox-
yphenylpyruvate, p-hydroxyphenyllactate, p-hydroxybenzoate, and ho-
mogentisate are not inhibitory.
Other Pathways of Tyrosine Metabolism
Little is known about the effects of analogs on tyrosine decarboxyla-
tion, although this might well be an important site to block if one wished
TYROSINE METABOLISM
307
to reduce the tissue tyramine concentration. Mardashev and Semina
(1961) found that the tyrosine decarboxylase from Streptococcus fecalis is
inhibited 20% by 8.3 mM cysteine and 25% by 8.3 mM homocysteine
when substrate concentration is 2.8 mM. This is a general phenomenon seen
with several amino acid decarboxylases and is presumably due to the for-
mation of complexes of the inhibiting amino acids with pyridoxal phosphate.
The tyrosine-activating enzyme of pig pancreas catalyzes the first
Amino acid + ATP + E -> E-amino acyl-AMP + PP
step in the incorporation of amino acids into proteins. This also would
be a very interesting step to investigate from the standpoint of analog
inhibition, but our present information is meager. Schweet and Allen
(1958) found that 3-fluoro-lL-tyrosine activated 50% the rate of L-ty-
rosine and does not inhibit. L-tyrosinamide inhibits weakly but tyramine
inhibits the phosphate exchange reaction 80% at 0.2 mM, this inhibition
being reduced by increase in substrate concentration. The importance of
the p-hydroxyl group is indicated by the fact that no inhibition is seen
with phenylethylamine.
Dihydroxyphenylalanine (Dopa) Decarboxylase
This enzyme is on the pathway leading from tyrosine to the important
catecholamines, epinephrine and norepinephrine, and lately has been the
subject of much investigation because of the possible clinical applications
of producing a selective block at this step. Inhibitors have indeed been
found which are effective in vivo, reduce amine formation, and lower the
blood pressure. The sequence of reactions for the formation of amines
from dopa is the following:
HO
HO
NH3
-CH— COO'
dopa
decarboxylase
HO
CH^ CH,— NH,
Dopa
Dopamine
dopamine /3-oxidase
OH
1 H
CH— CH,— NH,
Epinephrine
308
2. ANALOGS OF ENZYME REACTION COMPONENTS
A block of dopa decarboxylase would thus decrease the rate of formation
of these three physiologically active amines in the tissues.
Of historical interest are the following observations on the analog inhi-
bition of this enzyme: dopamine (Blaschko, 1942), epinephrine (Schapira,
1946), various aromatic amines (tryptamine, tyramine, phenylethylamine,
etc.) (Polonovski et al., 1946), and the various hydroxy, methoxy, and
dimethoxy derivatives of phenylethylamine, the dimethoxy analogs being
the most potent (Gonnard, 1950).
These studies were extended in important ways by Sourkes (1954),
who discovered the potent inhibiting activity of certain a-methylphenyl-
alanines (Table 2-10). Especially inhibitory is a-methyldopa and this sub-
Table 2-10
Inhibition of Dopa Decarboxylase from Pig Kidney Cortex "
Inhibitor
Concentration
(mM)
% Inhibition
Relative
— /iF oi binding
(kcal/mole)
a-Methyldopa
0.01
22
5.94
0.1
71
0.5
98
a-Methyl-3-hydroxy-PA
0.05
45
5.57
0.5
74
5
95
a-Methyl-3-hydroxy-4-
4.3
92
4.86
methoxy-PA
a-Methyl-3,4-dimethoxy-PA
0.01
16
4.65
0.1
20
iV-Acetyl-3,4-dimethoxy-PA
3.6
44
3.32
A'^-Methyldopa
0.8
11
3.12
1.7
30
Diiodotyrosine
2.3
15
2.68
3,4-Dimethoxy-PA
5
25
2.59
2,4-Dimethoxy-PA
5
22
2.49
3-Methoxy-4-hydroxy-PA
5
18
2.33
a-Methyl-3-methoxy-PA
5
18
2.33
a-Methyltyrosine
5
16
2.25
a-Methyl-PA
2
6
2.13
A'^-Methyl-3-methoxy-4-
1.6
0
—
hydroxy-PA
" Concentration of DL-dopa 4 mM, pH 6.8, preincubation with inhibitor 15 min.
PA — phenylalanine, and dopa = 3,4-dihydroxyphenylalanine. Relative binding ener-
gies calculated on the basis of competitive inhibition, which may not be strictly true;
in any event, these values give a better indication of the relative inhibitory potency
than the per cent inhibition at different concentrations. (Data from Sourkes, 1954.)
TYKOSINE METABOLISM 309
stance has been thoroughly studied biochemically and pharmacologically
during recent years. It is interesting that these analogs have very little
inhibitory activity toward tyrosine decarboxylase and that a-methyl-
tyrosine does not inhibit dopa decarboxylase strongly, both facts pointing
to the importance of the 3-hydroxyl group in the binding to the enzyme.
This is also seen by comparing or-methylphenylalanine and its hydroxylated
derivatives: The addition of a 4-hydroxyl has little effect, whereas a
3-hydroxyl increases the binding energy over 3 kcal/mole. A 3-methoxy
group seems to be ineffective.
The inhibition by a-methyldopa was shown to be pseudoirreversible
by varying the enzyme concentration and using the graphic procedure of
Ackermann and Potter (1949). At concentrations of 0.01-0.03 mM, the
inhibition being 15-25%, the behavior is fairly reversible, but at concen-
trations of 0.1 mM or above there is marked nonlinearity of the curves.
As pointed out by Sourkes, these data indicate merely that K^ is low and
the affinity for the enzyme is high. The binding might be to the apoenzyme,
to a great extent through the phenolic groups, or the inhibition could be
the result of reaction with pyridoxal phosphate. The former mechanism
was favored by Sourkes on the basis of the following evidence against a
reaction with the coenzyme. (1) The inhibition is reversible by dialysis.
(2) The rate of nonenzymic reaction of a-methyldopa with pyridoxal
phosphate is too slow at inhibiting concentrations to be significant. (3) In-
crease in pyridoxal phosphate concentration does not alter the inhibition
significantly. (4) Analysis for pyridoxal phosphate at the end of inhibition
experiments showed no loss. (5) Tyrosine decarboxylase is also a pyridoxal
phosphate enzyme and is not inhibited. None of this evidence is completely
conclusive and it is possible that a-methyldopa can form a reversible
complex with pyridoxal phosphate on the enzyme surface, so that increase
in coenzyme concentration would not be effective and analysis for total
coenzyme would not detect the small amount combined. 5-Hydroxytryp-
tophan decarboxylase is also potently inhibited by a-methyldopa (it is
possible that the decarboxylases for dopa, 5-hydroxytryptophan, trypto-
phan, tyrosine, and phenylalanine in mammalian tissues represent a single
enzyme) and S. E. Smith (1960 a) has investigated the mechanism, using
the mouse brain enzyme. Plots of 1/(S) against l/v showed pure competitive
inhibition with respect to substrate at higher coenzyme concentrations
(above 0.01 mM), but at low coenzjTne concentrations the inhibition be-
comes noncompetitive with substrate. In contrast to dopa decarboxylase,
increase of coenzyme concentration leads to a reduction in the inhibition
(Fig. 2-3). Smith inclines to a coenzyme inactivation mechanism but admits
that the inhibition is incompletely explained. If a-methyldopa forms a
complex with pyridoxal phosphate on the enzyme surface, which it can do
because it is decarboxylated slowly, it might be considered to be an in-
310
2. ANALOGS OF ENZYME REACTION COMPONENTS
hibitor which enters into" the catalytic sequence of reactions but is not
able to complete the process readily. This mechanism is also suggested by
the results of Lovenberg et al. (1963) on kidney aromatic amino acid de-
carboxylase, using tryptophan as the substrate. When a-methyldopa is
preincubated with the enzyme in the absence of pyridoxal-P the inhibition
is noncompetitive and potent, but when pyridoxal-P is added during the
preincubation period the inhibition is competitive with respect to substrate
0.001
■ METHYL - OOPA
Fig. 2-3. Inhibition of mouse brain 5-hydroxytryptophan
decarboxylase by a-methyldopa. The sohd curves show
the rate of formation of serotonin in //g/g/hr, and the
dashed curves show the fractional inhibition. The curves
X-X-X-X show the results without addition of pyridoxal-P,
and the curves 0-0-0-0 show the results after addition
of 0.008 mM pyridoxal-P. (From S. E. Smith, 1960 a.)
and is less potent. If no preincubation is done and a-methyl dopa is added
with the substrate, competitive inhibition is observed. The data suggest
that a-methyldopa interacts specifically with the enzyme-pyridoxal-P com-
plex (the enzyme as isolated contains tightly bound pyridoxal-P), and the
protection or reversal of the inhibition by exogenous pyridoxal-P may be
due to the reactivation of the enzyme-bound coenzyme. The reaction of
the a-methyldopa with pyridoxal-P may involve the cyclization of a
Schiff base (Mackay and Shepherd, 1962).
Another comprehensive study of dopa decarboxylase was made by Hart-
man et al. (1955), who determined the inhibitory activities of some 200
compounds. The results, some of which are presented in Table 2-11, enable
TYROSINE METABOLISM
311
Table 2-11
Inhibition of Dopa Decarboxylase from Pig Kidney Cortex"
Inhibitor
Concen-
tration
(mM)
%
Inhibition
Relative
- AF of
binding
(kcal/mole)
Cinnamates
O
CH=CH— COO'
3-Mercapto-
0.1
78
6.90
3,4-Dihydroxy- (ethyl ester)
0.3
90
6.78
2, 6-Dihydroxy-
0.2
84
6.70
3, 4-Dihydroxy-
0.4
74
5.90
2-Hydroxy-3, 5-dibromo-
0.4
65
5.63
3-Hydroxy- (methyl ester)
0.4
60
5.50
2-Hydroxy- (methyl ketone)
0.2
31
5.19
3-Hydroxy- (methyl ketone)
0.2
30
5.15
3-Hydroxy-
0.4
37
4.92
a-Methyl-3-hydroxy-
0.4
31
4.75
3-Hydroxy-6-sulfonate-
0.4
31
4.75
a-Ethyl-3-hydroxy-
0.4
27
4.64
3 -Hydroxy- (amide)
0.4
19
4.35
2-Hydroxy-
2
52
4.30
2, 4-Dichloro-
2
41
4.03
2-Chloro-
2
36
3.91
4-Hydroxy-
0.4
7
3.65
3-Nitro-
2
15
3.19
3-Amino-
2
10
2.91
2, 4-Dihydroxy-
2
9
2.83
2, 3-Dimethoxy-
2
8
2.75
4-Chloro-
2
8
2.75
Unsubstituted
2
0
~
Hydrocinnamates <( \— CH2CH2— COO"
V /
3-Hydroxy-2, 4, 6-triiodo-
0.2
90
7.04
a-Methyl-3-hydroxy-2, 4, 6-triiodo-
0.4
91
6.68
3, 4-Dihydroxy-
0.44
63
5.52
a-Methyldopa
0.2
37
5.35
4-Hydroxy-
2
9
2.83
3-Hydroxy-
2
0
-
312 2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-11 (continued)
Inhibitor
Concen-
tration
(mA'i)
%
Inhibition
Relative
- AF of
binding
(kcal/mole)
d-Hydroxy-
Unsubstituted
Phenylacetates
3, 4-Dihydroxy-
2, 5-Dihydroxy-
3, 4-Dimethoxy-
3 -Hydroxy -
Unsubstituted
CH,— COO"
2
88
5.49
2
28
3.68
2
18
3.32
2
11
2.97
2
0
_
Phenylglycines
3, 4-Dihydroxy-
3-Hydroxy-
4-Hydroxy-
Unsubstituted
=\ NH,
CH
\
COO
2
16
2
0
2
0
2
0
3.24
Phenylpyruvates
3, 4-Dihydroxy-
3-Hydroxy-
4-Hydroxy-
2, 5-Dihydroxy-
Unsubstituted
CH5— CO— COO'
0.2
1
2
2
2
60
72
60
60
60
5.92
5.26
4.50
4.50
4.50
Benzoates (\ n — - COO"
2-Hydroxy-3, 5-diiodo-
2-Hydroxy-3, 5-dibromo-
3, 5-Dibromo-
3,4, 5-Trihydroxy-
3-Hydroxy-4, 6-dibromo-
2 -Hydroxy- 5-amino-
2-Hydroxy-4-nitro-
0.2
47
5.60
0.2
33
5.24
0.2
10
4.33
2
42
4.07
2
36
3.91
2
28
3.68
2
26
3.62
TYROSINE METABOLISM
313
Table 2-11 (continued)
Inhibitor
Concen-
tration
(niiW)
%
Inhibition
Relative
- AF of
binding
(kcal mole)
2, 5-Dihydroxy-
2
21
3.44
2, 4, 6-Trihydroxy-
2
16
3.24
3, 4-Dihydroxy-
2
0
-
2. 4-Dihydroxy-
2
0
-
Miscellaneous
5- (3. 4-Dihydroxycinnamoyl)salicylate
0.002
87
9.69
HO
COO"
CH=CH-CO
OH
° Concentration of dopa 2 mM and pH 6.8. (Data from Hartman et al., 1955. )
one to speculate further about the nature of the binding to the active
center. Several inhibitors more potent than a-methyldopa were found.
The basic structure for inhibition was written as:
HO
(HO)
O
I I II
c=c-c— X
where X is OH, 0-alkyl, alkyl, or aryl. It is rather surprising that the
negatively charged carboxylate group is not necessary, esters and amides
being as potent as the acids, and it may be that the CO group is critical.
The positively charged amino group is also not necessary, since 3,4-dihy-
droxyhydrocinnamate is a good inhibitor, and this would make it likely
that the binding of the inhibitors is not too much dependent on pyridoxal
phosphate. The importance of the 3- and 4-hydroxyls is again evident and
all the potent inhibitors have phenolic groups; apparently only the sulf-
hydryl group can replace the hydroxyl. Halogens have the ability to aug-
ment binding when they are the only substituents but particularly when
a hydroxyl group is also present; 3,5-dibromobenzoate and 2,4-dichloro-
cinnamate are bound reasonably well (at least 2 kcal/mole more than the
unsubstituted compounds). The only unsubstituted inhibitor is phenyl-
pyruvate, which must be significant, although of what is not clear. The
interaction of the side chain must be complex and involve different types
of forces. If one compares all the 3,4-dihydroxy derivatives, it is seen that
314 2. ANALOGS OF ENZYME REACTION COMPONENTS
inhibitory activity increases with the length or bulk of the side chain.
Also it may be noted that the Hnear cinnamate derivatives are generally
more potent than the hydrocinnamates. One must conclude that the most
important binding groups are the hydroxyls, the ring, the side-chain car-
bonyl, and any more terminal groups, which just about includes all of the
molecule. The specificity of these inhibitors may well be quite high, since
3-hydroxycinnamate and 3,4-dihydroxycinnamate (caffeate) were tested
on tyrosine decarboxylase, glutamate decarboxylase, histidine decarbo-
xylase, and succinate dehydrogenase, and found to be without effect.
Tyrosinase is inhibited somewhat, especially by caffeate.
There is evidence in patients with phenylpyruvic oligophrenia of a dis-
turbance in tyrosine metabolism (hypopigmentation), and it is possible
that the phenyl acids which are abnormally high might be inhibiting some
step or steps in these pathways. Fellman (1956) therefore studied the ef-
fects of such substances on the dopa decarboxylase from beef adrenal
medulla. The order of inhibitory potency is phenylpyruvate > phenyllac-
tate > phenylacetate > phenylalanine. Phenylpyruvate inhibits 77% when
equimolar (3.3 milf ) with L-dopa. The low plasma epinephrine levels found
in these patients thus might be due to such an inhibition, but another point
of attack would have to be adduced for a suppression of melanin formation.
It is interesting that this enzyme is inhibited quite strongly by norepi-
nephrine and dopamine, whereas epinephrine exerts no effect (Fellman,
1959). The susceptibilities of dopa decarboxylases from various tissues
are obviously different, since the results of Fellman are often different
from those of previous workers.
cf-Methyldopa and related analogs can effectively inhibit decarboxylases
in vivo, thereby interfering with amine formation and modifying tissue
function. Direct evidence for an in vivo inhibition was obtained by intra-
muscular injection of «-methyldopa into guinea pigs and demonstration
of a marked depression of the decarboxylation of both dopa and 5-hydroxy-
tryptophan in isolated kidney 15-30 min afterward (Westermann et at.,
1958). Indeed, the inhibition of 5-hydroxytryptophan decarboxylation is
complete and after 90 min is 83%. More indirect evidence has been obtain-
ed by showing that these analogs prevent the pharmacological actions of
dopa and 5-hydroxytryptophan, these actions being dependent on decar-
boxylation of these substances to dopamine and serotonin, respectively.
Injection of dopa leads to a rise in the blood pressure which is probably
primarily due to dopamine (although some norepinephrine and epinephrine
may also be formed). This pressor response can be blocked by several de-
carboxylase inhibitors, including 5-(3-hydroxycinnamoyl)salicylate (Po-
grund and Clark, 1956) and a-methyldopa (Dengler and Keichel, 1958).
There is no effect on the response to dopamine or norepinephrine. The
increase in cardiac contractility induced by dopa is also completely blocked
TYROSINE METABOLISM 315
by a-methyldopa. 5-Hydroxytryptoplian causes bronchoconstriction in
guinea pigs and central excitation in mice (if brain monoamine oxidase is
blocked), these effects being due to serotonin, and pretreatment of the
animals with a-methyldopa prevents these actions (Westermann et al.,
1958).
Decarboxylase inhibition should lead to a decrease in the tissue concen-
trations of certain amines and this has been demonstrated. The degree
of reduction will depend on the relative rates of formation and metabolism
of the amines, as well as on the magnitude of the decarboxylase inhibition
(which will depend in part on the penetration of the analogs into the tis-
sues), since we are dealing with steady-state multienzyme systems. Injec-
tion of a-methyldopa (200 mg/kg) into dogs leads to a lowering of serotonin
in the caudate nucleus (1.03 to 0.43 //g/g at 3 hr), a more prolonged lowering
of norepinephrine (2.41 to 1.77 /ngjg at 24 hr) (Goldberg et al., 1960), and
a fall of total catecholamines in the brain stem (0.17 ^- 0.08 //g/g), heart
(0.59 -^ 0.26 //g/g), and spleen (0.94 -^ 0.43 //g/g) (Stone et al, 1962). In
the mouse, brain serotonin is reduced but norepinephrine is unaffected
(S. E. Smith, 1960 a). The urinary amines in four hypertensive patients
were decreased by a-methyldopa (1-4 g per day): the reductions were
81% for t>Tamine, 63% for serotonin, and 55% for tryptamine (Gates
et al, 1960).
Altering these amine levels should produce physiological disturbances.
It has been found that a-methyldopa lowers the blood pressure and causes
sedation in the dog (Goldberg et al, 1960), decreases coordinated activity
and produces miosis in mice (S. E. Smith, 1960 a), and in a variety of animals
induces a syndrome similar to that produced by reserpine (including hy-
pothermia), a drug releasing amines from the tissues (S. E. Smith, 1960 b).
a-Methyldopa is being studied clinically for the reduction of hypertension
and a preliminary report (Gates et al., 1960) indicated its effectiveness,
doses of 1-4 g/day for 1 week leading to a fall in supine blood pressure
from 187.0/115.4 to 173.4/108.1 and in standing blood pressure from 177.7/
119.3 to 138.6/98.0, the controls being given a placebo in a double-blind
study.
It is thus clear that a-methyl-dopa can inhibit certain amino acid de-
carboxylases in vivo, can alter amine levels in tissues, and can produce
physiological disturbances that could reasonably be attributed to the in-
hibition. Recently, however, more detailed studies of tissue amines have
indicated that other mechanisms are possibly operative. Injections of
a-methyl-3-hydroxyphenylalanine into guinea pigs lead to a reduction
in brain amines (Fig. 2-4), the degree of lowering and the duration of the
effect depending on the amine. a-Methyldopa acts similarly but is slightly
more potent. Simultaneously there is an inhibition of amino acid decar-
boxylase (Fig. 2-5). Cardiac norepinephrine is even more potently reduced
316
2. ANALOGS OF ENZYME REACTION COMPONENTS
100
75
50
25-
% OF
NORMAL
SEROTONIN /
/
/ /
/dopamine
^^
/
y"^ norepinephrine
\y
1 1
20
time (HOURS) —
40
60
80
100
Fig. 2-4. Effects of a-methyl-w-tyrosine on brain amine concentrations in the
guinea pig following an intraperitoneal dose of 400 mg/kg. (From Hess e< a?., 1961.)
and remains at a lower level for a longer time than in brain. The results
on serotonin and dopamine levels in the brain correspond as expected to
the time course of decarboxylase inhibition, but the prolonged depletion
of norepinephrine is difficult to explain on this basis. Since dopamine levels
return to normal long before norepinephrine, there must be either an in-
hibition of the /?-hydroxylation of dopamine to norepinephrine or an in-
terference with the tissue binding of norepinephrine. Hess et al. (1961)
showed that these analogs inhibit /5-hydroxyIation only at relatively high
concentrations, which might have been produced soon after injection but
certainly would not occur several hours later, and thus inclined to the sec-
ond explanation. The lack of inhibition of dopamine /5-oxidase has been
confirmed by Creveling et al. (1962)
The time course for catecholamine depletion in mouse brain and heart
following administration of these analogs is similar to that following re-
serpine, except the return toward normal in the brain is somewhat faster.
Porter et al. (1961) examined different analogs for ability to reduce nor-
epinephrine in the brain and heart, and compared these results with their
effectiveness in inhibiting decarboxylation of 5-hydroxytryptophan in
kidney (Table 2-12). Some lack of correlation between the two activities is
TYROSINE METABOLISM
317
Table 2-12
Effective Doses of Analogs in Inhibiting Kidney Decarboxylase
AND Lowering Brain and Heart Norepinephrine in Mice "
Analog
ED^o (mg/kg)
Inhibition
of renal
decarboxylase
Depletion
of norepinephrine
Brain
Heart
L-a-Methyl-3,4-dopa 2.63 32 21
L-a-Methyl-2,3-dopa 1.35 >100 >100
L-a-Methyl-3-hydroxy-PA 11.7 12 1
L-a-Methyldopamine No inhibition >100 5
L-a-Methyl-3-hydroxyphenylethylamine No inhibition 33 0.7
" Analogs injected intraperitoneally. Kidney decarboxylase activity determined
45 min after injection. Doses required to half deplete tissues of norepineplirine in
last two columns. Since it has been shown that only the L-isomers are active, results
are given on this basis for more convenient comparison. (Data from Porter et al., 1961.)
100
0 5
TIME (HOURS)
Fig. 2-5. Inhibition of amino acid decarboxylase in guinea pig tissues by a-methyl-
TO-tyrosine injected intraperitoneally at 400 mg/kg. (From Hess et al., 1961.)
318 2. ANALOGS OF ENZYME REACTION COMPONENTS
evident and led to the conclusion that an action other than decarboxylase
inhibition is involved, this probably being an interference with the binding
of amines in the tissues. Since the a-methylamino acid analogs can be
slowly decarboxylated to the corresponding a-methylamines in the body,
and since these amines have the ability to deplete norepinephrine, it was
suggested that at least part of the tissue amine lowering is due to displace-
ment by the a-methyl analogs of the amines. This mechanism has been
subscribed to by several recent workers. Maitra and Staehelin (1963) ad-
ministered a-methyldopa to rats and guinea pigs and found the cardiac
catecholamine levels to be insignificantly altered. They detected an increase
in the a-methylnorepinephrine level, however, and a corresponding de-
crease in norepinephrine, indicating the displacement of the normal catechol-
amine with its analog. MuschoU and Maitra (1963) further demonstrated
that a-methylnorepinephrine stored in the sympathetic nerve endings is
released by nerve stimulation and is active on various adrenergic receptors.
Pletscher et al. (1964) after injecting a-methyldopa into rats, found marked
reduction of brain serotonin several hours later and felt that inhibition of
the decarboxylase could not explain the results. They inclined to the view
that a-methyldopa must also release or displace stored amines, and might
also interfere with the uptake of amino acids by the brain. However, S. E.
Smith (1963) had shown that a-methyldopa is only a very weak inhibitor
of 5-hydroxy tryptophan uptake in brain slices (50% inhibition at 6.7 vaM),
although it inhibits the decarboxylase potently (50% inhibition at 0.00056
mM). It may be noted that other analogs may inhibit uptake more than
decarboxylation. Day and Rand (1964) showed that a-methyldopa can
restore the activity in animals whose catecholamine levels have been de-
pleted by treatment with reserpine, presumably by the formation of a-
methylnorepinephrine, which is generally only 1/9-1/2 as pharmacologically
potent as norepinephrine; this does not provide direct evidence for the
mechanism of inhibition by a-methyldopa, but clearly shows that it forms
an active amine analog.
The principles involved in the interpretation of these results are impor-
tant in the general field of analog inhibition and the disturbances produced
in tissue function, and, furthermore, the foregoing experiments might be
carelessly construed as invalidating the decarboxylase inhibition mechanism;
thus some critical comments may not be out of place.
(1) It is unfortunate that Porter etal. (1961) did not determine decarbox-
ylase inhibition in brain and heart for comparison with amine depletion
in these tissues, since the inhibition in kidney may be quite different. In
the first place, the penetration of the analogs into the three tissues may
vary. Indeed, Hess et al. (1961) found that a-methyldopa concentrations
in brain, heart, and kidney are in the ratio 1:1.66:3.28 at 1 hr and 1:1.8:9.8
at 5 hr after injection. The concentrations of the other analogs in the tis-
TYROSINE METABOLISM 319
sues are not known, but it was demonstrated that a-methyldopa does not
penetrate into brain. In the second place, the decarboxylases from the var-
ious tissues may have different susceptibilities to the analogs, as we have
noted above. Perhaps cardiac decarboxylase is resistant to a-methyldopa
while the renal enzyme is more sensitive.
(2) Monoamine oxidase inhibitors were administered during norepineph-
rine depletion, and norepinephrine levels immediately rose (Hess et al.,
1961). It was concluded that "biosynthesis of norepinephrine can still
occur in animals treated with or-methylamino acids." All these results
mean is that inhibition of the decarboxylase was not complete. In any
sequence
AillB^C
an inhibition of (1) will lower B and an inhibition of (2) will raise B (see
Chapter 1-7).
(3) It is stated that the decarboxylase step is the most rapid in the
over-all sequence and therefore cannot be rate-limiting (Hess et al., 1961).
The conclusion was that the decarboxylase must be inhibited very strongly
for any effect to be observed. First, it is very difficult to establish that
very very
Tyrosine — > dopa — > dopamine — > norepinephrine
sloiv fast slow
these are the relative rates of the reactions in vivo, where the concentra-
tions and states of the enzymes are quite different than when isolated from
the cells. Second, the rate of formation of norepinephrine in a steady state
is controlled by the first reaction (or a previous reaction) since these
reactions are virtually irreversible. An inhibition of dopa decarboxylase
will not alter the rate of norepinephrine formation as long as a steady
state is maintained; the concentration of dopamine will also be unchanged
in the steady state. However, it has been demonstrated that dopamine
concentration falls, indicating that a departure from a steady state has
occurred. One must also consider the other possible metabolic pathways
for dopamine (e.g., oxidation and 0-methylation), since this is a divergent
sequence. A certain depression of the decarboxylation need not be reflected
in the same depression of norepinephrine formation, even under nonsteady-
state conditions; the latter can be either greater or less than the inhibition
of dopamine formation. In the case of serotonin formation, the decar-
boxylation is the last step, and whether it will be inhibited or not will
depend on the degree to which 5-hydroxytryptophan concentration can
rise to overcome the block. In any event, it has been shown that the
in vivo inhibition of decarboxylase by these analogs can be very high
and sometimes complete.
320 2. ANALOGS OF ENZYME REACTION COMPONENTS
(4) The fact that certain a-methyl analogs of the catecholamines can
deplete tissues of the amines, although they do not inhibit the decarboxy-
lase, is not evidence against a decarboxylase inhibition mechanism for the
a-methylamino acids, but indicates another mechanism, which may play
a role in the prolonged lowering of norepinephrine levels without neces-
sarily being involved in the initial rapid fall in tissue amines. The relatively
rapid return of serotonin and dopamine levels to normal (Fig. 2-4) suggests
that there is no generalized disturbance in tissue amine binding, but that
the effect is specifically on norepinephrine. The most satisfactory position
at the present time might be the following: the initial marked fall in tissue
amines brought about by the a-methyl analogs is primarily due to an in-
hibition of decarboxylation (perhaps supplemented at peak concentrations
by inhibition of other steps, such as /?-hydroxylation), and further distur-
bances in amine binding are progressively produced by the a-methylamines
formed from the inhibitors, so that even when the decarboxylase is normally
active again the tissues cannot concentrate certain of the normal catechol-
amines.
Dopamine p-Hydroxylase
This enzyme catalyzes the synthesis of norepinephrine from dopamine
and, as we have seen, its inhibition by analogs could be both theoretically
and practically important. Hess et al. (1961) found that a-methyl-3-hy-
droxyphenylalanine does not inhibit at 2 tclM. but inhibits 50% at 4 toM.
The concentration of a-methyldopa 1 hr after injection is given as 376
//g/g in the heart and this could mean a concentration around 750 //g/ml
of intracellular fluid (assuming extracellular fluid has a low concentration
at this time). This is approximately equivalent to 4 mM so that appreciable
inhibition might occur. Inhibition data indicate that 3-methyl-3-hydroxy-
phenylalanine concentrations in the tissues are roughly the same as a-
methyldopa concentrations. Until more is known about the nature of the
inhibition of this enzyme, it might be safe to conclude that it plays some
role in the effects of the or-methyl analogs of phenylalanine.
Various amines can inhibit this enzyme (Goldstein and Contrera, 1961).
When dopamine concentration is 0.26 niM, the following inhibitions are
observed: tyramine at 2.9 raM (75%), /?-phenylethylamine at 3.3 vaM
(45%), amphetamine at 5.9 milf (35%), and 3-methoxydopamine at 4.8
mM (15%). None of these inhibitors appears to be potent enough to be
practically important in reducing norepinephrine synthesis and, further-
more, these amines are so pharmacologically active that their use is limited.
Benzyloxy amine, and particularly the p-hydroxyl derivative, inhibit this
enzyme rather potently, 0.01 mM of the latter blocking almost completely
after 90 min (van der Schoot et al., 1963), this being attributed to the iso-
steric relation between phenethylamines and benzyloxyamines.
i
TRYPTOPHAN METABOLISM 321
TRYPTOPHAN METABOLISM
Tryptophan is involved in several important metabolic pathways, form-
ing active substances as well as being incorporated into proteins, so that
many attempts to block these pathways specifically with analogs have
been made. Growth inhibition and physiological disturbances are readily
produced by many of these analogs. (See scheme on page 322).
Synthesis of Tryptophan
L-Tryptophan is a potent feedback inhibitor of the conversion of 5-
phosphoshikimate to anthranilate, an early reaction in tryptophan biosyn-
thesis, and 5-methyltryptophan also inhibits, although not so strongly,
a phenomenon (i.e., inhibition of a biosynthetic step by an analog) termed
false feedback inhibition by Moyed (1960). It is likely that this mechanism
explains the bacteriostatic activity of this analog. The condensation of
anthranilate and 5-phosphoribosyl-l-pyrophosphate is not inhibited by
5-methyltryptophan, but 6-fiuorotryptophan is inhibitory. A later reac-
COO
NH,
Tryptophan Anthranilate
tion in this sequence, the conversion of anthranilic deoxyribonucleotide to
indoleglycerol-3-phosphate, is inhibited by a variety of anthranilate de-
rivatives, especially the 3- and 4-methyl analogs (Gibson and Yanofsky,
1960). The final reaction, the condensation of indole and serine to form
tryptophan, catalyzed by tryptophan synthetase, is a major site of the
attack by 4-methyltryptophan, which is a bacterial growth inhibitor
(Trudinger and Cohen, 1956). The 5- and 6-methyl indoles are fairly potent
competitive inhibitors, with K^ values near 0.1 roM (Hall et al., 1962).
They are also antibacterial. The growth depression of E. coli is counteracted
by tryptophan (Fig. 2-6). At least two sites for the inhibition have been
demonstrated. Tryptophan synthetase is inhibited competitively, but there
is also a block of the much earlier formation of anthranilate. There are no
effects on the immediate metabolism of anthranilate or on tryptophanase,
which indeed readily splits the analog to 4-methylindole. The bacteriostatic
action is probably due mainly to suppression of tryptophan synthesis rather
than to a disturbance of tryptophan utilization. Thus several steps in the
biosynthesis are susceptible to analogs and it is quite possible that other
322
2. ANALOGS OF ENZYME EEACTION COMPONENTS
™ rt
O) T3
+ a
x:
a
o
S S S
^ eg CO
o
x:
a,
TRYPTOPHAN METABOLISM
323
unstudied reactions are likewise inhibited. A more indirect mechanism is
the inhibition of the synthesis of tryptophan synthetase in growing cells,
whereby tryptophan formation is further reduced.
100
Fio. 2-6. The inhibition of E. coli growth by 4-methyltryptophan
at 0.01 mM and its antagonism by increasing tryptophan con-
centrations. (From Trudinger and Cohen, 1956.)
Tryptophanase
This bacterial and fungal enzyme may be involved in the fermentation
of tryptophan and is responsible for the putrefactive reaction in the in-
testines. It is potently and competitively inhibited by the product indole
and less potently by some analogs (as shown in the accompanying tabula-
Inhibitor
Relative — AF of binding
(kcal/mole)
Indole
/?-3-Indolylpropionate
3-Indolylacetate
/?-3-Indolylethylamine
DL-/3-l-Methyl-3-indolylalanine
5.40
4.05
3.76
2.55
1.44
324 2. ANALOGS OF ENZYME REACTION COMPONENTS
tion) (Gooder and Happold, 1954). The importance of the indole N in
binding is indicated by the weak inhibition with the last analog (tryptophan
with the indole N methylated) and the failure of indene (the hydrocarbon
/
.CHoCOO" /\ /CH,— CH
NH^
IJ COO'
N' "^ N
I I
H CH3
3-Indolylacetate 0-1 -Methyl- 3 -indolylalanine
.CHoCHoNH,
N
I
H
0-3-Indolylethylamine
analog of indole) to inhibit, while the importance of the carboxylate group
is reflected in the weak inhibition by the indolylethylamine. The potency
of indole may be attributed to the fact that it may not have to be oriented
in a manner necessary for reaction of the side chain.
Tryptophan Pyrrolase (Tryptophan Peroxidase)
This enzyme initiates one of the most important catabolic pathways of
tryptophan and is readily inhibited by certain analogs. Hayaishi (1955 b)
found the Pseudomonas enzyme to be sensitive to the hydroxytryptophans,
and calculated the values of K; shown in the following tabulation. Since
Substance
Ki or Kg Relative — Zli^ of binding
(milf) (kcal/mole)
5- Hydroxy tryptophan 0 . 002 8 . 06
7-Hydroxytryptophan 0.12 5.55
L-Tryptophan 0.4 4.81
5-hydroxytryptophan is normally formed from tryptophan on the pathway
to serotonin, its potent inhibition of the pyrrolase suggests that it may
play a role in regulating tryptophan metabolism. The enzyme from rat
liver is also inhibited by 5-hydroxytryptophan and even more potently
TRYPTOPHAN METABOLISM 325
by serotonin (Frieden et al., 1961). The K/s in the following tabulation
indicate 3-indolylacrylate to be the most effective inhibitor. Other inhibi-
tions observed when (S) = (I) = 3 mM are: indole 69%, tryptazan 50%,
Inhibitor
K,
Relative — Ji" of binding
(mM)
(kcal/mole)
3-Indolylacrylate
0.012
6.99
Serotonin
0.067
5.93
5-HydroxytrjT)tophan
0.094
5.71
3-Indolylbutyrate
0.16
5.39
Tryptamine
0.20
5.25
3-Indolylpropionate
0.29
5.02
3-Indolylacetate
0.91
4.32
^-Methyltryptophan
1.1
4.20
D-Tryptophan
1.6
3.97
6-Fluorotr>i)tophan
2.0
3.83
5-Fluorotryptophan
2.2
3.77
5-methyltryptophan 38%, and 6-methyltryptophan 33%. The analogs with
altered side chains are competitive while the others are mainly noncompeti-
tive. The roughly equivalent binding of tryptamine and 3-indolylpropionate,
and of serotonin and 5-hydroxytryptophan, might indicate that the binding
is primarily with the indole ring, the side chains contributing little, and this
is substantiated by the fact that indole is bound approximately as well
as 3-indolylpropionate. The stronger binding of 3-indolylacrylate compared
to 3-indolylpropionate (about 2 kcal/mole) is thus difficult to account for
unless there is a modification of the interaction of the indole N. Tryptophan
pyrrolase is an inducible enzyme in the rat but none of these analogs is
active, although Sourkes and Townsend (1955) found a-methyltryptophan
to induce after subcutaneous injection.
Tryptophan Hydroxylase (Phenylalanine Hydroxylase)
Excessive feeding of phenylalanine leads to low blood serotonin, a low
excretion of 5-hydroxyindoleacetate, and a decrease in brain serotonin,
and this has usually been attributed to an inhibition of 5-hydroxytrypto-
phan decarboxylase. However, no accumulation of 5-hydroxytryptophan
has been demonstrated and it is possible that the site of the inhibition might
be earlier, perhaps on the hydroxy lation of tryptophan (Freedland et al.,
1961). Hydroxylating preparations from rat liver are indeed quite potently
inhibited by L-phenylalanine, and also by phenylpyruvate and phenyUac-
tate (K,„ for L-tryptophan is 29 mM, and K^ for L-phenylalanine is 0.22 mM).
326 2. ANALOGS OF ENZYME REACTION COMPONENTS
It is likely that the same enzyme is responsible for the hydroxylation of
both phenylalanine and tryptophan, since the K^ for phenylalanine is close
to the iii,,, when it is the substrate; the affinity for tryptophan is, however,
much less. These results may help to explain some of the changes observed
in phenylpyruvic oligophrenia (see pages 329 and 429).
Tryptophan-Activating Enzyme
An activating enzyme from pancreas is specific for tryptophan with
respect to other normal amino acids but can activate certain analogs of
tryptophan (Sharon and Lipmann, 1957). The analogs tested fall into three
categories:
Group I are activated (tryptazan, azatryptophan, 5-fluorotryptophan,
and 6-fluorotryptophan).
Group 11 are inhibitory (tryptamine, D-tryptophan, /5-methyltryptophan,
5-hydroxytryptophan, 5-methyltryptophan, and 6-methyltryptophan).
Group III are inactive (indole, indoleacetate, 6-methyltryptazan, and
iV-acety Itry ptophan ) .
CH,— CH ^-^^ CH^— CH
^coo' f H H coo'
N N N
I I
H H
r
Tryptazan Azatryptophan
Reciprocal plots were said to indicate competitive inhibition but actually
do not show pure competition, and it might better be designated as partially
competitive inhibition. For analogs to be activated they must match the
size of tryptophan, and the introduction of bulkier groups prevents reaction
with ATP but allows binding and inhibition. Azatryptophan and tryptazan
are both incorporated into proteins. In E. coli, azatryptophan permits syn-
thesis of proteins and nucleic acids but many of the enzymes are not in
the normal forms (Pardee and Prestidge, 1958). The synthesis of adaptive
enzymes, such as /?-galactosidase, is inhibited very rapidly, and phage for-
mation is blocked more readily than bacterial growth. The induced synthe-
sis of maltase in yeast is also very potently suppressed by tryptazan, in-
corporation of all amino acids being simultaneously blocked (Halvorson
etal., 1955). 5-Methyltryptophan and 6-methyltryptazan inhibit moderate-
ly, while 6-methyltryptophan is inactive.
GLUTAMATE METABOLISM
327
GLUTAMATE METABOLISM
Glutamate occupies a central position in many important metabolic
pathways and serves to link amino acid metabolism with the tricarboxylic
acid cycle. Its relationship to the biochemically active glutamine and the
physiologically active /-aminobutyrate (GABA) makes possible specific
inhibitions of glutamate reactions of great interest. The reactions catalyzed
by enzymes studied with respect to analog inhibition are shown in the
following scheme.
glutamylhydroxamate
D -glutamate
(4)
o-ketoglutarate
(7)
(3) (8)
glutamine
(6)
(1,2)
L-glutamate —
(5)
r
>' -aminobuty rate
(10)
proline
A'-pyrroline-5-carboxylate
a
(10)
glutamic -7 -semialdehyde
(1) L-glutamate dehydrogenase
(2) glutamate transaminases
(3) glutamate racemase
(4) D-glutamate oxidase
(5) glutamate decarboxylase
(6) glutamine synthetase
(7) glutaminase
(8) formylglycinamidine phosphoriboside synthetase
(9) 7 -glutamyl transferase
(10) A'-pyrroline-5-carboxylate dehydrogenase
Glutamate Decarboxylase
Several analogs of glutamate inhibit its utilization by Lactobacillus ara-
binosus, and thus a study of decarboxylase from E. coli was undertaken
by Roberts (1953). The two most potent inhibitors are a-oximinoglutarate
and a-methylglutamate, but the former is probably active by virtue of
its hydrolysis to hydroxylamine which inactivates pyridoxal phosphate.
The latter analog inhibits competitively when added with the substrate,
but if it is preincubated with the enzyme the inhibition becomes progres-
sively more noncompetitive and difficulty reversible. The rates of combina-
tion with the enzyme and dissociation from the enzyme are very slow.
The methyl group interferes with the normal binding of the molecule so
that decarboxylation does not occur, but by some unknown mechanism
brings about a type of binding that is very strong, a behavior seen with
some other a-methylamino acids. The decarboxylation of glutamate in
rat brain homogenates is also inhibited competitively by aspartate {K„^
328 2. ANALOGS OF ENZYME REACTION COMPONENTS
= 21 mM, and K^ = 23 milf ) (Wingo and Awapara, 1950), which is rather
surprising because of the shorter intercarboxylate distance.
Ghitamate decarboxylase from the squash Curcurbita moscliata is inhib-
ited competitively by a variety of organic acids (see accompanying tabu-
lation), and the results are of some interest with regard to structure and
Inhibitor (13.6 mM) % Inhibition
Monocarboxylates
Formate 17
Acetate 42
Propionate 10
7i-Butyrate 18
Isobutyrate 8
n-Valerate 25
Iso valerate 15
n-Caproate 24
Isocaproate 20
Dicarboxylates (saturated)
Oxalate 0
Malonate 0
Succinate 0
Glutarate 24
Adipate 49
Pimelate 58
Suberate 37
Dicarboxylates (unsaturated)
Fumarate 0
Maleate 14
Citraconate 0
Mesaconate 7
Itaconate 11
Tricarboxylates
cis-Aconitate 29
irans-Aconitate 18
intercarboxylate distance (Ohno and Okunuki, 1962). Glutamate concen-
tration was 27.2 mM in all cases. Many amino acids examined are weakly
inhibitory or without effect. The maximal inhibition by pimelate in its
series probably indicates that a cambering of the molecule is necessary for
binding of the two carboxylate groups, or possibly that the cationic groups
of the enzyme are farther apart than in glutamate. The relatively high
Relative
— Zli^of bindin
g (kcal/mole)"
Hanson (1958)
Tashian (1961)
3.15
4.16
—
4.26
2.62
2.62
2.53
3.73
1.75
3.83
0.60
—
0.53
—
GLUTAMATE METABOLISM 329
inhibition by acetate is surprising; if the monocarboxylates interact with
the cationic groups, one might expect propionate or butyrate to be more
inhibitory.
The problem of the abnormal brain development in phenylpyruvic oligo-
phrenia prompted an investigation of the effects of the phenyl acids on
brain glutamate decarboxylase by Hanson (1958); the results are presented
in the tabulation below in which they are compared with those of Tashian
Inhibitor
p-Hydroxyphenylacetate
o-Hydroxyphenylacetate
Phenylpyruvate
Phenylacetate
p-Hydroxyphenylpyruvate
Phenylalanine
Phenyllactate
° Relative — AF's of binding adjusted so that value for phenylpyruvate is the same
in each series.
(1961), who also used rat brain. There are some rather striking differences,
part of which may be due to the procedures used since these analogs,
although stated to be competitive, present deviations from classic kinetic
formulations (and for this reason the binding energies calculated from ap-
parent K/s are probably not very reliable). If these analogs, which are
present in high concentrations in the blood enter the brain readily, it is
possible that they depress the formation of y-aminobutyrate (GABA)
which may be essential for normal neurological development. In branched-
chain ketonuria (maple sugar urine disease) various keto and hydroxy
fatty acids accumulate in the body, which Tashian (1961) showed also
inhibit glutamate decarboxylase (relative — JF's of binding for a-hydroxy-
isovalerate, a-ketoisovalerate, and the corresponding isocaproates from
3.58 to 4.50 kcal/mole on the scale above). The enzyme from E. coli, on
the other hand, is relatively insensitive to any of these analogs.
L-Glutamate Dehydrogenase
The oxidative deamination of L-glutamate in beef liver homogenates is
catalyzed by a NAD-linked dehydrogenase, the inhibition of which at
pH 8.4 was thoroughly studied by Caughey et al. (1957). The accompanying
tabulation shows the K^ values for competitive analogs, from which the
330 2. ANALOGS OF ENZYME REACTION COMPONENTS
Inhibitor
(mM)
Relative — Zli*' of binding
(kcal/mole)
5-Bromofuroate
0.059
6.00
5-Chlorofuroate
0.063
5.96
5-Nitrofuroate
0.17
5.35
m-Iodobenzoate
0.46
4.74
»n-Bromobenzoate
0.54
4.65
Isophthalate
0.66
4.62
Glutarate
0.58
4.60
a-Ketoglutarate
0.73
4.45
w -Chlorobenzoate
1.02
4.25
D-Glutamate
2.0
3.83
m-Nitrobenzoate
3.4
3.51
Trimesate
4.0
3.40
Fumarate
6.8
3.08
Succinate
11
2.78
Adipate
16
2.55
relative binding energies have been calculated. Inhibitors were classed as
competitive if the interaction constant a is greater than 10; trimesate and
D-glutamate are only partially competitive, with a = 1.7. It was postulated
CCXD'
^O. ^COO"
COO
Furcate Isophthalate
COO
COO'
OOC \^ COO
Trimesate Indole-3-carboxylate
that the enzyme contains two cationic groups at a separation optimal for
interaction with the carboxylate groups of glutamate, glutarate, and iso-
phthalate. The effects of some other inhibitors for which the ^/s were
not calculated are shown in the following tabulation; most of these are
GLUT AM ATE METABOLISM 331
relatively weak (fumarate is included for comparison with values in the
preceding table). L-Glutamate was 2 mM in each case. The following are
not inhibitory at 10 mJ-l: L-glutamine, L-diethylglutamate, /?-methylglu-
tarate, citrate, o- and p-hydroxybenzoate.
Inhibitor
Concentration
(mM)
% Inhibition
Benzoate
10
12
Furoate
10
26
Phthalate
10
17
Terephthalate
6.7
16
Isophthalate
2
50
<ran.5-Aconitate
8
20
Indole-2-carboxylate
7.5
21
Indole-3-carboxylate
8.3
38
m-Hydroxybenzoate
10
27
Pyridine-2,6-dicarboxylate
10
27
Fumarate
10
29
In the various substituted benzenes the 7neta compounds are invariably
the most potent inhibitors, presumably because the intergroup distances
are close to the enzyme intercationic separation. The dipoles of the halogen
and nitro compounds may interact with the cationic group since they may
be represented as:
However, there is no correlation of inhibitory activity with dipole moment.
One might think that the dipole-cation interaction would be weaker than
the carboxylate-cation interaction, but the hydration of the inhibitors must
also be considered. Less water needs to be displaced when the dipoles ap-
proach the enzyme cationic group. It was pointed out that all good in-
hibitors are reasonably planar and the presence of bulky groups protruding
lowers the affinity. The particularly good binding of the furoates may be
related to some interaction of the ring 0 with the enzyme. The reverse
reaction from or-ketoglutarate to glutamate is inhibited less than the for-
ward reaction by glutarate, isophthalate, and 5-bromofuroate, and the
inhibitions are noncompetitive.
The L-glutamate dehydrogenase from cockroach muscle mitochondria
332 2. ANALOGS OF p:;nzyme reaction components
is similarly inhibited (see' accompanying tabulation), and here glutarate
also appears to present a relatively good fit to the active site (Mills and
Cochran, 1963). Glutamate was 15 mM in all cases. The reverse reaction
catalyzed by the glutamate dehydrogenases (both NAD- and NADP-
Inhibitor (3 mM)
% Inhibition
Succinate
15
Fumarate
15
Malate
15
Glutarate
65
Adipate
30
D-Glutamate
55
L- Aspartate
20
D-Aspartate
10
linked) from Fusarium oxysporum is also inhibited by glutarate, the K,^
for a-ketoglutarate being 2.1 mM, and the K^ for glutarate 1.52 mM
(Sanwal, 1961).
Glutaminase
The deamidation of glutamine is inhibited by the product glutamate
and this is not a reversal of the equilibrium but a competition for the
active center, as first pointed out by Krebs (1935). L-Glutamate and d-
glutamate inhibit guinea pig kidney glutaminase equally (98% at 25 mM
when glutamine is 8.7 mM). Inhibition by glutamate has been confirmed
for the enzyme from guinea pig kidney (van Baerle et al., 1957), pig kidney
(Klingman and Handler, 1958), dog kidney (Sayre and Roberts, 1958),
and rat brain (Blumson, 1957). The inhibition has generally been found
to be noncompetitive with respect to glutamine, but oddly is competitive
with phosphate on the phosphate-activated glutaminase from dog kidney.
The ammonium ion is, however, strictly competitive with glutamine on
both the pig and dog kidney enzymes. Another type of glutaminase (called
glutaminase II), which is transaminating in the presence of pyruvate and
is obtained from guinea pig kidney, is not inhibited by even 100 mM
glutamate (Goldstein et al., 1957). Sayre and Roberts (1958) pictured the
active center as containing two cationic groups, one binding the phosphate
and one the glutamine carboxylate group; the negatively charged phosphate
also interacts with the positive or-amino group of glutamine. Since the active
enzyme is the phosphate complex, it is easy to see why phosphate would
antagonize inhibitions produced by certain substances (e.g., dyes such as
bromosulfalein or bromcresol green which complex with both enzyme
cationic sites), but it is difficult to understand why glutamate inhibits
GLUTAMATE METABOLISM 333
competitively with respect to phosphate. It was stated that for a substance
to compete with glutamine it should have affinity for the enzyme-phosphate
complex, and it would seem that glutamate may fall into this category.
Few other analogs have been tested on this enzyme. Krebs (1935) ob-
served a mild inhibition by DL-/?-hydroxyglutamate (22% at 80 mM when
glutamine 40 mM), and Girerd et al. (1958) reported inhibition by ethyl
D-glutamate and DL-/5-methylglutamate. Because of the postulated role
of glutaminase in renal function, these two latter analogs, along with
L-glutamate and bromosulfalein, were tested in vivo. These inhibitors re-
duce diuresis in rats around 50% at 50 mg/kg subcutaneously, whereas a
group of five less potent glutaminase inhibitors actually increase diuresis.
Formylglycinamidine Phosphoriboside Synthetase
Glutamine participates in purine biosynthesis by contributing its amide
N. Azaserine and 6-diazo-5-oxo-L-norleucine (DON) are potent inhibitors
of inosinate biosynthesis and lead to the accumulation of formylglycina-
mide phosphoribotide (FGAR). These substances may be considered as
analogs of glutamine and have been shown to inhibit formylglycinamide
ribonucleotide amidotransferase competitively with respect to glutamine
N:=N=CH-
-CO— O-CH2-
Azaserine
-CH
COO'
N=N=CH-
NH3
/
-CO— CH2CH2— CH
^COO
DON
(Levenberg et al., 1957). The K„, for glutamine is 0.615 mM and the K^s
for azaserine and DON are 0.034 mM and 0.0011 mM, respectively. Once
azaserine binds to the enzyme, however, an irreversible reaction occurs,
due perhaps to an alkylation of the enzyme. French et al. (1963 a) pointed
out that 50% inhibition can be obtained with a (S)/(I) ratio of 2100 with
DON. Phosphoribosyl-PP amidotransferase, another enzyme catalyzing
the transfer of the amide nitrogen of glutamine, is also inhibited compe-
titively by DON, with a K^ of 0.019 mM, and much more weakly by aza-
serine (Hartman, 1963 b). A slow covalent binding of DON to the enzyme
occurs following the initial reversible attachment, and this is accelerated
by the presence of phosphoribosyl-PP and Mg++ on the enzyme, indicating
that the active site for the reaction of glutamine, or the binding of DON,
is partly dependent on the other substrate and the cofactor. Blocking of
an SH group prevents the attachment of DON to the enzyme, suggesting
that this SH group is catalytically functional in the nitrogen transfer and
the irreversible binding of DON, as French et al. (1963 b) concluded from
their work with azaserine on the formylglycinamide ribonucleotide amido-
transferase.
334 2. ANALOGS OF ENZYME REACTION COMPONENTS
Glutamate Transaminases
This enzyme apparently possesses two cationic groups properly separated
to interact with glutamate and the other dicarboxylates, because it is inhib-
ited best by glutarate of all the saturated dicarboxylates, as shown in the
accompanying tabulation (aspartate =1.7 mM and a-ketoglutarate = 6.7
mM) (Jenkins et al., 1959). This is one instance in which the «-methyl
analog has no affinity for the enzyme. Very similar results were obtained
Inhibitor (40 mM)
Malonate
Succinate
Glutarate
Adipate
Piraelate
Suberate
Maleate
a-Methylaspartate
a-Methylglutaniate
by Velick and Vavra (1962), glutarate inhibiting the most potently of the
saturated dicarboxylates; from their values for K, one can calculate that
glutarate is bound 0.85 kcal/mole more tightly than succinate, and 0.32
kcal/mole more tightly than adipate. Of the three phthalates, o-phthalate
is the most inhibitory {K^ = 5 mM), m-phthalate intermediary (K^ = 8
mM), and j^-phthalate the least active {K, = 10 mM). The results of the
studies above were obtained on pig heart transaminase, and from the lim-
ited data reported by Goldstone and Adams (1962) it would appear that
the enzyme from rat liver is different in that glutarate is about 10 times
more potent than maleate as an inhibitor. The alanine:a-ketoglutarate
transaminase from rat liver is inhibited moderately and competitively by
certain amino acids, such as leucine and valine, and also by maleate, but
no data were given for comparison with the aspartate: a-ketoglutarate trans-
aminase (Segal et al., 1962). Fluorooxalacetate inhibits the latter enzyme
from heart competitively with respect to oxalacetate (and to a-ketoglutarate
and aspartate when the reverse reaction is run), K^,, being 3.5mMforof-
ketoglutarate and 0.5 mM for aspartate, and /f, being 0.1 mM (Kun et
al., 1960). It is also slowly transaminated to fluoroaspartate which likewise
inhibits the enzyme.
Inhibition
Relative
- AF (kcal/mole)
0
<0.17
31
1.49
72
2.56
62
2.29
0
<0.17
0
<0.17
78
2.76
35
1.60
0
<0.17
ARGINASE 335
Other Enzymes in Glutamate Metabolism
Analog inhibition has been reported for several other enzymes involved
in the metabolism of glutamate but quantitative studies from which struc-
ture-action relationships may be derived have not been made. Some of
these results are summarized in Table 2-13. An interesting inhibitor is the
convulsant isolated from agenized flour, methionine sulfoximine, which is
inhibitory to the incorporation of methionine into proteins in bacteria
and brain. Since these actions are to a great extent antagonized by methio-
nine, this substance has generally been considered as a methionine antago-
nist, but glutamine also is antagonistic. Sellinger and Weiler (1963) have
shown that methionine sulfoximine inhibits brain glutamine synthetase
competitively with respect to glutamate, some inhibition being seen at
0.01 mM and around 50% inhibition at 1 mM with low glutamate concen-
trations {K^ = 0.05-0.064 mM). The relation between this inhibition and
the convulsant action is not clear but it was postulated that methionine
sulfoximine interferes in some vague manner with glutamine synthesis
in an intracellular compartment in the brain.
ARGINASE
The hydrolysis of arginine:
\ / (+H2O) + / \
C— NH— CHoCH^CH,— CH ^ ^-^- H3N— CHoCHoCHz— CH + CO
+ <^ \ - \ - /
HgN COO COO H2N
Arginine Ornithine Urea
is catalyzed by the Mn++-activated enzyme arginase and is a step in the
urea cycle. Arginase is inhibited by the product ornithine, as first shown
by Gross (1921) and confirmed by Bach and Williamson (1942), who found
that the inhibition is much more marked in liver extracts than in slices
(50% inhibition given by 5.3 mM ornithine in extracts and by 15.9 mM
in slices when argmine is 3.56 mM). Edlbacher and Zeller (1936) noted that
several amino acids inhibit, but ornithine is the most potent with lysine
running a close second.
These inhibitions were compared and subjected to quantitative treat-
ment by Hunter and Downs (1945) in the publication wherein the first
use of the single-curve plot (type F) was made (see Chapter 1-5). Orni-
thine and lysine inhibit competitively {K , = 4.1 mM and 4.8 mM, respec-
tively) but other amino acids are usually only partially competitive (since
the DL-forms of the inhibitors used and only the L-isomers are active, it is
likely that these constants should be halved). Calculations of relative
336
2. ANALOGS OF ENZYME REACTION COMPONENTS
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ARGINASE 337
interaction energies from the constants given are shown in the following
tabulation. Two types of inhibition may be possible: (1) competitive inhi-
bition by diamino compounds that are probably oriented as the substrate,
and (2) partially competitive or noncompetitive inhibition by monoamines
^ . ., . Relative — Ji^ of binding
Inhibitor ,, ,, , ,
(kcal/mole)
L-Ornithine 3.82
L-Lysine 3 . 72
L-Norvaline 2 . 53
L-Isoleucine 2 . 48
L- Valine 1.98
L-Cysteine 1 . 94
L-Leucine 1.81
L-a-AminobutjTate 1.40
L-Phenylalanine 1 . 09
L-Xorleucine 0 . 87
L-Proline 0.78
L- Aspartate 0 . 67
L- Alanine 0.61
L-Citrulline 0.47
L-Serine 0.25
L-Tryptophan 0.21
L-Histidine -0.82
Glycine —0.99
that may be oriented otherwise. Inhibitory activity increases in the straight-
chain series from glycine to norvaline; each additional methylene group
contributes 1.17 kcal/mole to the binding energy, a value similar to that
found in other series, and undoubtedly due to dispersion forces. It is odd
that there is a sudden and marked drop in the affinity on adding another
methylene group to form norleucine. As pointed out by Hunter and Downs,
it is difficult to establish structural correlation in the series of substituted
alanines (shown in the accompanying table); why, for example, is cysteine
bound so much more tightly than serine? Substitution in the a-amino group
(carbamyl or formyl) always reduces the activity; hence this probably con-
stitutes one binding group. The experiments were done at pH 8.4 and
therefore all carboxyl groups were essentially completely ionized, but there
would be some variation between the inhibitors with respect to the fraction
of the amino groups protonated {pK^'s for these inhibitors run from 8.2
to 10.6).
338 2. ANALOGS OF ENZYME REACTION COMPONENTS
_,,,., , Relative — AF oi bmdim
bubstituent group
(kcal/mole)
—Imidazole —0.82
—Indole 0.21
-OH 0.25
-H 0.61
-COO- 0.67
-Phenyl 1.09
-CH3 1.40
-SH 1.94
The relative sensitivities to these amino acids probably vary with the
source of the arginase. For example, the mouse liver enzyme is inhibited
somewhat more strongly by L-lysine than L-ornithine, and even D-lysine
is a weak inhibitor (inhibitions at 1 mM are 47% for L-lysine, 42% for orni-
thine, and 8% for D-lysine) (Nadai, 1958). Johnstone (1958) stated that
there is evidence in intact ascites cells that ornithine can interfere with the
transport of arginine into the cells, as well as inhibit arginase intracellularly,
so that this additional site of inhibition must be borne in mind.
An attempt was made by Sen (1959) to determine the effects of L-lysine in
vivo in order to evaluate its possible use in uremia. Bilaterally nephrec-
tomized dogs show an increase of blood urea at a rate of 15-16 mg/day and
die in 80-84 hr. When L-lysine is injected daily and intravenously at a
dose of 1 g, the blood urea rise only 3-4 mg/day and the animals survive
for 274-278 hr. Thus it would seem possible to reduce urea formation in
vivo with this competitive inhibitor, but the clinical benefit of this remains
to be tested.
L-AMINO ACID OXIDASES
These enzymes from snake venoms and mammalian tissues oxidize the
L-isomers of the monoamino-monocarboxylate amino acids, the substrate
used often being L-leucine. The snake venom enzyme was shown by Zeller
and Maritz (1944) to be inhibited by various aromatic sulfonates. Some of
the results are shown in Table 2-14, from which it may be seen that
p-nitrodiphenyl sulfonate is the most potent inhibitor studied. The inhi-
bitions seem to be competitive and a complex between the sulfonate group
and an enzyme amino group was postulated. Benzoate is a rather weak
inhibitor of rat kidney L-amino acid oxidase, 10 mM inhibiting 28%
(Blanchard et al., 1944). However, the ammonium ion is a surprisingly good
inhibitor, 12 mM producing 69% depression of activity.
L-AMINO ACID OXIDASES
339
Table 2-14
Inhibition of Snake Venom l-Amino Acid Oxidase by Sulfonates"
Inhibitor
Sulfonate
(mM)
L-Leucine
(mA/)
Inhibition
HO
N=N-
SO,
2.9
0.01
//_\Vn=n^/ Vso;
2.9
0.01
76
2.9
2.9
2.9
0.01
0.01
0.01
66
55
50
0,N
0.33
1.67
31
0,N
// \V_N„_// V-so;
0.07
0.01
21
a From Zeller and Maritz (1944).
340 2. ANALOGS OF ENZYME REACTION COMPONENTS
The L-amino acid oxidase from the hepatopancreas of Cardium tubercula-
tum is more specific than the venom or kidney oxidases, since many L-amino
acids are not oxidized but are inhibitory (Roche et al., 1959). The inhibitions
by 16.7 mM L-leucine are shown in the following tabulation. At pH 7.6
the inhibitions are competitive, but not at pH 9.2. Apparently this enzyme
can complex with both l- and D-isomers although in no case is a direct
comparison possible.
% Inhibition at:
Amino acid
pH7.6
pH9.2
DL- Alanine
89
71
D-Alanine
39
—
L-Serine
54
73
Glycine
.51
56
L-Glutamate
26
—
L- Proline
24
74
L- Valine
24
30
L-Threonine
20
14
L- Aspartate
13
12
D-Histidine
12
—
D- Leucine
5
—
D-AMINO ACID OXIDASE
D-Amino acid oxidase is also inhibited by certain amino acids, but no
thorough studies have been reported. The enzyme from pig kidney oxidizing
D-leucine is inhibited by DL-leucinamide, DL-leucylglycine, glycyl-DL-
leucine, and DL-leucylglycylglycine, but not glycylglycine (Heimann-
Hollaender and Lichtenstein, 1954). It is interesting that the oxidation
of D-phenylalanine is inhibited by DL-iV-ethylphenylalanine and the oxida-
tion of D-leucine by DL-A^-ethylleucine, since A'-substituted amino acids are
usually not inhibitory for any enzymes acting on amino acids. D-Lysine is
a good inhibitor of this enzyme oxidizing D-alanine (Ky„ = 3.3 mM, and
K^ = 5 mM), but L-lysine is completely inactive (Murachi and Tashiro,
1958). The D-amino acid oxidase from pig kidney with glycine as a substrate
is competitively inhibited by L-leucine {K, = 1 mM) (Neims and Hellerman,
1962). Pyruvate not only competes with D-alanine {K^ = 43 mM) but accel-
erates the photodecomposition of FAD (Yagi and Natsume, 1964). However,
the most interesting and best studied inhibition of D-amino acid oxidase is
that of benzoate and its derivatives, and the opportunity will be taken in
this section of discussing not only the actions of the benzoates on this en-
zyme but also on other enzymes and metabolism in general.
D-AMINO ACID OXIDASE
341
Benzoates and Related Compounds on D-Amino Acid Oxidase
The oxidation of D-alanine in slices and homogenates of rat liver and
kidney was shown to be markedly inhibited by 1 mM benzoate by Klein and
Kamin (1941). A preparation of D-amino acid oxidase from pig kidney was
thus obtained and found to be inhibited 79% by 0.1 mM, this being rever-
sible upon dialysis. Several substituted benzoates are also inhibitory but
all are less potent than benzoate; benzamide is inactive. The inhibitions of
a lamb kidney D-amino acid oxidase by benzoate and j9-amino-benzoate
were shown by Hellerman et al. (1946) to be competitive with substrate.
The rate of spontaneous inactivation of the apoenzyme is reduced by either
substrate or FAD, and benzoate was shown by Burton (1951 a) to have a
comparable action, indicating combination with the active center.
Before discussing the more detailed mechanism of the inhibition we
shall turn to three studies providing information on the structural require-
ments for inhibition. Bartlett (1948) compared many substituted benzoates
(Table 2-15) and found only four to be more potent inhibitors than benzoate,
Table 2-15
Inhibition of Pig Kidney d-Amino Acid Oxidase by Substituted Benzoates "
Substituent
Relative
— Ji^ of binding
(kcal/mole)
ortho
meta
para
F
4.26
6.25
CI
3.27
6.81
5.39
Br
2.11
6.40
5.05
I
2.84
5.39
3.90
OH
4.55
5.20
3.27
NHa
4.26
4.26
2.84
NO,
2.52
5.12
4.90
CH3
2.11
6.25
5.12
OCH3
2.11
3.90
3.78
COOH
1.85
2.11
2.84
None
—
5 . 57
—
" The substrate is DL-alanine at 30 mM and the pH 7.6. Binding energies calculated
from concentrations for 50% inhibition. (Data from Bartlett, 1948.)
pure competitive inhibition with respect to substrate being observed.
J. R. Klein (1953, 1957) demonstrated inhibition, often potent, by various
aromatic carboxylates (Table 2-16), and Frisell et al. (1956) provided further
342 2. ANALOCxS OF ENZYME REACTION COMPONENTS
Table 2-16
Inhibition of Pig Kidney d-Amino Acid Oxidase by Aromatic Acids "
Inhibitor
Ki Relative — AF oi binding
(mM) (kcal/mole)
2-Chloromethyl-5-hydroxy-l,4-pyrone
Kojate
TO-Toluate
Pjrrrole-2-carboxylate
Benzoate
Furan-2 -acrylate
p-Toluate
Nicotinate
Cinnamate
Furan-2-carboxylate
l,2-Pyrone-5-carboxylate (coumalate)
Indole-3-acetate
Hydrocinnamate
Phenylacetate
l,4-Pyrone-2,6-dicarboxylate (chelidonate)
o-Toluate
0.004
7.68
0.021
6.65
0.022
6.62
0.026
6.52
0.048
6.14
0.052
6.08
0.08
5.82
0.11
5.63
0.56
4.62
0.60
4.58
0.60
4.58
2.3
3.74
5.5
3.21
6.4
3.12
15
2.59
29
2. IS
"The substrate is DL-alanine and the pH 8.0-8.3. (Data from J. R. Klein, 1953, 1957.)
data on aliphatic and heterocyclic carboxylates (Table 2-17). Some of the
conclusions regarding relations between structure and inhibition derived
from these investigations will be summarized.
(1) A negatively charged anionic group is necessary for activity. This is
seen from the lack of inhibition by benzamide, nicotinamide, and cinnama-
mide. It is likely that the COO" group of the inhibitors reacts with the en-
zyme cationic site normally reacting with the amino acid C00~ group. A
SO3"" group can replace the COO" but is less effective.
(2) Klein has emphasized the importance of a positive charge at a distance
from the COO" group approximating the separation in amino acids. Most
of the potent inhibitors can be written in structures possessing such a
positive charge by virtue of resonance effects. Benzoate, for example,
resonates between the following structures:
- -J-
/ \ / / \
v>\ ^%
0
/ \
0
/ \
0
/
// ^
V /
/ \
/
_
)=c -
+( )
= c
\
\ /
'^ \
\ /
\
0
\ /
0
\ /
0
D-AMINO ACID OXIDASE
343
Table 2-17
Inhibition of Lamb Kidney d-Amino Acid Oxidase"
Inhibitor (3 mM)
Structure
% Inhibition
Crotonate
Butyrate
Dimethylacrylate
Isovalerate
Fumarate and maleate
CHj— CH=CH— COO'
CH3— CH2— CHj— COO'
H,C
^ \
C=CH— COO
H3C
H3C
CH— CH,— COO"
H3C
'OOC - CH= CH— COO'
99
0
70
Cinnamate
r y— CH=CH— COO'
100
Hydrocinnamate
CH,— CHo^COO
55*
Cinnamamide
(^ y — CH = CH-CONH2
Phenylacetate
CHo— COO
15
/>-Toluenesulfonate
H,C-
SO,
34
Sulfanilate
-h,nV/ V
so.
344
2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-17 (continued)
Inhibitor (3 mM)
Nicotinate
Structure
N-
// W
COO
Inhibition
65
Nicotinamide
N — V
CONH,
2-Furoate
O^ ^COO
W
100
Indol e - 2 - ca rbox yl ate
H
N^ COO'
100
Benzoate
/»-Methoxybenzoate
o - Methoxybenzoate
/J-Carboxybenzoate
o - Carboxybenzoate
Hz-Carboxyben^oate
Cyclohexanecarboxylate
€y
COO
( V-coo'
100
100
67
74
3
20
53*
Hydroquinone
Tribromophenol
/>-Aminophenol
Triiodophenol
Catechol
3, 5-Dihydroxyphenol
2, 3-Dihydroxyphenol
Resorcinol
73
40
30
19
19
5
0
0
" The substrate is o-alanine at 6.25 mM and the pH 8.3. The inhibitions marked
with an asterisk are at least partially due to contamination with the unsaturated
compounds. (Data from Frisell (7 (v/., 1956.)
D-AMINO ACID OXIDASE
345
and some of the other inhibitors may be written as:
,/0
Nicotinate
^O
2-Furancarboxylate
HOH,C
W A-""-""=V
HCH^=CH-CH=C:f -
O
O
Cinnamate
Crotonate
Indole - 2 - carboxylate
Such structures would be less possible or impossible for hydrocinnamate,
l,4-pyrone-2,6-dicarboxylate, cyclohexanecarboxylate, phenylacetate, and
some of the other weaker inhibitors. The orientation relative to the sub-
strate, according to Klein, would be represented as:
+
/C-CC
Substrate
Inhibitor
where X is carbon, oxygen, or nitrogen.
(3) Frisell and his co-workers, on the other hand, emphasized the impor-
tance of a double bond near the carboxylate group. Saturation of crotonate,
cinnamate, dimethylacrylate, and benzoate definitely reduces the inhibition.
They postulated that this double bond might correspond in position on the
enzyme to the double bond of the iminoketonic form of the dehydrogenated
product, and thus according to their theory the structural correspondence
would be:
Product
— C>
^c-c:
,o
Benzoate
-'^--c-cr"
Aliphatic carboxylate
(4) These two theories are not incompatible, since we have seen that
the presence of a positive charge generally depends on resonance, which in
turn requires double bonds and either conjugation or hyperconjugation.
346 2. ANALOGS OF ENZYME REACTION COMPONENTS
(5) The different potencies of the substituted benzoates would be explained
on the basis of the effects, such groups would have on resonance and the
magnitude of the positive charge, but in addition other factors must play
a role, for example the dipole moments of the ring X bonds and the possible
interactions of the substituent groups themselves. One might expect ortho
groups to decrease the binding and it is true that all are less inhibitory than
benzoate. Bartlett pointed out that the inhibition increases markedly with
the electronegativity of the halogens. It is very interesting to attempt to
interpret the results in Table 2-15 but without more knowledge about the
nature of the binding any hypotheses must be vague for the time being.
(6) It is unlikely that the degree of ionization of the carboxyl group is
important here because all of this work was done between pH 7.6 and 8.3
where a negligible fraction is undissociated. The p/iT^'s of all the substituted
benzoates tested run from 2.85 to 4.65. However, the series of phenols
studied by Frisell et al. (1956) may have to be considered in terms of the
ionization of the OH groups when relating structure to activity.
(7) The K-^ for some selected inhibitors were determined by Frisell
et al. (1956) and from these the relative binding energies may be estimated
(see accompanying tabulation). The 1.45 kcal/mole difference between cin-
Source of enzyme Inhibitor
A", Relative — AF oi binding
{mM) (kcal/mole)
Lamb kidney
Cinnamate
0.076
5.85
Crotonate
0.038
6.28
Pig kidney
Cinnamate
0.22
5.20
Benzoate
0.021
6.65
Indole-2-carb
axylate
0.0034
7.77
namate and benzoate (it is 1.52 kcal/mole from Klein's data) may be
attributed to a more satisfactory location of the positive charge in benzo-
ate. The lesser affinity of the enzyme for nicotinate compared with benzoate
(0.51 kcal/mole) could be due to the reduction in resonance brought about
by the asymmetry of the former. If the 1.94 kcal/mole difference between
pyrrole-2-carboxylate and furan-2-carboxylate is related to the different
amounts of positive charge on the nitrogen and oxygen atoms, this would
not be surprising since pyrrole should resonate more effectively. Finally,
if the positive charge theory is valid, the 3.02 kcal/mole difference between
benzoate and phenylacetate might indicate the amount of binding contri-
bution from the positive charge. The positive charge may, of course, be
stabilized somewhat by an enzyme anionic site with which it interacts.
D-AMINO ACID OXIDASE 347
Although the concept that these inhibitors bind to the enzyme and com-
pete with the substrate is very reasonable, some recent evidence may
indicate that the situation is a little more complex. The values of K^ for
benzoate should be the same for every substrate, according to the classic
treatment, but Klein (1956, 1960) has found that they are not, although
all the 1/v — 1/(S) plots are definitely competitive. The A;/s may vary as
much as almost 3-fold (see tabulation below). One possibility is that in
Substrate
Relative F,„
Km
[mM)
{mM)
Alanine
1.00
6.3
0.059
Proline
1.66
5.8
0.070
Phenylalanine
1.39
14.0
0.092
Valine
0.60
4.6
0.096
Isoleucine
0.79
4.1
0.104
Methionine
1.32
5.3
0.163
the aqueous extracts of pig kidney there are different oxidases for each
substrate but Klein prefers to assume that the inhibitors may react with
the ES complex to release the substrate:
ES + I ^ EI 4- S
The equilibrium constant K = (ES) (I)/(EI) (S) depends on the substrate
used and in the usual rate equation for competitive inhibition, {l)KJK^
would be replaced by {l)IK. Since K = KJK„ the determined values of K
should be inversely proportional to K,. They are not inversely proportional
to K„„ but is K„, = K, in this case? It may be noted that this mechanism is
not quite the same as uncoupling inhibition, since there an ESI complex is
formed and the substrate is not forced off. I must admit that I cannot
easily visualize how a competitive inhibitor can actively displace an enzyme-
bound substrate molecule.
Although Hellerman et al. (1946) reported the inhibition of D-amino acid
oxidase by benzoate to be independent of FAD concentration, there is
more recent evidence that certain aromatic carboxylates and phenols not
only can compete with substrate, but can also either compete with FAD
or complex directly with FAD (Yagi et al., 1957, 1959, 1960). The constants
for each of these reactions are given for a few of these inhibitors in the fol-
lowing tabulation. Only the carboxylates compete with substrate, while
the phenols act by the other two mechanisms; the substances with both
COO- and OH groups react in all three ways, although competition with
the substrate is the most important. Yagi and his group have recently
348 2. ANALOGS OF ENZYME REACTION COMPONENTS
K^ (mi/)
Inhibitor
Competition
Competition
Complex
with substrate
with FAD
with FAD
0.0145
0.35
1.05
0.65
6.35
15
75
0.305
7.25
—
—
0.85
—
—
9.15
—
—
56.5
32
—
28
110
—
5.0
0.5
—
3.9
0.02
—
2.3
0.04
Benzoate
Salicylate
^-Aminosalicylate
p-Arainobenzoate
p-Nitrobenzoate
Aniline
Phenol
m-Aminophenol
p-Nitrophenol
2,4-Dinitrophenol
2,6-Dinitrophenol
2,4,6-Trinitrophenol — 0.64 0.012
crystallized the complex of apoenzyme, FAD, and benzoate, and found
equimolar amounts of each component present.
It would be interesting to have more data on the effects of these inhibitors
on L-amino acid oxidases. Benzoate inhibits the L-amino acid oxidases
from rat kidney (Blanchard et al., 1944) and snake venom (Zeller and Maritz,
1945), but does not inhibit the enzyme from Neurospora even at 10 niM
(Burton, 1951 b).
Benzoate on Other Enzymes and Metabolism
The endogenous ammonia formation and respiration of rat kidney slices
are inhibited 52% and 66%, respectively, by 25 mM benzoate (Herner,
1944). One might assume that the former might be attributed to inhibition
of amino acid oxidases, but this is unlikely because benzamide, phenyl-
acetate, and /?-phenylpropionate are even more potent inhibitors than
benzoate. The deamination of various l- and D-amino acids in kidney is,
however, strongly inhibited by benzoate, at least in part competitively.
The inhibition of respiration by benzoate was first observed by Grif-
fith (1937) in slices and minces of various tissues in the presence of glucose,
but no analysis of the site of action has been published, although in con-
nection with recent studies on the salicylates some effects on mitochondria
have been investigated. Benzoate inhibits the oxidations of citrate, a-
ketoglutarate, and succinate in rat kidney homogenates but is invariably
less active than salicylate (E. H. Kaplan et al., 1954); and neither the Og
uptake nor the P : 0 ratio is affected markedly in rat liver mitochondria,
although salicylate uncouples strongly (Brody, 1956). Endogenous phosphor-
D-AMINO ACID OXIDASE 349
ylation in liver mitocliondria is inhibited 55% by 10 mM benzoate (Wein-
bacli, 1961). One can conclude from this limited material that benzoate is
certainly a weak inhibitor of cycle oxidations and phosphorylations. Bosund
(1959, 1960 a, b) has investigated the effects of benzoate on the metabolism
of glucose and pyruvate in Proteus vulgaris in attempting to elucidate
the mechanisms for the bacteriostatic activity. There is no interference
with glucose metabolism to the acetate level, and acetate was found to
accumulate. The respiratory quotient is increased from 1.24 to 1.82 by
benzoate during the oxidation of pyruvate and the Oa/pyruvate ratio is
decreased. The oxidation of pyruvate in yeast is quite strongly inhibited
by benzoate (50% at 0.4 mM), especially at low pH's where penetration is
better, but acetate oxidation is less sensitive. It is quite possible that these
inhibitions play a role in the suppression of growth, which for yeast requires
5 mM benzoate at pH 5.1 and 60 mM at pH 6. It is clear that much more
work must be done before the mechanisms of respiratory inhibition are
understood.
Benzoate can also interfere in lipid metabolism, as demonstrated many
years ago by Jowett and Quastel (1935 a, b), but the mechanism is still
unknown. In liver slices it was claimed that benzoate at around 0.5-2
mM inhibits specifically the oxidation of fatty acids, and the oxidation of
crotonate 63% at 1 mM. There is progressively less effect on the higher
fatty acids, little inhibition of decanoate being observed. It is possible
that the benzoate ring simulates the aliphatic chains of butyrate or croton-
ate enabling it to compete with these substrates for some enzyme; it would
be interesting to know if benzoate can participate in any of these reactions
(e. g., if benzoyl-CoA is formed) and deplete the systems of some cofactor.
Benzoate is a weak inhibitor of tyrosinase (Ludwig and Nelson, 1939),
chymotrypsin (Foster and Niemann, 1955 b), p-aminobenzoate acetylation
(Koivusalo and Luukkainen, 1959), and NADPH dehydrogenase (Kasa-
maki et al., 1963); it does not effect shikimate dehydrogenase (Balinsky and
Davies, 1961 b) or D-glutamate oxidase (Mizushima and Izaki, 1958) at
1 mM, or a-ketoisocaproate decarboxylase at 4 mM (Sasaki, 1962).
Kojic Acid
The potent inhibition of D-amino acid oxidase by kojic acid is interesting
in light of the central nervous system effects observed in dogs, rabbits, and
rats, namely, ataxia, excitement, and convulsions (Friedemann, 1934).
Kojic acid was first isolated by Saito in 1907 from Aspergillus oryzae and
has since been found in many species of Aspergillus. It is a weak antibiotic,
inhibiting growth of most bacteria at 2-15 mM, but is particularly active
against Leptospira, complete growth inhibition being observed at 0.007 m.N
(Morton et al., 1945). Toxic effects are produced in dogs by 150 mg/kg and
in mice by 250 mg/kg when injected parenterally; the LD50 for mice is
350 2. ANALOGS OF ENZYME EEACTION COMPONENTS
1.5-2.0 g/kg. Leucocytic activity and phagocytosis are not affected by 18
raM kojic acid. A biochemical study was undertaken by Klein and Olsen
(1947), who found that the respiration of muscle and heart mince is resistant
to kojic acid, whereas 10 mM suppresses the respiration of liver 40%,
kidney 20%, and brain 15%. The convulsant dose corresponds to a tissue
concentration around 4-50 mM. The oxidation of both l- and D-amino
acids in liver homogenates is quite strongly inhibited in a competitive
fashion: for example, 50% inhibition is given by 0.04 niM for L-methionine
and by 0.12 mM for l- and D-phenylalanine. Xanthine oxidation is also
inhibited (50% at 0.7 mM). It was suggested that kojic acid may be an
inhibitor of flavin enzymes, and it is possible that some direct complexing
with FAD may occur. Nevertheless, FAD does not influence the inhibition
of D-amino acid oxidase. There is no inhibition of the oxidation of succinate,
tyramine, L-proline, choline, or urate at 5 mM kojic acid. Although the
metabolic effects are interesting, it is impossible to correlate any of these
inhibitions with either the central effects in animals or the bacteriostatic
activity.
ANALOG INHIBITION OF THE METABOLISM
OF VARIOUS AMINO ACIDS
Many reports indicate the influence of analogs on various enzymes con-
cerned with amino acid metabolism, but in most cases insufficient work
has been done to draw clear conclusions about the mechanism of the bind-
ing to the enzymes. It will suffice to present some of the results in Table
2-18. Probably many of these inhibitions are competitive but they have
been so indicated only when graphical analysis has shown this to be true.
A few of these inhibitions may be significant in feed-back control or in the
general regulation of amino acid metabolism.
Generally speaking, there are certain types of amino acid analog that
have proved to be effective inhibitors. Mcllwain (1941) pointed out that
aminosulfonate analogs of amino acids are frequently bacteriostatic and
that this inhibition is reduced by adding the normal amino acids. If staphy-
lococci are trained to be independent of exogenous amino acids, the a-
aminosulfonates no longer inhibit. However, the exact sites of action
of these analogs have not been determined. Umbreit (1955 b) has discussed
the general inhibitory properties of the a-methylamino acids and pointed
out that the inhibitions are often competitive only for a short interval
if both substrate and inhibitor are present, whereas they are noncompetitive
if the inhibitor is added first. This is a matter of terminology; the inhibitions
are probably competitive but appear to be noncompetitive because of the
very high affinities of some analogs for the enzymes (an inhibition can be
competitive even though irreversible but the substrate must be given an
ANALOG INHIBITION OF THE METABOLISM OF VARIOUS AMINO ACIDS 351
opportunity to compete). A third group of interesting analogs is the halogen-
substituted amino acids, which are often potent inhibitors of protein syn-
thesis and growth. The fluoroamino acids are particularly active. The
m-, 0-, and p-fluorophenylalanines all inhibit the formation of the adaptive
maltase in yeast, the last being the most potent (Halvorson and Spiegelman,
1952). On the other hand the p-chloro- and js-bromophenylalanines do not
inhibit. The incorporation of phenylalanine and other amino acids into
ascites cell proteins is competitively inhibited by o-fluorophenylalanine;
this is in part due to a depression of transport into the cells and in part due
to block of some unknown steps in the incorporation (since labeled phenyl-
alanine accumulates in cells) (Rabinovitz et al., 1954). There is also an inhi-
bition of protein synthesis in rat liver in vitro by the fluorophenylalanines,
this leading to a net breakdown of tissue proteins since the constant bal-
ance of synthesis and degradation is disturbed (Steinberg and Vaughan,
1956). a-Amino-/?-chlorobutjTate is an analog of valine and inhibits valine
incorporation into rabbit reticulocyte proteins, including hemoglobin; it
was suggested that the analog enters a precursor protein which is unable
to assume the proper configuration of hemoglobin and thus there is accu-
mulation of protein intermediates (Rabinovitz and McGrath, 1959). As
pointed out previously, some of these analogs are incorporated into cell
proteins. p-Fluorophenylalanine-C^* is incorporated into the proteins of
muscle, blood, and liver when fed to rabbits, this being a replacement of
phenylalanine (Westhead and Boyer, 1961). The replacement of phenyl-
alanine in aldolase is 25% and in 3-phosphoglyceraldehyde dehydrogenase
16%, and in each case the enzyme activities are normal. Despite this ap-
preciable incorporation, the rabbits suffer no obvious biochemical or physi-
ological disturbances, so that mammals may well differ from microorgan-
isms in the response to this analog.
Feedback inhibition in the biosynthetic pathways of amino acids is an
important aspect of regulation but we can touch only briefly on this prob-
lem. An interesting example of this has been studied in connection with
the synthesis of histidine, since the enzyme inhibited is the first in the
pathway and catalyzes a reaction not involving substrates structurally
similar to histidine (Martin, 1963). This enzyme is phosphoribosyl-ATP
pyrophosphorylase and the reaction catalyzed is:
5'-P-ribosyI-PP + ATP :^ N-l-(5'-P-ribosyl)-ATP + PP
Histidine is a surprisingly potent and specific inhibitor with K, = 0.1 mM.
Related compounds inhibit weakly or not at all; 2-methylhistidine, for
example, exhibits weak inhibition with K^ = 2.4 mM. The inhibition varies
with the pH but maximal inhibition is exerted at physiological pH. HgClg
at 0.03 mM does not inhibit the enzyme but blocks almost completely the
inhibition by histidine. This coupled with the fact that the inhibition by
352
2. ANALOGS OF ENZYME REACTION COMPONENTS
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ANALOG INHIBITION OF THE METABOLISM OF VARIOUS AMINO ACIDS 353
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ANALOG INHIBITION OF THE METABOLISM OF VARIOUS AMINO ACIDS 355
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ANALOG INHIBITION OF THE METABOLISM OF VARIOUS AMINO ACIDS 357
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358 2. ANALOGS OF ENZYME REACTION COMPONENTS
histidine is noncompetitive indicates that the histidine site is different
from the catalytically active site. ATP and 5'-P-ribosyl-PP protect the
enzyme against inactivation by trypsin, and histidine eliminates this pro-
tection, so it is possible that histidine modifies the enzyme configuration
by combining with the feedback site.
Aminooxyacetate
An interesting analog studied recently in connection with the physiolog-
ically important y-aminobutyrate (GABA) is aminooxyacetate (+H3N —
0 — CH2COO"). This substances inhibits the GABA: a-ketoglutarate trans-
aminase of E. coli very potently (40% at 0.0033 mM and 100% at 0.33
mM when both substrates are 27 mM), so it was tested on a similar enzyme
from brain and found to inhibit as strongly (Wallach, 1960). Aminooxyace-
tate in doses of 6-50 mg/kg elevates the brain GABA in several species as
much as 4- to 5-fold, peak levels being reached about 6 hr after administra-
tion and high levels remaining up to 24 hr (Wallach, 1961 a; Schumann et al.,
1962). Reinvestigation of the transaminase inhibition led to a K^ of 0.0075
mM (Z,„ is 9.66 mM for a-ketoglutarate and 27.6 mM for GABA); the
inhibition is competitive with respect to both substrates. The alanine:
«-ketoglutarate transaminase from rat liver and heart is also potently
inhibited, around 60-80% by 0.0001 mM aminooxyacetate at pH 6.8
(Hopper and Segal, 1964). Indeed, this transaminase seems to be more
sensitive than either the GABA:a-ketoglutarate or aspartate:or-ketoglutar-
ate transaminase.
Inasmuch as GABA has been implicated in certain convulsive disorders
(e. g., GABA formation in epileptic brains is apparently depressed), amino-
oxyacetate was administered to animals made convulsive with thiosemi-
carbazide and methionine sulfoximine (DaVanzo et al., 1961). Anticonvul-
sant activity was observed but there is some doubt if this is correlated with
the brain GABA levels since the time relations are not correct. The effects
of aminooxyacetate on central nervous system function are complex.
There is first a progressive depression and muscular relaxation with loss of
certain reflexes, but at high doses tonic-clonic convulsions occur (DaVanzo
et al., 1964 a). Pyridoxal-P antagonizes these convulsions and it was postu-
lated that oxime formation occurs between aminooxyacetate and pyri-
doxal-P. Pyridoxal-P does not reverse the transaminase inhibition in the
brain (Wallach, 1961 b). W^allach also suggested that a depletion of suc-
cinic semialdehyde, which arises from GABA by transamination, might
also play a role in the convulsant action. Since a pyridoxal deficiency is pro-
duced in rats by the administration of aminooxyacetate, DaVanzo et al.
(1964 b) postulated another possible mechanism of action, namely, the
inhibition of pyridoxal kinase, inasmuch as McCormick and Snell (1961)
had shown this enzyme to be rather potently inhibited by the condensation
ANALOG INHIBITION OF THE METABOLISM OF VARIOUS AMINO ACIDS 359
product of aminooxyacetate and pyridoxal. It may be that aminooxyacetate
should be classed as a carbonyl reagent which reacts with pyridoxal-P,
rather than strictly as an amino acid analog, but it is sometimes difficult
to distinguish these actions. Nevertheless, the effects of aminooxyacetate
on GABA levels in the tissues will undoubtedly make it a useful compound
for the study of the physiological role of GABA.
Cycloserine
D-Cycloserine (orientomycin, Oxamycin) is a tuberculostatic antibiotic
isolated from Streptomyces which can be considered as a cyclic form of
aminooxyalanine or 0-aminoserine. It is written in the zwitterion state be-
I I
H nh;
Cycloserine
cause there is some evidence that this is the inhibitory form (Neuhaus and
Lynch, 1964). Its actions are similar to aminooxyacetate in many respects
but differ occasionally in interesting ways. D-Cycloserine inhibits the growth
of many bacteria and this is often well antagonized by D-alanine, and
competitively inhibits the incorporation of D-alanine into a uridine nucleo-
tide necessary for the synthesis of cell wall material. The growth of myco-
bacteria is 50% suppressed, for example, by D-cycloserine at 0.03-0.045 mM
and this is reversed by D-alanine but not by L-alanine (Zygmunt, 1963).
L-Cycloserine is also inhibitory to certain bacteria and this is antagonized
by L-alanine. It is interesting that *S. aureus can develop a 50-fold resistance
to D-cycloserine, no cross-resistance with other antibiotics being observed
(Howe et al., 1964). In animals it produces sedation, lethargy, muscular
relaxation, ataxia, an accentuated startle response, and, above all, epi-
leptic convulsions (Dann and Carter, 1964; Holtz and Palm, 1964). In these
respects it at least superficially acts like aminooxyacetate and, furthermore,
these effects are antagonized by pyridoxal.
D-Cycloserine inhibits certain enzymes dependent on pyridoxal-P, such
as the transaminases and glutamate decarboxylase, and it has been postu-
lated that it simply reacts with pyridoxal-P to form a substituted oxime.
However, this is not the usual reaction in which a Schiff base is produced,
but involves an opening of the cycloserine ring. The inhibition of GABA:
or-ketoglutarate transaminase is initially competitive with respect to GABA
{K^ is 0.25 TCiM for the enzyme from cat brain and around 0.6 mM for the
enzyme from E. coli), but a secondary progressive irreversible inhibition
also occurs (Dann and Carter, 1964). This may be related to the hypothesis
of Khomutov et al. (1961) that the decyclicized cycloserine forms an oxime
360 2. ANALOGS OF ENZYME KE ACTION COMPONENTS
bond with enzyme-bound pyridoxal-P, and also an acyl bond with a cat-
ionic group at the active site, these two groups thus being connected by a
bridge would prevent access to the site. The L-asparagine:a-ketoglutarate
I
HC = NH— 0— CHa— CH— CO
I I
Pyr X
Apoenzyme
transaminase is inhibited by L-cycloserine with respect to L-asparagine
{K^ = 0.0001 mM) but D-cycloserine is a much weaker inhibitor {K^ =
1 mM) (Braunstein et al., 1962). L-Cycloserine likewise competitively
inhibits the L-alanine: a-ketoglutarate transaminase {K^ = 0.008 mM).
This taken with previous work indicates that the cycloserines inhibit
specifically those enzyme attacking substrates of the same optical isomerism.
Another possible site of action of D-cycloserine in bacteria would be the
D-alanyl-D-alanine synthetase, a block of which would prevent the incor-
poration of D-alanine into cell wall material. Alanylalanine is often able to
reverse the cycloserine inhibition of bacterial growth, sometimes more ef-
fectively than alanine. One example of this is the inhibition of the prolif-
eration of agents of the psittacosis group in chick embryo yolk sac (Moulder
et al., 1963). Chick embryos infected with the mouse pneumonitis organism,
for example, are well protected by D-cycloserine at 0.004-0.008 mM, and, of
all the possible reversors tested, only alanylalanine is effective. The d-
alanyl-D-alanine synthetase of Streptococcus fecalis is inhibited competi-
tively with respect to D-alanine {K^ = 0.022 mM), and Neuhaus and
Lynck (1964) felt that this enzyme may well be the major site of inhibition
in certain bacteria. It is unfortunate that cycloserine and aminooxyacetate
have not been accurately compared in any study.
DIAMINE OXIDASE (HISTAMINASE)
Enzymes in this group exhibit different degrees of substrate specificity
depending on the source, but most oxidatively deaminate diamines of the
type +H3N — (CH2),i — NH3+ (with maximal rates when n is around 5) and
histamine, in all cases primary amines being attacked. The usual substrate
in most studies has been cadaverine {n = 5).
The diamines have been written as cations because the p^^'s are usually
between 8.5 and 10.5. The amidines and guanidines exist almost entirely
as cations at physiological pH since the p^^'s for these groups are around
13 to 14. The structures have been written in a rather unconventional way
DIAMINE OXIDASE (hISTAMINASE)
361
H3N— (CH2)— NHa""
Cadaverine
Putrescine
H3N— (CH2 )3— ^fH— (C H2 )4— NH — (CH2 )3— NH^
*2M
Spermine
CH3— (CH3)^C^ +
Monoamidines
HjN^ , , /NH2
HzN'^ NH2
Diamidines
/=\ /NH3
Dibenzamidines
H,N— C
^NH— CH3
Methylguanidine
H,N-C
'NH^
-NH— NH2
Aminoguanidine
H^N^ /NH2
+ C— NH— (CH,);,— NH— C +
Diguanidines
+ /NH2
H3N— (CHs).— NH-C +
Agmatine
H^N^ /NH2
+ C— S — (CHs)^— S-C- +
HgN^ NH2
Diisothioureas
H2N /NH2
+ C-NH — (CH2)4— NH-C +
HgN^ NH2
Arcaine
to indicate the equivalence of the C — N bonds and the states of the amino
groups, since there is resonance between forms such as the following for
the guanidinium ion:
+ /NH2
H,N— C
+
NH,
^NH,
H,N-C,
,NH,
NH,
+ ^
Alkyl substitution does not reduce the resonance appreciably.
Aliphatic monoamines, such as amylamine, are not substrates nor are
362
2. ANALOGS OF ENZYME REACTION COMPONENTS
they readily bound to the enzyme since they inhibit weakly (Zeller, 1940).
The short aliphatic diamines are very poor substrates but inhibit quite
well; thus both ethylenediamine and trimethylenediamine inhibit the
oxidation of cadaverine by pig kidney diamine oxidase (Zeller, 1938).
If the amino groups of these inhibitors react with the same anionic enzyme
sites as does cadaverine, these anionic groups must be fairly close so that
cadaverine would have to assume a very bowed configuration. On the other
NH3+
\ ^
CH,
\
CH,
/ '
NH+
H.
C
HgC
CH2
/
\
HgC
CH2
+ \
/+
H3N
NH3'^
-
-
H,C-
^H,N
-CH,
NH,
(a)
(b)
(c)
hand, the anionic groups may be separated by a distance corresponding
to the amino groups in cadaverine and the inhibitors react with only one
of the anionic sites ,the other amino group interacting with some anionic
site outside the active center (as in (c)).
Certain guanidine derivatives are more potent inhibitors. Guanidine
itself is rather weak, inhibiting cadaverine (2 mM) oxidation 42% at 10
mM, but methylguanidine inhibits 63% at 1 mM (Zeller, 1938). Although
Zeller noted that in the pig kidney preparation methylguanidine did not
inhibit histamine oxidation very well, Waton (1956) found marked inhibi-
tion of histaminase activity in cat kidney, 0.01 mM inhibiting 42% and
0.1 mM 75%. The oxidations of putrescine and agmatine are both well
inhibited by methylguanidine (Zeller, 1940). An even more potent inhibitor,
however, is aminoguanidine. Apparently the diamine oxidases differ in
sensitivity to aminoguanidine; 50% inhibition is given by 0.00005 mM for
the enzyme from pig kidney (Schuler, 1952), by 0.001 mM for the enzyme
from cat kidney (Waton, 1956), by 0.01 mM for the enzyme from rabbit
liver (Kobayashi, 1957), and the enzyme from mouse liver is not inhibited
even by 0.1 mM. The nature of the inhibition is not clear, inasmuch as
aminoguanidine is also a derivative of hydrazine and might act by attacking
carbonyl groups: hydrazine and semicarbazide are, indeed, potent inhibitors
of diamine oxidase (Schuler, 1952; Waton, 1956). A further complication is
that aminoguanidine hydrolyzes to form semicarbazide and eventually
hydrazine. It behaves chemically more like a hydrazine than a guanidine,
and reacts with carbonyl groups without being hydrolyzed (Lieber and
Smith, 1939). It is also possible that the NHNHg group simulates the
CH2NH2 substrate group, as in the monoamine oxidase inhibitors, and forms
a tight bond to the enzyme. In vivo inhibition of diamine oxidase by amino-
DIAMINE OXIDASE (HISTAMINASE)
363
guanidine was demonstrated by Schayer et al., (1954) by injecting labeled
cadaverine into mice and determining the C^^Og respired (Fig. 2-7). It is
much more effective than isoniazid or agmatine. The metabolism of hista-
mine should also be blocked by aminoguanidine. This was shown in three
psychiatric patients by administering labeled histamine and finding that
100
-1.5 -0.5
LOG INH DOSE (^g/G)
+ 0.5
+ 1.5
+ 2.5
+ 3.5
Fig. 2-7. Effects of diamine oxidase inhibitors on the oxidation of cadaverine in
mice, as determined by the formation of C'^Oj from labeled cadaverine. (Data from
Schayer et al., 1954.)
urinary imidazoleacetate-C^^ is reduced by aminoguanidine at 0.1-1 mg/kg
(Lindell et al., 1960). A larger fraction of the histamine is excreted as
methylhistamine, demonstrating a diversion of metabolic pathways by
this inhibitor.
Diamine oxidase is not strongly inhibited by monoamidines, but diami-
dines, diguanidines, and diisothioureas of the proper chain lengths are
quite potent inhibitors (Blaschko et al., 1951). The data for these series
are summarized in Fig. 2-8. The correlation between chain length and
inhibition is not nearly so clear as for monoamine oxidase (Blaschko and
Duthie, 1945; Blaschko and Himms, 1955), and in some cases, as the dia-
midines, there is surprisingly little variation of inhibition with chain length.
For diamidines of n = 10-16, one wonders if the binding is actually to
364
2. ANALOGS OF ENZYME KEACTION COMPONENTS
Fig. 2-8. Inhibition of pig kidney diamine oxidase with cadav-
erine (5 niM) as the substrate and all the inhibitors at 1 mM.
(From Blaschko et ah, 1951.)
NH,
(A) Monamidines
H3C — (CHa)^ C
NH
(B) Diamidines
H,N
NH,
(C) Dibenzamidines
(D) Diguanidines
H2N
HN
H,N
0-(CH2)„-0— (
NH,
NH,
W
NH
C-NH-(CH2)— HN-C^
HN ^NH
(E) Diisothioureas
H,N
NH,
C-S- CH,L— S— C
// \\
HN NH
CARBOXYPEPTIDASE, AMINOPEPTIDASES, DIPEPTIDASES 365
the substrate site entirely, or possibly to two substrate sites, or even to
anionic groups outside the substrate site. The enzyme may have binding
sites for the imidazole ring of histamine since imidazole inhibits 11%,
imidazolelactate 20%, and histidine 4% at 6.7 mM when cadaverine is
3.3 mM (Zeller, 1941). Urate also competitively inhibits the oxidation of
histamine, but rather weakly. An excellent review of the structure-action
relationships among the amidine derivatives is by Fastier (1962).
CARBOXYPEPTIDASE, AMINOPEPTIDASES,
AND DIPEPTIDASES
Certain aspects of the inhibition of carboxypeptidase by substrate
analogs were discussed in Volume I (page 292) to illustrate how certain
interaction contributions could be estimated. We shall now attempt to
visualize more clearly the orientation of these analogs on the enzyme sur-
face. The data indicate that a three-point attachment of the substrate
is necessary for catalysis. The enzyme sites may be indicated as follows
(see Fig. 2-9 for hypothetical orientation of substrate): (A) the peptidatic
site contains the mechanism of the electron displacement necessary for hy-
drolysis and is probably positively charged, (B) the cationic site is a positively
charged group that interacts electrostatically with the C00~ group, and
(C) the electrokinetic site is perhaps a lipophilic region capable of reacting
with alkyl or phenyl groups by dispersion forces. It is easy to see why
D-substrates are not reacted since the peptide bonds would not be able to
approach the peptidatic site. There is also an enzyme region near the projec-
tion direction of the fourth asymmetric carbon bond that sterically prevents
attachment of groups larger than an amino group, and thus the D-isomers
usually do not bind and are not inhibitors. Only two-point attachment is
necessary for inhibitors, and most that have been studied bind at the cationic
and electrokinetic sites.
The relative binding energies for inhibitors in Table 1-6-26 were calculated
from the data of Smith etal. (1951). Earlier studies by Elkins-Kaufman and
Neurath (1949) provide additional information on the competitive inhibitors
in the accompanying tabulation. It is interesting that D-phenylalanine is
Inhibitor
(mM)
Relative — AF oi binding
(kcal/mole)
^-Phenylpropionate
0.062
5.96
Phenylacetate
0.39
4.83
y - Phenylbutyrate
1.13
4.17
D - Phenylalanine
2
3.82
2?-Nitrophenylacetate
2.5
3.68
366
2. ANALOGS OF ENZYME REACTION COMPONENTS
A.
Acyl-L-phenyl-
alanine (substrate)
(3"'-^l
L- Phenylalanine
/^ X NH3 Vc
r hV
Benzoate
\^ y CHj
D- Phenylalanine
Ql
Phenylacetate
Propionate
Q-
3 - Pheny Ipr opionat e
(hydrocinnamate)
Valerate
y- Pheny Ibuty rate 3-Indoleacetate
Fig. 2-9. Possible orientations of substrate and inhibitors at the active center of
carboxypeptidase. (A) is peptidatic site, (B) is cationic site, and (C) is electrokinetic
site. The molecular configurations are only approximate.
CARBOXYPEPTIDASE, AMINOPEPTIDASES, DIPEPTIDASES 367
a better inhibitor than L-phenylalanine under the usual conditions, since
the NH3+ group of the latter is repelled by the positively charged pep-
tidatic site while in the former it does not encounter serious steric in-
terference (Fig. 2-9). However, /5-phenylpropionate (hydrocinnamate) is
bound more tightly than D-phenylalanine by 2.14 kcal/mole, suggesting
that some steric repulsion of the latter analog occurs. iV- Substitution in-
creases the repulsion markedly and inhibitory activity is lost. A comparison
of the possible orientations of some inhibitors in Fig. 2-9 with the relative
binding energies may give some idea of the structural requirements for
potent inhibition. It may be noted that benzylmalonate is an effective
inhibitor but is somewhat less well bound than /5-phenylpropionate; this is
surprising because it might be thought that additional energy would be
contributed by interaction of one of the C00~ groups with the peptidatic
site. Neither cis- nor <rans-cinnamate inhibits and it was suggested that the
double bond restricts the orientation of the ring so that adequate binding
cannot occur. The linearity of these molecules may also be a factor, since
the active site is probably not flat as is implied by the two-dimensional
representations in the figure.
Competitive inhibition by the following analogs has been demonstrated
more recently: 3-indolepropionate, e-aminocaproate, (5-amino-w-valerate
(Folk, 1956; Greenbaum and Sherman, 1962), y-aminobutyrate, S-guani-
dinovalerate, argininate (Folk and Gladner, 1958), iV-acetyl-L-tyrosine,
D-leucinyl-L-tyrosine, glycyl-L-tyrosine, and other dipeptides (Yanari and
Mitz, 1957). The inhibition apparently sometimes depends on the substrate
used; for example, 3-indolepropionate inhibits the hydrolysis of carbo-
benzoxyglycyl-L-phenylalanine but not the hydrolysis of a-iV-benzoyl-
glycyl-L-lysine, where s-aminocaproate exhibits just the opposite behavior.
Relatively little work has been done on analog inhibition of dipepti-
dases and aminopeptidases, and it will suffice to mention a few isolated
observations. Yeast dipeptidase is inhibited by various amino acids; for
Concentration for
Amino acid
50% inhibition
(mM)
L-Leucine
1.5
L-Isoleucine
1.8
L-Tryptophan
6.0
L-Histidine
8.0
L-Leucinamide
9.0
L-Arginine
18
DL- Valine
22
DL-Phenylalanine
22
DL-Serine
>100
368 2. ANALOGS OF ENZYME REACTION COMPONENTS
example, L-leucine, D-alanine, and glycine inhibit the hydrolysis of alanyl-
glycine and glycylglycine (Grassmann et al., 1935). This has been investi-
gated more quantitatively by Nishi (1960) and his results are summarized
in the accompanying tabulation (substrate is glycylglycine at 50 mM).
These inhibitions are competitive with respect to substrate and uncompe-
titive with respect to Co++. Some interesting inhibitions of pig kidney
leucine aminopeptidase have been reported by Hill and Smith (1957).
The hydrolysis of substrates of the type R— CH(NH3+)— CONH— R'
depends on a three-point attachment of the R group, the NH3+ group,
and the amide N. The R groups interact by van der Waals' forces; further
energy is contributed from the hydrogen bonding of water molecules dis-
placed from the hydrophobic surfaces. The inhibitions given in the follow-
ing tabulation are for L-leucinamide as substrate at 50 mM and at pH 8.50-
Inhibitor
Concentration
{mM)
% Inhibition
L-Leucine
50
45
100
61
L-Leucinol
50
38
100
48
Isocaproamide
25
41
50
56
Isocaproate
100
47
200
65
w-Hexylamine
100
0
a-Ketoisocaproamide
25
21
L-a-Hydroxyisocaproamide
50
68
8.65. Every good inhibitor contains an R group that should give nearly
optimal interaction with the electrokinetic site of the enzyme; in addition,
at least one group that will react with the bound Mn++ is present.
CHYMOTRYPSIN AND OTHER PROTEOLYTIC ENZYMES
Chymotrypsin hydrolyzes various amides and esters of the general type:
NH— R2
Rj — CH2 CH
bo— R3
where R^ represents the side chains of amino acids (phenylalanine, tyrosine,
and tryptophan most commonly used), Rg is an acyl group (acetyl, benzoyl,
CHYMOTRYPSIN AND OTHER PROTEOLYTIC ENZYMES 369
or nicotinyl), and Eg is a group forming either an amide or ester bond.
A typical substrate is benzoyl-L-tyrosinamide:
NH— CO
KO—(. />— CH,— CH
\\ // Vo-NH
o
In addition, the NH — R2 chain may be replaced by H, CI, or OH groups.
Only the derivatives of L-amino acids are hydrolyzed. The R^ and Rg
groups are important in binding to the enzyme and thus, with the esteratic
(paptidatic) site, one may again visualize a three-point attachment. Analogs
either devoid of susceptible amide or ester bonds, or having in their
place bonds resistent to hydrolysis, are often inhibitory. The R^ group is the
most important for binding, as is shown by the strong inhibitory activity
of /5-phenylpropionate (hydrocinnamate) (Kaufman and Neurath, 1949).
The necessity for at least one aromatic ring in one of the side chains was
pointed out by Neurath and Gladner (1951). Their data on the /^-substituted
propionates indicate the ring groups to have the following order of inhibitory
activity:
Indole > napthyl > phenyl > 2,4-dinitrophenyl > cyclohexyl
The distance between the COO" group and the Rj group is also of impor-
tance. The inhibitions are summarized in Table 2-19. The weaker binding
of cyclohexyl derivatives compared to phenyl compounds (0.5-1 kcal/mole)
could be explained by either the smaller polarizability of the cyclohexyl
ring or the inability of the cyclohexyl ring to approach the enzyme surface
as close as the phenyl ring. The strong binding of the indole compounds
was explained on the basis of the enhancement of hydrogen bonding by the
ring N. In fact, Neurath and Gladner interpreted the inhibitions by most
of the analogs in terms of hydrogen bonding. Even the C00~ may not
interact electrostatically with an enzyme cationic group since the binding
energies are quite low; indeed, it may serve as a hydrogen acceptor. The
equivalent bindings of
. /y — CH2CH2— COO and (^ /)— O-CH^CH,— OH
l3-Phenylpropionate 2-Phenoxyethanol
would be difficult to explain otherwise; however, the latter compound can
act as a hydrogen donor in forming a hydrogen bond. These two com-
pounds have essentially the same molecular dimensions but the electronic
configurations of the terminal groups differ.
370 2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-19
Competitive Inhibition of a-CnYMOXRYPSiN by Analogs "
Inhibitor
Apparent iC, Relative — AF oi binding
(mM) (kcal/mole)
3-Indolepropionate 2.5 3.57
3-Indolebutyrate 3.6 3.35
a-Naphthylpropionate 4.0 3.28
2,4-Dinitro-^-phenylpropionate 5.3 3.08
^-Phenylpropionate 5.5 3.07
2-Phenoxyethanol 5.8 3.05
y-Phenylbutyrate 14 2.53
Cyclohexylpropionate 30 2.08
Cyclohexylbutyrate 35 1.99
Benzoate 42 1.88
Phenylacetate 42 1 . 88
a-Naphthylmethylmalonate 55 1 . 72
a-Benzylmalondiamide 78.8 1.51
Cyclohexylacetate 86 1.46
" The substrate is acetyl-L-tyrosinamide (K^ = 27 mM). Experiments at pH 7.8
and 25°. (Data from Neurath and Gladner, 1951.)
For the past several years Niemann and his associates have conducted
excellent quantitative investigations of the inhibition of chymotrypsin
by a variety of analogs and their results, some of which are presented in
Table 2-20, provide a basis for the interpretation of the binding mechanisms.
Although final conclusions must await completion of their work, some tenta-
tive ideas may be expressed.
(A) Despite the fact that the derivatives of L-amino acids are hydro-
lyzed by chymotrypsin, the D-isomers of several inhibitors are bound on an
average of 0.45 kcal/mole more tightly than the corresponding L-isomers
(see also page 271). Although three-point attachment may be important for
substrate binding, it is evident from this difference and other data that
it is not for inhibitor binding.
{B) Comparing the derivatives of the three amino acids, it is seen that
the phenylalanine and tyrosine analogs are equally bound, whereas the
tryptophan analogs are bound some 1.19 kcal/mole more tightly, and this is
probably to be attributed to the greater affinity of the enzyme for the
CHYMOTRYPSIN AND OTHER PROTEOLYTIC ENZYMES
371
Table 2-20
Competitive Inhibition of q-Chymoteypsin by Analogs "
Inhibitor
Apparent K^ Relative — AF oi binding
(mil/) (kcal/mole)
Tryptophan series
L-Tryptophanamide
D-Tryptophanamide
Acetyl-D-tryptophanamide
Trifluoroacetyl-D-tryptophanamide
Acetyl-L-tryptophanmethylamide
Acetyl-D-tryptophanmethylamide
Benzoyl-D-tryptophanamide
Nicotinyl-D-tryptophanamide
p-Methoxybenzoyl-D-tryptophanamide
Acetyl-L - tryptophanate
Acetyl-D -tryptophanate
Acetyl-D-tryptophan isopropylester
Tryptamine
Acetyltryptamine
Trifluoroacetyltryptamine
Indole
Indoleacetate
Indolepropionate
Indolebutyrate
Indolepropionaraide
Tyrosine series
Acetyl-D-tyrosinamide
Trifluoroacetyl-D-tyrosinamide
Chloroacetyl-D-tyrosinamide
Nicotinyl-D-tyrosinamide
Acetyl-L-tyrosinemethylamide
Formyl-L-tyrosinemethylamide
Nicotinyl-L-tyrosinemethylamide
Benzoyl-L-tyrosinemethylamide
Acetyl-L-t>Tosinate
Fluoroacetyl-L-tyrosinate
Chloroacetyl-L-tyrosinate
Acetyl-D-tyrosine ethyl ester
Nicotinyl-D-t>Tosine ethylester
D-Tyrosinehydroxamide
Acetyl-D-tyrosinehydroxamide
8.5
2.93
4.0
3.40
2.4
3.72
4.0
3.40
6.5
3.10
1.8
3.89
0.7
4.46
1.6
3.96
0.6
4.55
9.5
2.87
7.5
3.01
0.8
4.39
2.3
3.74
1.8
3.89
1.2
4.13
0.8
4.34
18
2.47
15
2.58
23
2.32
2.3
3.74
12
2.72
20
2.40
6.5
3.10
9
2.90
61
1.72
31
2.14
8.8
2.92
6.4
3.12
110
1.36
120
1.30
150
1.16
4.7
3.30
0.8
4.39
40
1.98
7.5
3.01
372 2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-20 {continued)
Inhibitor
Apparent K^ Relative — Zli^ of binding
(mM) (kcal/mole)
Phenylalanine series
Acetyl-D-phenylalaninamide 12 2 . 72
Nicotinyl-D-phenylalaninamide 9 2.90
Acetyl-D-phenylalanine methylester 2.3 3.74
Benzoate 150 1.16
Benzamide 10 2 . 83
Phenylacetate 200 0.99
Phenylacetamide 15 2.58
Phenylpropionate 25 2 . 26
Phenylpropionamide 7.0 3 . 05
Phenylbutyrate 60 1.73
Phenylbutyramide 12 2.72
" The values of K^ are taken from work at pH 7.9. A", varies with the pH and thus
the degree of ionization may be of importance in some instances. The relative binding
energies are therefore subject to some error but may provide an initial basis for discus-
sion. Various substrates have been used in the different studies and this may introduce
further uncertainties. (Data from Foster et al., 1955; Foster and Niemann, 1955 a,
b; Lands and Niemann, 1959.)
indole ring system. This is confirmed by comparing the phenyl and indole
acids and amides, the indole derivatives being bound 0.77 kcal/mole more
strongly on the average.
(C) Comparing the iV-substituted Rg groups one may calculate that the
order of binding is:
A = 0.35 A = 0.68
(kcal/mole) (kcal/mole)
Benzoyl > nicotinyl > acetyl
and it appears that the acetyl derivatives are better inhibitors than the
analogs with a free NH3+ group. It is likely that these groups interact
with the enzyme surface in a nonspecific fashion.
(D) Analogs with a CONH2 group are bound around 1.10 kcal/mole
more tightly than those with a free C00~ group. This might indicate that
the peptidatic site is in an electric field arising from surrounding negative
charges, but it could also mean that hydrogen bonds between the pep-
tide linkage and the enzyme are important. The rather tight binding of
CHYMOTRYPSIN AND OTHER PROTEOLYTIC ENZYMES 373
tryptaraine indicates also a more favorable field for positive than negative
ionic groups, but the positive charge is not important since acetyltrypt-
amine is bound even more readily.
Foster and Niemann (1955 a) determined the values of K^ for several
inhibitors at pH 6.9 and 7.9. The acetyltryptophanates and indolepropio-
nate are bound 1.0-1.4 kcal/mole more tightly at pH 6.9 than at pH 7.9,
whereas the binding of various amides is not significantly affected. This is
interpreted as pointing to the development of a negative charge in the vicin-
ity of the active site as the pH rises from 6.9 to 7.9; this charge would repel
the negatively charged tryptophanates and other carboxylates. It is pos-
sible that the lack of effect of pH on amide binding is due to the simulta-
neous deprotonation of the CONH3+ with rise in pH (p^^ = 7.5). The con-
cept of a negative charge on or near the active center was first postulated
by Neurath and Schwert (1960) on the basis of the suppressing effect of a
carboxylate group on the hydrolysis of an adjacent peptide bond. Whether
this enzyme negative group participates in the hydrolysis along the lines
suggested by Steam (1935) and Vaslow and Doherty (1953) is not certain.
The nitrogen analogs of substrates are usually not hydrolyzed and can
act as inhibitors (Kurtz and Niemann, 1961). In these compounds an a-
methine group is replaced by a N atom, or an or-methylene group by an
NH group. Thus ethyl l-acetyl-2-benzylcarbazate is an analog of ethyl
^COO— Et
CH2— N
NH- COCH3
acetyl-L-phenylalaninate and is an inhibitor, with K^ = 20 raM. Also
9)— CH2CH2— COO— CH3 and (7^— NH— CH2— COO— CH3 are substrates of
chymotrypsin whereas cp — CHg — NH — COO — C2H5 is an inhibitor [K^ =
6 m.M). It is suggested that an a-N= or a-NH — group leads to a restriction
of the rotation around the bond joining it to the CO group compared to
that of a C — C bond, this constraint leading to loss of substrate activity or
weakening of inhibitor binding.
Wallace et al. (1963) published the results of their studies of the interac-
tions of 136 compounds with chymotrypsin, and it is seen that several
substances not directly related to amino acids are fairly potent inhibitors;
such are quinoline and the methylquinolines, hydroxyquinolines, and ami-
noquinolines, various acridines, a-naphthol, and the naphthylamines, all
with K^ values less than 1 mM . They summarized their conclusions with re-
spect to structure and inhibitory activity in ten postulates, from which the
following comments are extracted. Aromatic compounds are more effective
than the corresponding saturated derivatives, and monosubstituted benzene
derivatives with polarizable uncharged groups are more inhibitory than the
374 2. ANALOGS OF ENZYME REACTION COMPONENTS
parent compounds; both facts indicate the importance of polarization of
the inhibitor molecule in the field of ionic groups on the enzyme. The ac-
tive site has a locus for interaction with the aromatic nucleus, and vicinal
to this there is at least one anionic group; the orientation of the inhibitor
is determined by the interaction of the polarizable group with a sublocus.
Molecules presenting a larger planar area are more inhibitory, and it seems
that the site at which the aromatic compounds act is mainly flat, of greater
length than breadth, and not straight but curved along the enzyme surface.
The different loci involved in the interactions perhaps have different prop-
erties; e. g., it was postulated that the active site may be hermaphroditic*
in that one locus may be electron-deficient and another electron-rich.
The chymotrypsin reaction proceeds in two steps, the acetylation of the
enzyme:
EH + AcB z^ (EH— AcB) ^ (EAc-HB) ^ EAc + HB
and the solvolysis of the acetylchymotrypsin:
EAc + HOR ^ (EAc-HOR) ^ (EH— AcOR) 5=t EH + AcOR
The competitive inhibitors tabulated here were shown by Foster (1961)
to block the acetylation reaction and not the solvolysis. The inhibition by
indole is not strictly competitive (Applewhite et al., 1958).
A'i
Relative — AF of binding
Inhibitor
(mi/)
(kcal/mole)
Skatole
0.5
4.69
Indole
0.7
4.43
p-Nitrobenzoyl - d - try ptophan
1.5
4.01
Tryptamine
2.0
3.84
p-Nitrobenzoyl-L-tryptophan
2.3
3.74
Acetyl-D-tryptophan
4.0
3.41
Acetyl-L-tryptophan
6.0
3.16
D-Tryptophan
10
2.84
L-Tryptophan
20
2.41
The relation of chymotrypsin or a chymotrypsin-like enzyme to hista-
mine release and the anaphylactic reaction was examined by Austen and
Brocklehurst (1960) in guinea pig lung sensitized to ovalbumin. Inhibitors
such as /5-phenylpropionate, indoleacetate, and indolepropionate depress
anaphylactic histamine release more than 50% at 2.5 mM, and indole and
skatole are effective at 1 mM or below. Antigen apparently activates some
proteolytic enzyme necessary for the release of histamine.
* This terminology connotes an entirely new way of looking at enzyme catalysis.
CHYMOTRYPSIN AND OTHER PROTEOLYTIC ENZYMES 375
The hydrolysis of a-benzoyl-L-argininamide by papain is inhibited com-
petitively by carbobenzoxy-L-glutamate (Kimmel and Smith, 1954; Stockell
and Smith, 1957). The inhibition is very sensitive to pH between 3.9 and
5, the inhibitory activity almost disappearing at the upper end of this range
(see Fig. 1-14-6). The y-carboxyl group has a pTiT,, of 4.4 so it is possible
that the active form is un-ionized:
coo"
/
HOOC— CH2CH2— CH /^^
Vh^— COO-CH2 — (\ /)
It is rather surprising that the ionization of this group should be so im-
portant in the binding, especially as it is at some distance from the other
probable binding groups, and the enzyme as a whole is quite positively
charged at these pH's (isoelectric point near 8.75). Perhaps a hydrogen-
bonding function must be attributed to the COOJI group instead. Benzoyl-
L-arginine {K^ = 60 mM) and benzoyl-L-ar'gininamide (iiC, = 54 mM)
also inhibit the hydrolysis of benzoyl-L-arginine ethyl ester (K,,, =23 mM)
by ficin at pH 5.5 (Bernhard and Gutfreund, 1956).
Beef spleen cathepsin C is competitively inhibited by various amides,
esters, and dipeptides when glycyl-L-tyrosinamide is the substrate (Fruton
and Mycek, 1956). L-Phenylalanine amides and esters are good inhibitors
(as shown in the accompanying tabulation) but L-phenylalanine, acetyl-L-
Inhibitor
(mM)
L-Phenylalaninamide
8.3
L-Phenylalanine ethyl ester
14
L-Phenylalanyl-L-phenylalanine
17
L-Tyrosinamide
22
DL-Phenylalanylglycine
25
D-Phenylalaninamide
68
phenylalanine, and glycyl-DL-phenylalanine are inactive. ^-Acetylation or
a free C00~ group seems to prevent binding. The interaction between the
inhibitors and the enzyme may involve several groups with possible hydro-
gen bonding of the carbonyl group in a manner similar to that proposed for
chymotrypsin. Decarboxylation and A^-acylation of amino acids can lead
to inhibitors, as in the competitive inhibition by tosylagmatine of thrombin
(Ki = 13.2 mM) and trypsin {K, = 3.45 mM) (Lorand and Rule, 1961).
This inhibitor markedly slows clotting in a thrombin-fibrinogen system and
in whole plasma.
376 2. ANALOGS OF ENZYME KEACTION COMPONENTS
HEXOKINASES
We shall now turn to various aspects of carbohydrate metabolism and
begin with those enzymes responsible for the initial phosphorylation of
sugars. Hexokinases catalyze the reaction between two types of substrate
— hexoses and ATP — and analogs of either may inhibit. Discussion of cer-
tain particularly important glucose analogs (2-deoxy-D-glucose, 6-deoxy-6-
fluoro-D-glucose, and related compounds), whose actions may involve not
only hexokinases but other early steps in carbohydrate metabolism, will
be postponed to the following section. The remaining analog inhibitors may
be classed as (A) hexoses, (B) hexose phosphates, (C) glucosamine and deri-
vatives, and (D) nucleotides and polyphosphates. It should be remembered
that the values of K„, and K^ may often be composite in that the sugar or
its derivative may occur in solution in a variety of forms (for example, a-
or /^-isomers, or pyranose or furanose rings). These forms may have quite
different activities and different individual constants.
Inhibition by Hexoses
Most hexokinases are not specific for the phosphorylation of one sugar
but act at varying rates with different hexoses. Thus yeast hexokinase
phosphorylates glucose {K„i = 0.16 mM), fructose {K„j = 1.7 mM), and
mannose (K,,, = 0.1 mM); brain hexokinase is very similar (Slein et al.,
1950). If one active site on the enzyme is responsible, competition between
the substrates should be observed. At equimolar concentrations, glucose
and mannose almost completely inhibit the phosphorylation of fructose
(88-98%), whereas fructose inhibits the phosphorylation of the former
hexoses very little (15-18%), which would be expected on the basis of their
relative iiC,„'s. The calculated K/s are essentially the same as the J^,„'s.
The inhibition of Schistosoma fructokinase* by glucose and mannose is also
quite marked, whereas galactose is much less potent (Bueding and Mac-
Kinnon, 1955). The fructokinase of rat intestinal mucosa shows similar
specificity, the following inhibitions being observed with 6 niM inhibitor
when fructose is 10 mM: glucose (90%), mannose (80%), mannoheptulose
(65%), xylose (50%), allose (0%), and galactose (0%) (Sols, 1956). Ascites
cell glucokinase is inhibited rather well by talose (jK", = 3.2 mM) and altrose
{K^ = 6 mM), but not by allose, gulose, or idose (Lange and Kohn, 1961 b).
Yeast glucokinase is competitively inhibited by mannose {K^ = 0.11 mM)
(Fromm and Zewe, 1962). Mannoheptulose induces a temporary diabetic
state by blocking the uptake of glucose into the tissues through inhibition
of glucokinase (Coore and Randle, 1964).
* The terms "glucokinase," "fructokinase," etc., will be used to designate kexo-
kinases when the corresponding substrates are used without implying that the en-
zymes are specific for these substrates.
HEXOKINASES 377
Such inhibitions may be of importance in the metabolism of mixtures
of sugars. Fructose is usually phosphorylated more rapidly than glucose,
but in mixtures of the two the phosphorylation of fructose is markedly
suppressed and essentially only glucose is metabolized. Although little
is known of the nature of the binding of these hexoses to the enzyme, it
would appear that the configurations at C-3 and C-4 are particularly im-
portant; for example, allose differs from glucose only at C-3 and is not
readily bound, and galactose differs from glucose only at C-4 and is much
less bound.
Inhibition by Hexose Phosphates
Three hexose phosphates have been found to be fairly specific and inter-
esting inhibitors of hexokinases: these are a-D-glucose-6-P, a-L-sorbose-1-P,
and l,5-anhydro-D-glucitol-6-P. Inhibiting hexose phosphates should be
visualized in their pyranose or furanose forms, since it is likely that in-
teraction with the enzyme occurs with one side of these ring structures.
(See formulas on page 378).
Glyceraldehyde has been known for many years to be an inhibitor of
glycolysis and Lardy et al. (1950) in a study to determine the site of action
found that L-glyceraldehyde prevents the phosphorylation of glucose or
fructose in brain extracts, but yet has no direct effect on the hexokinase.
This paradox was resolved by showing that aldolase catalyzes the conden-
sation of L-glyceraldehyde with glyceraldehyde-3-P to give a mixture of
D-fructose-1-P and L-sorbose-1-P. The latter compound was found to be a
potent inhibitor of hexokinase (for example, 0.08 mM inhibits 67% the
phosphorylation of glucose at 35 mM), whereas L-sorbose and L-sorbose-6-P
are inactive. The conventionally written structure for L-sorbose- 1-P
(as above) bears little obvious resemblance to D-glucose-6-P, but if the
former structure is inverted it is seen that the molecules are identical from
one side and differ only by the transposition of a hydroxyl group, as illus-
trated by Lardy et al. with molecular models. D-Glucose-6-P is the product
CH3— O— PO'a'
O
HO^
Q- L-Sorbopyranose- 1-P
of the hexokinase phosphorylation of glucose and is an inhibitor of the
reaction (see below). It is surprising that the inhibition by L-sorbose-1-P
is not competitive with glucose; indeed, as the glucose concentration is
increased, the inhibition becomes somewhat greater. It is possible that
378
2. ANALOGS OF ENZYME REACTION COMPONENTS
7^
/
"^o^
K
pK
\
^^^
— O
u-
i
\^
HEXOKINASES
379
competition might be seen if rates of inhibition in the presence of various
concentrations of glucose were determined, but once L-sorbose-1-P has
combined with the enzyme the inhibition is essentially irreversible. It is
possible that glucose forms an intermediary tightly bound complex with
the enzyme- ATP during the reaction and that L-sorbose-1-P forms a similar
complex that is stable. Actually the affinities of the brain enzyme for
D-glucose-6-P (Kj = 0.4 mM) and L-sorbose-1-P {K^ = 0.7 mM) are close
(Crane and Sols, 1954). Other hexokinases may not be so susceptible to
L-sorbose-1-P, since Taylor (1960) found only slight inhibition by 0.5 mM of
glucose uptake by Scenedesmus, the primary transfer site being hexokinase
on the outside of the membrane.
A closely related inhibitor is l,5-anhydro-D-glucitol-6-P (1,5-D-sorbitan-
6-P), this lacking the 2-OH group in L-sorbose-1-P and binding somewhat
less tightly {K^ = 1 mM) to the brain hexokinase (Crane and Sols, 1954).
The nonphosphorylated compound is a very weak inhibitor (Sols, 1956).
Since there are very few potent and specific hexokinase inhibitors, Ferrari
et al. (1959) have recently investigated l,5-anhydro-D-glucitol-6-P in some
detail to see if it might be useful as a blocking agent of this enzyme in ho-
mogenates. It is stable to the enzymes attacking D-glucose-6-P, except for
hydrolysis by liver glucose-6-phosphatase. No inhibition of glucose-6-P de-
hydrogenase is evident, but it inhibits phosphoglucomutase variably (de-
pending on the concentrations of glucose-1-P, glucose-l,6-diP, and Mg++)
and phosphoglucose isomerase noncompetitively at higher concentrations.
At 6.25 mM it blocks glucose respiration in heart homogenates but has
no influence on the oxidation of glucose-6-P, indicating under these con-
ditions a rather specific inhibition of hexokinase.
Brain hexokinase is inhibited by glucose-6-P whereas yeast hexokinase
is not (L-sorbose-1-P also does not inhibit the yeast enzyme), and the inhi-
bition has been found to be noncompetitive with respect to both glucose
and ATP (Weil-Malherbe and Bone, 1951). Inhibitions by various hexose
phosphates have been studied thorougly by Crane and Sols (1953, 1954);
the accompanying tabulation summarizes their data. The following are
noninhibitory: /?-D-glucose-l,6-diP, D-mannose-6-P, D-fructose-6-P, D-fruc-
tose-l,6diP, D-arabinose-5-P, D-ribose-5-P, D-galactose-6-P, a-glucose-l-P,
Inhibitor
imM)
Relative
! — JjP of binding
(kcal/mole)
a-D-Glucose-6-P
0.4
4.80
a-D-Glucose-l,6-diP
0.7
4.46
a-L-Sorbose-1-P
0.7
4.46
l,5-Anhydro-D-glucitol-6-P
1.0
4.25
a-D-Allose-6-P
7.0
3.05
380 2. ANALOGS OF ENZYME REACTION COMPONENTS
D-altrose-6-P, glucuronate, and glucuronate-6-P. It may be noted that glu-
cose is the only hexokinase substrate that forms an inhibitory phosphate,
indicating the importance of the configuration at C-2 for inhibition. Thus
glucose phosphorylation in a closed system slows down progressively while
that of mannose is linear, a phenomenon which may be significant in
regulating the rate of sugar utilization. In a purer preparation of brain
hexokinase. Crane and Sols confirmed that the inhibition is not formally
competitive but that a reversible EI complex is formed. Phosphorylation
at C-6 (or at C-1 in the sorbose structure) seems necessary for inhibition;
e. g., glucose-1-P and 1,5-anhydro-D-glucitol lack inhibitory activity. It is
interesting that the inversion of the phosphate-carrying group at C-1 to
form /?-glucose-l,6-diP abolishes the inhibition, possibly due to a static
interference of the now closely apposed phosphate groups. Inversion of the
groups on C-2 (mannose-6-P), C-3 (allose-6-P), or C-4 (galactose-6-P) re-
duces or abolishes the inhibition; it was felt by Crane and Sols that the
configuration at C-3 influences the effect of an adjacent group and is not
directly concerned in the binding. It is difficult in most cases to decide if
the change in affinity on inversion of the groups is related to the hydroxyl
group as a binding site or as producing steric hindrance; thus the lack
of inhibition by galactose-6-P could be due either to the loss of hydrogen
bonding through the OH group (occurring in glucose-6-P) or to a protru-
sion of the OH group preventing approach of the pyranose ring. Compar-
ison with the corresponding deoxyglucose-6-P's might be informative. We
cannot do this for C-4, but at C-2 removal of the OH group (2-deoxy-
glucose-6-P) abolishes inhibition, pointing to the OH group as a binding
site. The weak inhibitory activity of 3-deoxyglucose-6-P substantiates the
idea that the 3-OH group is not involved in binding. The retention of
inhibition in l,5-anhydro-D-glucitol-6-P likewise indicates that the 1-OH is
not a binding site, but the loss of inhibition on C-1 methylation (a-meth-
ylglucoside-6-P) shows that steric repulsion occurs when the C-1 group
becomes too large. One may conclude that binding sites are at the 2-OH
and 6-phosphate groups, and possible at the 4-OH group. The inhibitors
thus attach to a different set of enzyme sites than the substrates, only
C-4 being common to both, and Crane and Sols visualized these differences
in the following structures, where the solid circles indicate necessary binding
positions:
► H,OH •H2— O— PO",'
H )r — \
OH
Substrate Inhibitor
HEXOKINASES
381
Rat intestinal mucosa hexokinase is inhibited by glucose-6-P but only
about one-tenth as readily as the brain enzyme (Sols, 1956). The hexokinase
of Schistosoma is strongly inhibited by glucose-6-P when glucose or mannose
is the substrate, but fructose phosphorylation is unaffected (Bueding and
MacKinnon, 1955). Ascites tumor hexokinase behaves like the brain enzyme
and the K, for glucose-6-P is 0.4 mM (McComb and Yushok, 1959). Thus
inhibition of various hexokinases by glucose-6-P has been observed, but the
original observation that the yeast enzyme is resistant cannot as yet be
explained.
Inhibition by D-Glucosamine and Derivatives
Glucosamine (2-amino-D-glucose) is phosphorylated by brain hexokinase
(Harpur and Quastel 1949) and it was postulated by Quastel and Cantero
(1953) that it might be carcinostatic through ATP depletion. However, it
also competitively inhibits glucose phosphorylation and any carcinostatic
activity it possesses would be more likely related to this. Maley and Lardy
(1955) thus attempted to find a more potent inhibitor among the A^-acylated
derivatives and were quite successful, as shown in the accompanying tabu-
lation. Furthermore, these derivatives are not phosphorylated.
Ki (mM)
Glucokinase
Fructokinase
iV^-(3,5-Dinitrobenzoyl)-
0.011
0.004
A''-(m-Nitrobenzoyl)-
0.033
0.0084
iV-(p-Nitrobenzoyl)-
0.04
0.05
iV-Benzoyl-
—
0.036
iV-( w? - Aminobenzoyl)-
0.15
0.081
i\/^-(p-Aminobenzoyl)-
0.2
0.11
iV-Acetyl
—
0.46
iV-Phenylacetyl-
—
0.86
CH2OH
nA
/H
\OH
HON
\
H
1/
OH
NH,
CHoOH
NH— CO
O
Q-D-Glucosamine
iV- Benzoyl- o-D -glucosamine
Before considering the nature of this inhibition, let us examine other
hexokinases to determine how widespread is the susceptibility. The fructo-
382 2. ANALOGS OF ENZYME REACTION COMPONENTS
kinases of Schistosoma (Bueding and MacKinnon, 1955) and rat intestinal
mucosa (Sols, 1956) and the glucokinase of Spirochaeta recurrentis (P. J. C.
Smith, 1960 b) are moderately sensitive to glucosamine (50-65% inhibition
by 6-10 mM) and more sensitive to A^-acetylglucosamine (75% inhibition
by 1-2 mM). The iC, for iV-acetylglucosamine and the glucokinase of ascites
tumor cells is 0.074 mM (McComb and Yushok, 1959), indicating a binding
about 1 kcal/mole tighter than for glucose-6-P. Furthermore, both glucos-
amine and iV-acetylglucosamine inhibit the metabolism of glucose-C^* and
fructose-C^* to glycogen and COg in rat liver slices (Spiro, 1958), and the
A'^-(2?-nitrobenzoyl) and A^-(3,5-dinitrobenzoyl) derivatives inhabit glucose
uptake by Scenedesmus (Taylor, 1960). The phosphorylation of glucosamine
in liver extracts is competitively inhibited by glucose {K^ =0.11 mM),
fructose, A'^-acetylglucosamine, and hexose and glucosamine phosphates,
illustrating mutual interference by these substrates and products (McGar-
rahan and Maley, 1962).
In all these instances the inhibitions are strictly competitive with glucose
or fructose. The question arises as to why the iV-acylated derivatives are
not phosphorylated. Maley and Lardy (1955) showed by molecular models
that the iV-acyl groups do not overlap the 6-position so that some other
explanation must be sought. It was suggested that the A^-acyl groups might
interfere with the binding of ATP to the enzyme, but it is also possible that
they shift the position of the pyranose or furanose rings sufficiently so
that the 6-position is not favorably oriented for phosphorylation. It may be
mentioned that no carcinostatic activity was noted with any of these sub-
stances when tested in sarcoma-bearing mice, perhaps due to the hydrolysis
of the iV-acyl compounds by tissue cathepsins.
Kono and Quastel (1962) confirmed the glucosamine inhibition of glyco-
gen formation in rat liver slices (50% inhibition by around 0.8 mM) and
showed there to be no depression of the entry of glucose into the cells. Hexo-
kinase, phosphoglucomutase, and UDP-glucose pyrophosphorylase are in-
hibited quite weakly by glucosamine, significant effects being exerted only
at concentrations over 20 mM. UDP-glucose-glycogen glucosy transferase
in inhibited by glucosamine but not by iV-acetylglucosamine, which inhi-
bits glycogen synthesis as does glucosamine. The isolated phosphorylase
is also resistant to glucosamine. Thus the enzymes involved in glycogen
formation are not directly inhibited by glucosamine and iV-acetylglucosa-
mine. However, each of these substances stimulates phosphorylase activity
in slices when added with glucose. Thus the explanation for the reduced
glycogen formation may be an acceleration of glycogen breakdown and not
a true inhibition. Silverman (1963) found a significant reduction of glucose
oxidation by glucosamine in the rat epididymal fat pad in the presence of
insulin and felt that an inhibition of glucokinase could not entirely account
for the results, so that one must assume some action from nonmetabolized
HEXOKINASES 383
products formed from glucosamine, possibly glucosamine-P. Glucosamine
depresses the respiration and oxidation of pjTuvate in ascites cells, as
does glucose (Crabtree effect), and this is relieved by 2,4-dinitrophenol
(Ram et al., 1963). Since this is presumably related to the phosphorylation
of glucosamine and the effects on ADP-ATP levels, it constitutes another
mechanism whereby glucosamine can alter carbohydrate metabolism.
Inhibition by Adenine Nucleotides and Polyphosphates
The reaction rate of hexokinases falls with time due to the accumulation
not only of glucose-6-P but also of ADP (Sols and Crane, 1954). Phosphate,
pyrophosphate, and AMP do not inhibit. The nature of the ADP inhibition
appears to vary with the source of the hexokinase. With yeast hexokinase
the inhibition is noncompetitive with respect to ATP since around 50%
inhibition is produced by 0.5 mM ADP at all levels of ATP used (Gamble
and Najjar, 1955), and with Schistosoma glucokinase the inhibition actually
increases slightly with ATP concentration (Bueding and MacKinnon, 1955).
The inhibition of liver fructokinase by ADP is reduced slightly by increasing
the ATP concentration from 5 to 10 mM, but not enough to indicate pure
competitive inhibition; Parks et al. (1957) stated it is noncompetitive but it
might better be designated as mixed. Echinococcus fructokinase, on the
other hand, is inhibited competitively (Agosin and Aravena, 1959).
Tripolyphosphate (PgOjo^") inhibits the fermentation of glucose by in-
tact yeast cells, glycolysis in cell-free extracts, and pure hexokinase (Vish-
niac, 1950). The inhibition of hexokinase is quite potent when (ATP) = 3.75
mM: 13% at 0.47 mM, 31% at 1.4 mM, 74% at 4.7 mM, and 93% at 14
mM tripolyphosphate. The inhibition is reversed by both ATP and Mg++.
Wheat germ hexokinase appears to be more resistant to tripolyphosphate,
only 8% inhibition being given by 5 mM (Saltman, 1953).
Inhibition by Giucosone
D-Glucosone may be formed from D-glucose by mild oxidation (e. g.
with Cu++) at C-2 but, although it has been known for over 75 years, its
structure is not completely understood (Becker and May, 1949). The
following forms are possible and it is difficult to choose between them:
CH,OH
O
a)
384 2. ANALOGS OF ENZYME KEACTION COMPONENTS
OH H OH
an) (IV)
There is some evidence that the enolic tautomer II is not important and the
behavior with enzymes might favor structure I. Hynd (1927) at St. Andrews
tested D-glucosone to determine if it could counteract insulin hypoglycemic
convulsions, as glucose does, but found that, if anything, the effect of
insulin is increased. Glucosone was then administered to normal mice giving
toxic symptoms within 5 min and a well-developed insulin-like reaction in
20 min. The lethal dose range is very narrow, 2.4 mg/kg being nonlethal and
2.6 mg/kg generally lethal. Glucose injected before or with the glucosone
reduces the effects somewhat. Moribund mice following lethal doses show
an elevation in blood glucose from 0.161 to around 0.240 mg%; thus the
symptoms are not due to a hypoglycemia. Although these results would
point to glucosone interference with the utilization of glucose, Hynd un-
fortunately assumed that glucosone is formed from glucose by the action
of insulin and that, indeed, it is responsible for the effects of insulin, the
raised blood glucose levels being unexplained. Similar reactions to glucosone
are seen in several species (Herring and Hynd, 1928). The theory that insulin
induces glucosone formation was made untenable by Dixon and Harrison
(1932), who found no glucosone in the blood during insulin convulsions.
The problem rested at this stage for 20 years and then was taken up at St.
Andrews (Bayne, 1952; Mitchell and Bayne, 1952; Johnstone and Mitchell,
1953), but the results were published in a series of short and incomplete
communications without adequate data. D-Glucosone effects in mice were
not seen with up to 10 mg/kg of any other osone, including D-galactosone,
D-arabinosone, D-xylosone, L-glucosone, and 3-methyl-D-glucosone. Turning
to yeast glucose fermentation, they found no inhibition by 50 mM D-gluco-
sone but marked inhibition at 200 mM, whereas L-glucosone has no effect
at 200 mM. Becker (1954) reported almost complete inhibition of the aero-
bic and anaerobic utilization of glucose by yeast when (glucosone)/(glu-
cose) = 5.
It was realized finally by Eeg-Larsen and Laland (1954) in Oslo that the
structural similarity of glucosone to glucose might allow the former to
interfere with the utilization of the latter by blocking its phosphorylation.
This was demonstrated with ox brain hexokinase; glucosone is not phospho-
rylated but inhibits glucose phosphorylation 50% at 0.35 mM and 100%
at 2.4 mM when glucose is 2.4 mM. They concluded that the inhibition is
noncompetitive, but the small range of glucose concentrations used makes
HEXOKINASES 385
it impossible to determine the type of inhibition; actually some decrease
in the inhibition with increasing glucose was observed. As would be expected,
glucosone does not produce a Crabtree effect but blocks it (Yushok and
Batt, 1957). The inhibition of glucose fermentation in yeast depends on the
pH and the buffer system present (Hudson rnd Woodward, 1958). At pH
6.5 in phosphate buffer an inhibition of 73% of the anaerobic fermentation
of glucose was found at (glucosone) /(glucose) = 2, whereas no effect was
found at pH 3.5-4.5. The fermentation and phosphorylation of fructose are
inhibited more readily than with glucose, due to the higher ^,„ for fructose,
and the inhibitions are completely competitive with K^ for glucosone around
0.061 mM. Marked inhibition of anaerobic glycolysis in rat tissues by glu-
cosone was noted, brain being much more sensitive than tumor tissue; in
brain complete inhibition occurs with (glucosone)/(glucose) = 0.0067. The
susceptibility of brain glycolysis to glucosone is certainly much greater than
of any hexokinase studied and possibly there is an additional site of action.
In any event, these results provide sufficient explanation for the central
toxic actions of glucosone. Despite the fact that glucosone can be formed
in certain organisms (e. g., molluscan crystalline styles and red algae) and
can be metabolized in streptococci and mammals, it would appear that it
is not an important substance in intermediary metabolism and is not gen-
erally on the pathway for the synthesis of glucosamine (Becker and Day,
1953; Topper and Lipton, 1953; Dorfman et at., 1955; Bean and Hassid,
1956). There is thus no evidence that glucosone can participate in the reg-
ulation of carbohydrate metabolism, but the high susceptibility of brain
glycolysis suggests that one should withhold final judgment until the ab-
sence of glucosone in the body under various conditions has been demon-
strated.
Inhibition of Phosphafructokinase by Cycle Intermediates
The phosphofructokinase from sheep brain is quite potently inhibited by
certain cycle intermediates (see accompanying tabulation) (Passonneau and
Lowry, 1963). Although this is not strictly analog inhibition, it is worth
Inhibitor
Ki (mJ/)
Citrate
0.03
cis-Aconitate
0.1
Isocitrate
0.2
Malate
0.6
Succinate
1.5
a-Ketoglutarate
2.5
Fumarate
>10
386 2. ANALOGS OF ENZYME REACTION COMPONENTS
mentioning because of the implications such actions have for a feedback
control of carbohydrate oxidation. A rise in the levels of the inhibitory-
intermediates would reduce the rate of formation of pyruvate. The steady-
state concentrations of the cycle intermediates are certainly high enough
to inhibit significantly, but the problem of compartraentalization arises
since it is generally assumed that the tricarboxylates particularly are
mostly confined to the mitochondria. Whatever the significance of this
type of inhibition, it emphasizes the importance of compartmentalization
in regulatory control of metabolism, a factor which has not always been
taken into account.
EFFECTS OF 2-DEOXY-D-GLUCOSE
ON CARBOHYDRATE METABOLISM
An inhibitor capable of specifically blocking the glycolytic pathway
would not only be a valuable tool in biochemical investigation but might
play a role in the chemotherapy of certain neoplasms. Glucose analogs,
especially those entering the pathway and forming inhibitory intermediates,
would be the most likely candidates, and 2-deoxy-D-glucose (2-DG) is the
most interesting and best understood substance of this type. The volume
of literature during the past 10 years on this analog precludes a complete
discussion and emphasis will be placed on the sites and mechanisms of the
inhibition in the glycolytic pathway. 2-DG was first examined by Cramer
and Woodward (1952) at the Franklin Institute in the course of searching
for carcinostatic glucose analogs, and they found that it does indeed produce
some regression of Walker carcinoma and terminates embryonic develop-
ment in rats. 2-DG differs from glucose in the substitution of the 2-OH
group by a hydrogen atom and may be represented in the pyranose form as:
2 -Deoxy-D -glucose
Absorption, Distribution, and Metabolism of 2-DG
It will be well to discuss the uptake and fate of 2-DG in cells before turn-
ing to the metabolic disturbance produced. 2-DG enters most cells readily;
this may involve a phosphorylation at the membrane in some cases, but
in others it is phosphorylated only after entry. Whatever the transport
mechanism there is usually competition between 2-DG and glucose, the
EFFECTS OF 2-DEOXY-D-GLUCOSE 387
uptake of 2-DG being depressed progressively as the glucose concentration
is increased in rat diaphragm (Nakada and Wick, 1956; Kipnis, 1958)
and lymph node cells (Helmreich and Eisen, 1959). Nakada and Wick
(1956) showed that insulin can double the rate of 2-DG uptake by dia-
phragm, and Kipnis and Cori (1959, 1960) studied this in greater detail. In
normal diaphragm the 2-DG taken up appears as 2-deoxy-D-glucose-6-phos-
phate (2-DG-6-P), the rate of phosphorylation being apparently greater
than the rate of penetration. Diabetic diaphragm takes up and phosphory-
lates 2-DG at a reduced rate but 2-DG does not accumulate in the cells,
indicating the penetration is still rate-limiting. Addition of insulin acceler-
ates the uptake and some free 2-DG appears in the cells so that the phos-
phorylation is not increased proportionately. Epinephrine does not in-
fluence the penetration but slows phosphorylation of 2-DG. Transport of
2-DG across the entire diaphragm is slow, being about one-fifth the rate
for glucose and one-twenty-fifth the rate for galactose (Ungar and Psy-
choyos, 1963). It is possible that it is trapped in the muscle as 2-DG-6-P
since insulin depresses the transfer. The uptake of 2-DG by yeast in glucose-
phosphate medium is 5-10 times faster aerobically than anaerobically;
when glucose is omitted the aerobic uptake of 2-DG is not altered, but an-
aerobically there is a loss of 2-DG from the cells, so that the uptake is de-
pendent on aerobic processes and probably on the level of ATP since 2,4-
dinitrophenol acts like anaerobiosis (Kiesow, 1959). Certain fungi, such as
Neurospora crassa and Aspergillus oryzae, can grow with 2-DG as the sole
source of carbon (Sols et al., 1960 b) but not as rapidly as with glucose;
indeed, growth with glucose or other sugars is inhibited by 2-DG. E. coli
will not grow with 2-DG as the only carbon source and there is some evi-
dence that it does not penetrate into the cells (Gershanovich, 1963). The
evidence for the lack of entrance is that glycolysis is not inhibited in intact
cells where it is in extracts. 2-DG diffuses across the intestinal wall but is
not actively transported as are glucose, 1-DG, and 3-DG, indicating the
importance of the 2-position in transport (Wilson and Landau, 1960),
nor does 2-DG have an effect on the short-circuit current through the in-
testinal wall, such being associated with transport (Schultz and Zalusky,
1964; Barry et al, 1964).
There is some information on the disposal of 2-DG in intact animals.
Injected into rabbits, it is rapidly distributed in the extracellular space
and some enters the tissues in eviscerated and nephrectomized animals, the
uptake being markedly stimulated by insulin (Wick et al., 1955). Since it
produces a block of glucose uptake for at least 8 hr, it is evident that little
2-DG is metabolized beyond the 2-DG-6-P stage in extrahepatic tissues.
This is confirmed by the finding that little or no C^^Oa is expired following
injection of 2-DG-Ci* (Wick et al, 1957). Blood levels of 2-DG are more
consistent after subcutaneous injection than when it is given intraperito-
388
2. ANALOGS OF ENZYME REACTION COMPONENTS
neally, due to erratic absorption from the peritoneum (Ball and Saunders,
1958). After subcutaneous administration there is a blood peak at 15 min,
after which there is a gradual fall over 6 hr. In human subjects infused
intravenously with 50-200 mg/kg of 2-DG over 30-min periods, approxi-
mately 30% is excreted in the urine (Landau et al., 1958).
The pathways of 2-DG metabolism have not been completely worked out.
The accompanying scheme shows some of the reactions encountered. The
phosphorylation by hexokinase to 2-DG-6-P would seem to be the most
important reaction, especially as 2-DG-6-P is not metabolized in most cells
and tends to accumulate. HeLa cells can oxidize 2-DG-6-P but much more
slowly than glucose-6-P (Barban and Schulze, 1961). Hexokinases for the
deoxy-disaccharides
2-deoxy-D-glucose
6 -deoxy-D-glucono lactone
^^ 2-deoxy-D-gluconate
\
2-deoxy-D-glucose-6-P »- 2-deoxy-D-gluconate- 6- P
i \
(oxidized) D-mannonate-6-P
formation of 2-DG-6-P have been found in brain (Sols and Crane, 1953),
kidney, intestine, liver (Lange and Kohn, 1961 a), skin (Brooks etal., 1959),
diaphragm( Kipnis and Cori, 1959), HeLa cells (Barban and Schulze, 1961),
ascites carcinoma cells (McComb and Yushok, 1959; Lange and Kohn, 1961
b), and Neurospora crassa (Sols et al., 1960 b). The Michaelis constants for
2-DG are usually higher than for glucose (see tabulation) but the rates of
Hexokinase K^ {mM)
Source
Glucose
2-DG
Brain
0.01
0.024
Ascites carcinoma
0.04
0.069
Intestine
0.065
0.09
Kidney
0.048
0.04
Liver
0.04
0.09
phosphorylation are often comparable. It is interesting that a strain of
HeLa cells resistant to 2-DG has been obtained, and they are defective in
hexokinase or contain a hexokinase inhibitor; the phosphorylation rates for
2-DG, glucose, fructose, and mannose are all lower than normal (Barban,
1961). Resistance is also associated with a 5- to 10-fold increase in alkaline
phosphatase activity and this may partly account for the slower rate of
EFFECTS OF 2-DEOXY-D-GLUCOSE 389
accumulation of 2-DG-6-P (Barban, 1962 a, b). How much of the 2-DG is
oxidized directly is generally unknown, but the glucose oxidase from
As'pergillus niger oxidizes it fairly well: the relative rates of oxidation are
glucose (100), 2-DG (20), 3-DG (1), 4-DG (2), 5-DG (0.05), and 6-DG (10)
(Pazur and Kleppe, 1964). The direct oxidation of 2-DG is apparently
catalyzed by a variety of enzymes, some of the notatin type (Sols and de
la Fuente, 1957) and some of the glucose dehydrogenase type (Williams
and Eagon, 1959). The further metabolism of 2-deoxy-D-gluconate probably
varies with the tissue and has been shown in the scheme above for skin
(Brooks et at., 1960). Other pathways for 2-DG metabolism may occur in
plants, since Kocourek et al. (1963) have provided evidence for (1) /?-
glucosidation probably on C-6, (2) oxidation on C-1 to form 2-deoxyhexo-
nate lactone, and (3) epimerization to 2-deoxygalactose in tobacco plants
taking up 2-DG through the roots. The last reaction involves three enzymes
and the epimerization occurs in a complex with UDP. The abnormal di-
saccharide, /5-D-fructofuranosyl-2-deoxy-D-glucose, has been insolated from
the excised leaves of several plants following incubation with 2-DG (Barber,
1959). It is possible that a number of abnormal polysaccharides containing
2-DG will eventually be found.
Effects of 2-DG and 2-DG-6-P on Glycolytic Enzymes
2-DG inhibits the utilization of glucose and other sugars in many organ-
isms and tissues, and we shall now attempt to localize the site of this
inhibition in the early phases of glycolysis. We must consider not only
2-DG but also its primary metabolic product, 2-DG-6-P, as inhibitors.
The most likely sites for inhibition would be (1) 2-DG on hexokinases, or
(2) 2-DG-6-P on phosphoglucose isomerase, 6-phosphofructokinase, or al-
dolase with respect to glucose metabolism. Since 2-DG is phosphorylated
about as well as glucose by hexokinases it is clear that some competitive
inhibition would be observed under certain circumstances, a suggestion first
made by Cramer and Woodward (1952). However, this would appear to
be generally an unimportant factor in the over-all glycolytic inhibition,
since 2-DG equimolar with glucose does not inhibit the glucokinase of ascites
cells (Nirenberg and Hogg, 1958) and at 10 times the glucose concentration
does not inhibit HeLa cell glucokinase (Barban and Schulze, 1961). The
situation may be somewhat more complex in certain tissues, however, in-
asmuch as rat liver contains two glucose-phosphorylating enzymes, called
glucokinase and hexokinase (Walker and Rao, 1963). The sensitivities of
these enzymes are quite different (see accompanying tabulation) and a
kinetic analysis was made, the hexokinase being studied without interfer-
ence by the glucokinase since fetal liver contains only the former. The
effect of the various inhibitors, which are all competitive, varies in a com-
plex fashion as the glucose concentration is changed because of the vary-
390 2. ANALOGS OF ENZYME REACTION COMPONENTS
Ki (mM)
Inhibitor
Rat liver
Rat liver
Guinea pig
glucokinase
hexokinase
hexokinase
2-DG
14
0.3
0.6
D-Glucosamine
0.8
0.3
0.2
iV-Acetyl-D-glucosamine
0.5
0.2
0.3
Glucose (K,„)
10
0.04
0.03
ing importance of each enzyme, and it was pointed out that it is very dif-
ficult to assess the effects of such analogs in adult liver.
Most of the emphasis recently has been placed on the inhibition of
phosphoglucose isomerase by 2-DG-6-P. This inhibition is competitive on
the enzyme from rat kidney (Wick et al., 1957), rat muscle (Ferrari et al.,
1959), and ascites cells (Nirenberg and Hogg, 1958). The inhibition is reas-
onably potent (see tabulation for kidney enzyme) and it would be quite
Glucose-6-P 2-DG-6-P
{mM) (mM)
% Inhibition
I 0.5 9
1 1 24
0.5 1 79
easy for the 2-DG-6-P concentration to become greater than the glucose-6-P
concentration in cells, especially as the former is usually not metabolized
and accumulates. The inhibition of the skeletal muscle enzyme is very simi-
lar. Unfortunately the inhibitions of 6-phosphofructokinase and aldolase
by 2-DG-6-P have not yet been adequately examined, so one is left with
phosphoglucose isomerase as the site of the primary block, the conclusion
of Wick et al. (1957). However, another possible site for inhibition is the
membrane transport system for glucose, as suggested by the work of Kipnis
and Cori (1959), and this might be by either 2-DG or 2-DG-6-P. Furthermore,
Nirenberg and Hogg (1958) reported that the metabolism of fructose-1,6-
diP is blocked by 2-DG-6-P and stated that some inhibition must occur
after the phosphofructokinase step (which could be on aldolase). Other
inhibitions on enzymes metabolizing glucose but not on the main glycolytic
pathway may be mentioned. Rat liver microsomal glucose-6-phosphatase
is inhibited weakly by 2-DG (Hass and Byrne, 1960), HeLa cell glucose-6-P
EFFECTS OF 2-DEOXY-D-GLUCOSE 391
dehydrogenase is inhibited noncompetitively by 2-DG-6-P (Barban and
Schulze, 1961), 2-DG-6-P competitively inhibits the activation of rat liver
glycogen synthetase by glucose-6-P (Steiner et al., 1961), and UDP6:
a- l,4-glucan-o;-4-glucosy transferase from dog muscle is inhibited by 2-DG-
6-P with Kj =1.3 mM (Rosell-Perez and Larner, 1964). The importance
of these inhibitions in the interference produced by 2-DG on glucose me-
tabolism is not understood.
The block of fructose utilization by 2-DG may well present a different
problem. Fructose phosphorylation is inhibited much more readily than
glucose phosphorylation, presumably due to the lower affinity of the hexo-
kinases for fructose (Sols, 1956; Nirenberg and Hogg, 1958; Barban and
Schulze, 1961). The inhibition of phosphoglucose isomerase could not explain
the suppression of fructose utilization inasmuch as the fructose pathway
bypasses this step. In rat adipose tissue 2-DG has very little effect on the
metabolism of fructose although glucose metabolism is quite strongly
depressed (Fain, 1964). With glucose at 2.8 mM and 2-DG at 1.4 mM, the
formation of COg is reduced 70% and of fatty acids 89%. Nevertheless, the
stimulatory effect of insulin on fructose utilization is blocked by 2-DG
whereas the effects of insulin on glucose are unaltered. Certainly different
tissued and organisms must have various transport and enzyme systems for
the metabolism of fructose, so one should not expect a uniform action of
2-DG. The enzymes involved in the metabolism of fructose- 1-P or fruc-
tose-6-P have not been studied with respect to 2-DG inhibition.
Effects of 2-DG on Carbohydrate Metabolism and Respiration
Investigations on isolated enzymes have indicated an important block
of phosphoglucose isomerase by 2-DG-6-P and contributory inhibition of
hexokinases under certain conditions. Let us now turn to studies on carbo-
hydrate metabolism in intact cells and tissues in order to determine if the
effects of 2-DG can be explained adequately on this basis, or to accumulate
evidence of blocks elsewhere. Anaerobic glycolysis, aerobic glycolysis,
and glucose respiration are inhibited by 2-DG but to very different degrees
(Fig. 2-10). Indeed, respiration is inhibited only with high concentrations,
usually 30-100 times that required to inhibit anaerobic glycolysis compara-
bly (Woodward and Hudson, 1954; Tower, 1958), so that some workers
have reported that respiration is not inhibited (Fridhandler, 1959; Taylor,
1960). In the case of sea urchin eggs, the inhibition can be almost completely
counteracted by increasing glucose concentration (Bernstein and Black,
1959). However, glucose must be present when the 2-DG is added and is
ineffective when the inhibition has developed. The respiration of guinea pig
skin in the presence of various substrates is inhibited by 2-DG moderately
and progressively (see accompanying tabulation) (Carney et al., 1962).
All substrates and 2-DG were 20 mM. There is no effect on the endogenous
392
2. ANALOGS OF ENZYME REACTION COMPONENTS
100
50
%
INH
ANAEROBIC
GLYCOLYSIS
0.1
(2-06)
Fig. 2-10. Effects of 2-DG on the glucose metabolism in cat brain slices.
(From Tower, 1958.)
respiration. The small effect on galactose respiration was felt to be due
perhaps to a different hexokinase being used for the phosphorylation of
galactose, since inhibition is exerted by 2- and 4-deoxy galactose.
Substrate
% Respiratory inhibition at:
0-2 hr
22
-24 hr
Glucose
12
53
Mannose
10
43
Fructose
8
50
Galactose
3
21
PjTuvate
0
1
Glycolysis as measured by the formation of C^^Og from glucose-u-C^^
is depressed by 2-DG in diaphragm (Nakada and Wick, 1956), kidney (Serif
and Wick, 1958), lymph node cells (Helmreich and Eisen, 1959), and adipose
tissue (Brooks et at., 1961). The variation of inhibition with 2-DG concen-
tration is shown in Fig. 2-11 for rat kidney slices. Glycolysis as measured
by unlabeled COg or lactate formation is also inhibited in yeast (Cramer
and Woodward, 1952), ascites carcinoma and leukemic cells (Laszlo et al.,
1958), and cerebral cortex slices (Tower, 1958). The depression of aerobic,
and anaerobic glycolysis in tumor tissue is counteracted by increasing glu-
cose concentrations (Woodward and Hudson, 1954). These results are quite
EFFECTS OF 2-DEOXY-D-GLUCOSE
393
consistent in showing an inhibition of the glycolytic pathway by 2-DG.
However, the formation of CO2 from glucose is not always depressed. Frid-
handler (1959) found that although 2-DG inhibits anaerobic glycolysis
in rabbit blastocysts, the rates of respiration and COg formation are not
significantly affected. The formation of C^^Og from glucose-1-C^* in human
fetal liver is actually increased by 2-DG, but this may be attributed to a
partial inhibition of glycolysis (Villee and Loring, 1961). In ascites carci-
noma cells 2-DG simultaneously inhibits the glycolysis of glucose and
X
INH
100
^^^--''^ - DE ox Y - D - GLUCOSE
.
^^^"^"^^^
50
■ /^^^
<
/ ^^ 6- DEOXY- 6- FLUORO-D-GLUCOSE
10
20
30
40
50
60
(I)
70
mM
Fig. 2-11. Inhibition of the formation of C^^Oa from glucose-u-C*
by glucose analogs in rat kidney slices. Glucose was 10 vaM. (From
Serif and Wick, 1958.)
increases its oxidation (Christensen et al., 1961), while the disappearance of
glucose from the medium is reduced (Fig. 2-12). The increased C^^Oj formed
from glucose coupled with the depressed glucose uptake must be taken to
mean that the pathway of glucose utilization has been markedly altered,
i. e., less glucose is going to lactate and more is being oxidized. One impor-
tant factor in such tissues must be the activity of the pentose-P pathway,
which is apparently not directly inhibited but is indirectly stimulated by
2-DG. The formation of C^^Og from glucose-6-C^* in calf thymus nuclei is
inhibited essentially completely by 2-DG (McEwen et al., 1963 b). However,
394
2. ANALOGS OF ENZYME REACTION COMPONENTS
the C-l/C-6 ratio in brain slices remains the same when ghicose uptake is
reduced to one third by 2-DG (Tower, 1958).
Since hexose uptake into cells is often coupled with phosphorylation,
one would expect 2-DG to inhibit this uptake by suppressing kinase activity
directly or indirectly. Lymph node cells treated with 2-DG until 2-DG-6-P
is formed and then washed free of 2-DG do not accumulate glucose, fructose,
or mannose, and lactate formation is markedly reduced (Helmreich and
Eisen, 1959). The uptake of glucose into chick embryo hearts is 35-45%
ISO
0, FORMED
30
60
90
(2-06)
120
mM
Fig. 2-12. Effects of 2-DG on the glucose metabolism in Ehrlich ascites carcinoma
cells. Glucose-u-C* was 10 mM. Control for residual glucose taken from nonincubated
flasks with no 2-DG. Experiments 2 and 4 were averaged. (Data from Christensen
et ah, 1961.)
reduced by 40 mM 2-DG (Modignani and Foa, 1963) and into carrot slices
is reduced to about the same degree by 10 mM 2-DG, this inhibition not
being overcome by increase in glucose concentration (Grant and Beevers,
1964). 2-DG interferes with galactose uptake into mouse strain L cells but
not potently {K^ = 7.2 niM) (Maio and Rickenberg, 1962). It is quite likely
that these inhibitions are exerted predominantly by 2-DG-6-P. On the
other hand, the accumulation of a-methylglucoside, which is transported
into E. coli by the glucose carrier, is well inhibited by 1-DG, poorly by
6-DG, and not at all by 3-DG (Hagihira et al, 1963). Apparently 2-DG
is not as potent an inhibitor here as 1-DG.
Another mechanism by which 2-DG could alter carbohydrate uptake and
metabolism is by changing the levels of Pj, ADP, and ATP, since the rates
EFFECTS OF 2-DEOXY-D-GLUCOSE 395
of hexose phosphorylation and glycolytic breakdown are controlled by these.
In brain slices, 10 mM 2-DG causes a 50% fall in creatine-P and almost
complete disappearance of the adenosine polyphosphates (Tower, 1958).
There is also a diversion of phosphate to the stable 2-DG-6-P so that a cer-
tain amount of phosphate is removed from glycolytic participation, as also
pointed out by Kiesow (1960 c). Furthermore, 2-DG has been shown to
inhibit the incorporation of P/^ into ADP and ATP in ascites carcinoma
cells 70% at 10 mM (Greaser and Scholefield, 1960). El'tsina and Beresot-
skaya (1962) determined P, and nucleotide levels in tumor cells exposed to
11 raM 2-DG and found marked decreases in ATP and ADP (see accompa-
nying tabulation). On the other hand, rat liver and kidney slices show no
Tumor (
components
Control
2-DG
Zahdel
hepatoma
ATP
69.2
7.7
ADP
28.9
13.2
AMP
—
11.7
P.
157
39
Sarcoma 37
ATP
74.7
1.7
ADP
40.5
5.4
AMP
14.2
3.5
P,
140
67
significant changes, a difference attributed to variations in the activity
and cellular location of hexokinases. McComb and Yushok (1964 a) also
reported marked falls in ATP in ascites cells within 12 min after exposure
to 2-DG, and a 65% net loss of the cellular adenine nucleotides. The disap-
pearance of nucleotides is at least partly accounted for by the phosphory-
lation of 2-DG by hexokinase, the formation of AMP mediated by adeny-
late kinase, the deamination of AMP to IMP, and the splitting of IMP
to inosine by 5 '-nucleotidase (McComb and Yushok, 1964 b). They observed
a steady rise in inosine, correlated with a rise and subsequent fall in IMP.
Changes in nucleotide levels should affect various phases of metabolism
which involve these substances, and this is well seen in the effects of 2-DG
on the oxidation of ethanol in yeast (Maitra and Estabrook, 1962). When
2-DG is added to previously starved yeast in the presence of ethanol,
there is acceleration of respiration and the oxidation of NADPH, accompa-
nied by a fall in ATP with an elevation of ADP. This stimulatory phase
lasts less than a minute and is followed by a depressed phase characterized
396 2. ANALOGS or enzyme reaction components
by very low levels of P^. During the oxidation of ethanol, the ATP/ADP
and P,/ADP ratios are high; when 2-DG is added it temporarily augments
respiration by lowering these ratios, but within a minute enough phosphate
has been trapped in 2-DG-6-P to cause a marked fall in the Pj/ADP ratio
(perhaps from 17 to 1.5). The P, may now be so low that it limits the respi-
ration.
The respiration of certain tissues is diminished by the addition of glucose,
a phenomenon often called the Crabtree, or reversed Pasteur, effect; it
is particularly evident in Ehrlich ascites carcinoma cells and most of
the studies of the mechanisms involved have been on these cells. It has
been stated that an acceleration of glycolysis inhibits the oxidation of
pyruvate, but there was no real evidence to link the entire EM pathway
with respiratory control. The effects of 2-DG are thus of particular impor-
tance, since it is phosphorylated but not further metabolized to any extent.
It was shown that 2-DG inhibits respiration to about the same degree as
glucose (Ibsen et al., 1958). Respiration and pyruvate decarboxylation are
reduced 50% by 10 milf 2-DG (Ram et al, 1963). There has been dis-
agreement as to whether glucose and 2-DG act by the same mechanism or
differently. Let us briefly compare the responses to these sugars. (1) 2-DG
depresses the respiration more slowly than does glucose. Yushok (1964)
has shown in a group of sugars that the rate of respiratory inhibition is
correlated with the rate of phosphorylation. One would thus expect 2-DG
to act more slowly than glucose, so this does not constitute a real difference
in action. (2) The inhibition by glucose is released when it is all glycolyzed
but the inhibition by 2-DG remains (Ibsen et al., 1962; Hofmann et al., 1962).
This does not seem to me to be valid evidence for different mechanism of
action. (3) The addition of glucose leads to the formation of lactate whereas
2-DG does not (Ram et al., 1963). This is what would be expected, of course,
but emphasizes that glycolysis, as defined classically, is not necessary for the
effect. (4) Glucose at 10 mM inhibits the respiration 40%, 2-DG at 20 mM
inhibits it 48%, and both together inhibit it only 23% (Wenner and Cereijo-
Santalo, 1962). This was interpreted to mean that the inhibitory mechanisms
are quite different. (5) It has been stated that amobarbital prevents the
inhibition of respiration by 2-DG but not by glucose (Wenner and Cereijo-
Santalo, 1962). This is true, however, only in the presence of succinate,
since the endogenous respiration is not affected by either glucose or 2-DG
in the presence of amobarbital (there is very little to be affected). (6) The
respiratory inhibition by glucose is released by 2,4-dinitrophenol, but
there is some disagreement as to the effect of the uncoupler with 2-DG,
Ibsen et al. (1962) stating that it releases the inhibition and Ram et al.
(1963) stating that it does not. The latter workers, however, did not feel
that this is evidence for different mechanisms and were inclined to attrib-
ute the differences to the availability of glycolytic intermediates. (7) Both
EFFECTS OF 2-DEOXY-D-GLUCOSE 397
glucose and 2-DG reduce the ATP level immediately and the ADP level
very soon (Ibsen et al., 1962). The most commonly accepted explanation
of the Crabtree effect is a depletion of ADP, since the rate of mitochondrial
oxidation in ascites cells depends on the level of ADP. (8) 2-DG causes the
loss of enzymes from the ascites cells (measured with lactate dehydrogen-
ase) whereas glucose does not (Hofmann et al., 1962). Indeed, glucose
seems to antagonize this action of 2-DG. Whether this observation has any
bearing on the Crabtree effect is not known. (9) Both glucose and 2-DG
still exert respiratory inhibition in the presence of sufficient iodoacetate
to block almost completely the glycolytic pathway (Ibsen et al., 1958;
Wenner and Cereijo-Santalo, 1962). It seems that although the Crabtree
effect is not abolished by iodoacetate, it may be diminished. If the initial
phosphorylation of hexoses is responsible for the Crabtree effect, iodoacetate
would not be expected to inhibit it, except as it might reduce ATP for the
kinase reactions. (10) Glucosone inhibits the Crabtree effect produced by
both glucose and 2-DG (Yushok, 1964), but this is probably the result of
the inhibition of hexokinase by glucosone. Summarizing these results, it
would appear that glucose and 2-DG inhibit respiration by basically the
same mechanism and that this is related to their phosphorylation. It is
difficult to understand the diminished Crabtree effect in the presence of
glucose and 2-DG together, observed by Wenner and Cereijo-Santalo (1962),
but this should be investigated further, inasmuch as Ram et al. (1963)
stated that the respiratory inhibition by glucose is not enhanced by 2-DG,
apparently no antagonism being noted.
Glucose and 2-DG not only depress the oxidation of pyruvate in ascites
cells but even more strongly the oxidation and C^'^Oa formation from label-
ed palmitate (Sauermann, 1964). Inhibition of palmitate oxidation occurs
to the extent of around 80% at the relatively low concentration of 1.8
mM 2-DG. The inhibitions are approximately 29% for acetate, 58% for
pyruvate, and 92% for palmitate at 18 mM 2-DG. There would thus appear
to be some effect on fatty acid oxidation which is exerted prior to the uti-
lization of acetyl-CoA. This action is not related to the inhibition of glyco-
lysis for several reasons, including the demonstration that it occurs in the
presence of iodoacetate. Some effect on fatty acid oxidation might be
expected from a lowering of the ATP level, but it was felt that this is not
the entire explanation because of the relatively small effect on acetate
oxidation. It is not necessary, however, that a fall in ATP should affect
acetate and palmitate oxidations equally. One awaits with interest the
elucidation of this interesting effect.
The Crabtree effect has been discussed in some detail because it illus-
trates one way in which 2-DG can alter carbohydrate metabolism through
the alteration of levels of P, and the adenine nucleotides. It is quite possi-
ble that part of the effect of 2-DG on tissues is the result of a lowering
398
2. ANALOGS OF ENZYME REACTION COMPONENTS
of available ATP, this secondarily reducing hexokinase activity, and the
uptake of sugars. The greater resistance of aerobic glycolysis and respira-
tion to 2-DG, compared to anaerobic glycolysis, may be due in part to the
higher ATP levels aerobically. The differential effects on the uptakes of
different hexoses (Fig. 2-13) might be explained on the basis of whether a
hexose is actively transported or not, and the steady-state rate of its
phosphorylation (e. g., galactose metabolism is not depressed as much as
that of glucose), and the different affinities of the hexoses for the hexoki-
nases.
20
15
10
5 ■
c 0^
(% OF
ADDED )
^
--^
■"-..^GLUCOSE
\
FRUCTOSE
GALACTOSE
^
-^-^^__
10
15
20
[ 2 - DG )
Fig. 2-13. Inhibition of hexose oxidation by 2-DG in isolated rat
diaphragm. Hexoses were 10 mM. (From Nakada and Wick, 1956.)
A summary of the sites and mechanisms in the inhibition of carbohydrate
utilization by 2-DG would then include: (1) primary competitive inhibition
of certain hexokinases by 2-DG, (2) possible direct interference in the ac-
tive transport of hexoses into the cell, (3) inhibition of phosphoglucose
isomerase by 2-DG-6-P, (4) secondary reduction in transport and hexose
phosphorylation through depletion of ATP, and (5) possible inhibitions by
2-DG-6-P of glycolytic enzymes not yet examined.
EFFECTS OF 2-DEOXY-D-GLUCOSE 399
Effects of 2-DG on Various Metabolic Pathways
Infusion of acetate-C^^OO" into rabbits and determination of the respired
C^^Og were done prior to and after 2-DG injections; no alteration of acetate
oxidation was observed (Wick d al., 1957). The total COg produced decreas-
ed, partly due to the hypothermia brought about by 2-DG. This was taken
as evidence that any block by 2-DG is previous to the tricarboxylate cycle.
An effect of 2-DG on citrate levels in ascites carcinoma cells in the presence
of pyruvate was reported by Letnansky and Seelich (1960). The citrate
begins to rise 2 min after addition of 18.3 mM 2-DG and eventually reaches
levels definitely higher than in untreated suspensions. The utilization of
pyruvate is also depressed around 50%. These observations might be in-
terpreted as originating from some action on the cycle, but more recently
(Seelich and Letnansky, 1961) it was shown that methylene blue reduces
the high citrate levels and promotes pyruvate utilization in the presence of
2-DG. It was postulated that this is caused by oxidation of NADPH to
NADP, which is necessary for isocitrate oxidation. The rise in citrate may
be due to a deficiency of NADP, possibly because lactate is not formed in
the presence of 2-DG and NADPH is not oxidized, and also due to a drop in
ATP. The results can thus be satisfactorily explained on the basis of the
mechanisms previously discussed without assuming any direct action on
the cycle.
Lipogenesis in human fetal liver from glucose is depressed by 2-DG,
incorporation of l-C^* being lowered 33% and of 6-Ci* 18% at 6.1 mM
(Villee and Loring, 1961). This can, of course, be attributed to an inhi-
bition of glycolysis. Plasma fatty acids in man rise as much as 330%
following intravenous infusion of 2-DG at 60 mg/kg, and this might be
interpreted as a depression of fatty acid synthesis so that the normal equi-
librium is disturbed and fatty acids are mobilized from the tissues (Laszlo
et al., 1961). On the other hand, there is evidence that 2-DG augments
epinephrine release, so that this could be partly responsible.
The appearance of labeled amino acids from labeled glucose in brain
slices is also depressed by 2-DG (Tower, 1958). 2-DG not only prevents the
accumulation of glutamate but causes a profound fall in the intracellular
level, glutamine decreasing less markedly. It is believed that glutamate
accumulation is important in K+ uptake by brain and it was found, as
expected, that this is inhibited around 60% by 10 mM 2-DG. The role of
such changes in the central actions of 2-DG in the whole animal is not as
yet understood. The incorporation of L-valine-C^* into protein in ascites
cells is almost abolished by 10 mM 2-DG; since this may be reversed by
glucose, it is likely that the inhibition is related to glycolytic suppression
(Riggs and Walker, 1963). The DNA content of a rat carcinoma is reduced
by feeding 2-5% 2-DG in the diet and possibly this is related to the observed
inhibition in growth (Sokoloff et al., 1956).
400 2. ANALOGS OF ENZYME REACTION COMPONENTS
Effects of 2-DG on Cell Division and Growth
The growth rate of E. coli is depressed at least 50% by 10 mM 2-DG
but the differential rate of inducible /5-galactosidase synthesis is not altered
(Cohn and Horibata, 1959). Although Neurospora and Aspergillus can
grow slowly on 2-DG alone, the growth is inhibited when the usual sugars
are present (Sols et al., 1960 b). The mutiplication of influenza virus in
chick embryos is markedly inhibited by 2-DG but in preliminary studies
there was no evidence that the development of lesions in mice is altered
(Kilbourne, 1959). Glucose and, to some extent, pyruvate are able to
counteract this depression. Sea-urchin egg cleavage is delayed by 2-DG
and development is stopped at various stages, depending on the concen-
tration: 1-10 mM prevents gastrulation and the eggs reach swimming
blastulae, 100 mM delays first cleavage but the early blastula stage is
eventually reached, 200 mM causes greater cleavage delay and develop-
ment stops before the blastula stage (Bernstein and Black, 1959). Glucose
can counteract the cleavage delay. The most sensitive tissue investigated
appears to be chick embryo heart fibroblasts, since the mitotic index is
reduced from 2.65 to 0.15 by 1.52 mM 2-DG (Ely et al, 1952). Glucose at
approximately equimolar concentration is not able to reverse this inhibition
significantly. The migration of cells in the 2-DG-treated cultures is also
limited and the cells become vacuolated. These observations all point to a
general growth-inhibiting action of 2-DG, which is not unexpected, but
there has been little study of differential growth effects.
The ability of tumors to derive a good part of their energy for growth
from glycolysis prompted the original study of 2-DG as a possible carcino-
static agent, but surprisingly little work has been done on this aspect.
It has been established that glycolytic inhibition of tumors in vivo is pos-
sible, in that patients with chronic myelogenous leukemia infused with
60 mg/kg 2-DG show 35-40% inhibition of glycolysis in the leukemic cells
(Laszlo et al., 1958). The growth of cultured HeLa cells from human carci-
noma is inhibited readily by 2-DG, 5 mM producing essentially complete
depression which is reversible up to 3 days but not afterward (Barban
and Schulze, 1961). Glucose or mannose will counteract the growth inhibi-
tion. Cells in a fructose medium are inhibited more readily than when
grown on glucose or mannose, which corresponds to the greater inhibition
of fructose utilization discussed previously. When 2-DG is added at 2-5%
to the diet of rats, the growth of carcinoma G-175 in reduced (Sokoloff et al.,
1956). Adult rats tolerate this dosage well but the growth of young rats is
retarded. The 2-DG can also be injected subcutaneously at 2-4 mg/kg/day.
Mouse sarcoma is similarly affected. The inhibition in either case is not
very marked. Solid, transplanted, and ascitic tumors in mice grow more
slowly when 2-DG is administered (Laszlo et al., 1960). A modest prolonga-
tion of the survival time of the animals was noted. Definite carcinostatic
EFFECTS OF 2-DEOXY-D-GLUCOSE 401
activity in vivo has thus been demonstrated but there should be much more
work to establish if sufficient differential depression can be achieved, and
other types of neoplasm should be studied.
Effects of 2-DG on Whole Animals
The intravenous infusion of 2-DG at doses of 50-200 mg/kg in cancer
patients produces a feeling of warmth, flushing, diaphoresis, headache,
drowsiness, tachycardia, a rise in blood glucose, and a fall in white cell
count (Landau et al., 1958). Hyperglycemia has been noted in all studies
and might be attributed to a reduced utilization of glucose brought about
by glycolytic inhibition. However, other factors must be considered. Brown
and Bachrach (1959) showed that the rise in blood glucose from 2-DG can
be partially prevented by demedullation of the adrenals, indicating that
2-DG may stimulate the release of epinephrine. Increases in urinary cate-
cholamines during 2-DG infusion have also been noted (Laszlo et al., 1961).
Hokfelt and Bydgeman (1961) felt that this epinephrine release is the
primary cause of the hyperglycemia and showed that 2-DG can deplete
the adrenals of half their epinephrine. Spinal transection reduces the 2-DG
effect, indicating epinephrine release to be mediated through the central
nervous system. Pretreatment with dihydroergotamine, which blocks the
effects of epinephrine, abolishes the 2-DG hyperglycemia (Altszuler et al.,
1963; Sakata et al., 1963). It is interesting in connection with the possible
effects of 2-DG on the central nervous system that 2-DG is transported from
the blood into the cerebrospinal fluid faster than glucose; there is also com-
petition between glucose and 2-DG for the carrier (Fishman, 1964). Landau
et al. (1958), on the other hand, were inclined to discount the role of epine-
phrine since no rise in blood pressure is observed during maximal hyper-
glycemia. The hyperglycemia probably is responsible for the decreased
stainability and degranulation of pancreatic islet cells produced by 2-DG,
indicating increased activity of these cells, rather than a direct action
(Hokfelt and Hultquist, 1961). The symptoms listed above must be due
in part to the restricted glucose utilization caused by 2-DG. The rise in
blood glucose must tend to counteract this inhibition, since glucose in-
fusions reduce mortality from 2-DG, but not sufficiently. It is interesting
that the anaphylactoid reaction to dextran and ovomucoid in rats is
inhibited by 2-DG at 200 mg/kg intravenously, although the response to the
histamine releasers is not affected, indicating an important role of glucose
uptake and metabolism in certain inflammatory reactions (Goth, 1959).
The LD50 in mice is around 2.5 g/kg for single intravenous injections and
5 g/kg subcutaneously or intraperitoneally (Laszlo et al., 1960). The animals
survive several hours to a day in most cases but may die within 10 min af-
ter intravenous injection. The major toxic effects are related to the central
nervous system and are weakness, convulsions, and coma.
402 2. ANALOGS OF ENZYME REACTION COMPONENTS
Effects of 2-DG on the Heart *
The contractile tension of rat atria is progressively depressed by 10 mM
2-DG when glucose is present at its usual concentration of 5.5 mM; the inhi-
bition is 25% at 20 min, 50% at 40 min, and 70% at 90 min (A. Gimeno and
M. Gimeno). If pyruvate is present, the rate of depression is slower and the
inhibition is 30% at 90 min; if pyruvate is added at 30 min when the depres-
sion by 2-DG is around 40-45%, there is partial recovery to the — 30%
level; if glucose and pyruvate are present and 2-DG is added at 30 min,
there is a depression to the — 30% level at 90 min. The atria in the absence
of glucose progressively fail so that at 30 min the contractile tension is
50% depressed (actually the course is quite similar to that when glucose
and 2-DG are present), but with 2-DG the rate of fall is much faster, in-
dicating that 2-DG can effect the endogenous metabolism. Addition of
glucose at 30 min when the depression is 80% or more results in partial
recovery, pyruvate is less effective, and both allow return to near the
— 30% level. The rapid cessation of the 2-DG depression brought about
by either glucose or pyruvate is noteworthy. The contractile levels reached
in 90 min may be summarized in the following tabulation. The failure of
(I) Glucose-free and glucose added at 30 min — 0%
(II) Glucose-free and pyruvate added at 30 min —15%
(III) Glucose + pyruvate + 2-DG (in any order) —30%
(IV^ Glucose + 2-DG (in any order) -70%
(V) 2-DG alone -90% to -100%
pyruvate to maintain normal contractions might indicate that some 25-30%
of the tension is dependent on glycolysis, but it is also possible that 2-DG is
interfering in some way with the utilization of pyruvate. The ability of
glucose to stimulate 30 min after depression by 2-DG at 10 mM shows
that the block of glycolysis is only partial or that glucose is acting by some
other mechanism. The atrial depression by anoxia (— 85% at 10 min) is
accelerated by 2-DG (— 93% at 5 min and — 100% at 10 min), and re-
covery upon readmission of Og is much less when 2-DG is present (J. La-
cuara). Rat ventricle strip contractility is not affected over 160 min by
0.5-2 milf 2-DG, but 4 mM causes a slow depression and partially prevents
the positive inotropic effect of ouabain (E. Majeski).
* Inasmuch as so little is known of the effects of 2-DG on tissue functions and
nothing has been reported relative to the heart, this section summarizes briefly some
of the recent work done in our department and not yet published at the time of sub-
mission of the manuscript.
EFFECTS OF 6-DEOXY-6-FLUORO-D-GLUCOSE 403
It is very interesting that the atrial depression is completely unassoci-
ated with demonstrable changes in the membrane electrical characteristics
(E. Ruiz-Petrich). 2-DG at 10 mM depresses the contractile tension around
50% at 20-30 min under the conditions of these experiments, glucose being
present, but there are no changes at all of the resting potential, the action
potential magnitude, or the rates of depolarization and repolarization. The
addition of 5 mM pyruvate rapidly stopped the progression of the contractile
depression and allowed slight recovery, again without detectable altera-
tions of the membrane characteristics. 2-DG is the only inhibitor with which
we have worked that is able to affect the contractile processes so selectively,
all other inhibitors decreasing the action potential duration to varying
degrees and producing other correlated changes in the potentials. If 2-DG
acts here by reducing the utilization of glucose or glycogen, these results
would point to a close relation between the contractile process and some as-
pect of glycolysis other than the generation of ATP. It has also been shown
that atrial K+ influx and efflux are only very slightly altered by 11 mM
2-DG, and that intracellular K+ is unchanged over a period during which
the contractile activity is depressed 50% (Chin, 1963).
EFFECTS OF 6-DEOXY-6-FLUORO-D-GLUCOSE
ON METABOLISM
Modification of hexoses at the 6-position should interfere with their
phosphorylation by hexokinase, and hence any inhibition of glucose metab-
olism observed would probably not be exerted beyond the hexokinase step.
Thus inhibition produced by 6-substituted sugars should be simpler than
inhibition by 2-substituted sugars, which can be phosphorylated and may
block at several sites. Brooks et al. (1961) compared 6-deoxy-D-glucose
with 2-DG on the oxidation of glucose-u-C^* by various tissues and found
it to be somewhat less potent. When glucose is 10 mM and 6-DG is 30 mM,
the inhibitions of C^'^Og formation are 43% for adipose tissue, 23% for
kidney, and 19% for diaphragm. The inhibition in adipose tissue is reversed
by increasing glucose concentration and appears to be competitive. 6-DG
is not metabolized by adipose tissue and no C^^Oa arises from 6-DG-u-C^^.
The site of inhibition was considered to be either the membrane transport
system or hexokinase. It has been found that galactose transport across
the intestine is inhibited by 6-DG (47% at 5 mM) and that the transport of
6-DG is depressed by glucose (Wilson et al., 1960).
The replacement of the 6-0II group with fluorine to give 6-deoxy-6-
fluoro-D-glucose (6-DFG) leads, as expected, to an interesting inhibitor
of glucose utilization. The original idea was apparently to produce a gly-
colytic inhibitor with fluorine analogous to the cycle inhibitor fluoroacetate.
The initial work was done by Blakley and Boyer (1955) at Minnesota; they
404 2. ANALOGS OF ENZYME REACTION COMPONENTS
showed that fermentation of glucose and fructose by yeast is competitively
inhibited by 6-DFG. The apparent constants are shown in the following
tabulation. 6-DFG is not fermented by intact cells or yeast extracts. In
Glucose
Fructose
Yeast
Km
(milf)
Ki
{mM)
Km
{n\M)
K,
(mM)
Baker's
Brewer's
1.8
6.9
7.3
2.7
5.0
27
3.3
2.3
extracts 6-DFG has essentially no effect even at 36 milf and hexokinase
is very weakly inhibited. Glucose utilization by rat diaphragm is not in-
hibited as well as yeast fermentation, and 6-DFG is not taken up as readily
as glucose by the muscle cells. Glucose oxidase (notatin) oxidizes 6-DFG at
about 3% the rate of glucose oxidation so that direct oxidation in the
tissues is probably negligible. It was concluded that the rate-limiting re-
action for glucose utilization is different in intact cells and extracts, and
that 6-DFG probably inhibits the membrane transport of normal hexoses.
The uptake and metabolism of 6-DFG may vary from tissue to tissue.
For example, although 6-DFG is not actively transported into rat diaphragm
and insulin has no effect on this transfer, it is well transported across the
intestinal wall whereas 2-DG is not (Wick et at., 1959; Wilson and Landau,
1960). And, although 6-DFG is not metabolized in most tissues, a particulate
preparation from Aerobacter aerogenes is able to oxidize it to the corres-
ponding gluconate (Blakley and Ciferri, 1961). It is fairly certain that the
compound is quite stable and that release of fluoride does not occur suffi-
ciently to inhibit glycolysis (Serif et al, 1958).
6-DFG inhibits the formation of C^^Oa from glucose-u-C^^ in liver, kidney,
and adipose tissue, without modifying the metabolism of acetate or lactate
(Serif and Stewart, 1958; Serif and Wick, 1958; Serii et al, 1958). In general
6-DFG is somewhat less potent than 2-DG (Fig. 2-11), but the relative
sensitivities depend on the tissue studied. Nevertheless, in the eviscerated
rat 6-DFG is able to inhibit the intracellular transport of glucose quite
appreciably, e. g., 42% from 200 mg/kg (Wick et al, 1959). There is no direct
evidence that 6-DFG acts elsewhere than on membrane transport and the
comparable inhibitions produced on the metabolism of glucose- 1-C^^, glu-
cose-6-C^*, and glucose-u-C^* would point to a block prior to the glyco-
lytic-pentose phosphate shunt division (Serif and Wick, 1958). However,
hexokinases from other than yeast have not been adequately tested. Al-
though the toxicities of 2-DG and 6-DFG have not been directly compared,
Blakley and Boyer (1955) found that 250 mg/kg 6-DFG intraperitoneally
INHIBITORS OF CARBOHYDRATE METABOLISM 405
in rats produces toxic symptoms with recovery. Since the LD50 for 2-DG
is probably around 10 times this, it would indicate that 6-DFG must af-
fect the central nervous system more than the other tissues which have
been studied.
VARIOUS ANALOG INHIBITORS
OF CARBOHYDRATE METABOLISM
Numerous inhibitions of enzymes in the glycolytic and pentose phosphate
pathways by analogs have been reported. Some of these may be important
in attempts to block carbohydrate metabolism specifically and some are
undoubtedly significant in mechanisms regulating the rates in these path-
ways. We shall discuss a few of the more important enzymes and inhibi-
tions, no effort being made to include all of the observations.
Phosphorylases
The enzymes involved in the synthesis and phosphorolysis of polysac-
charides, such as starch or glycogen, are very stereospecific with respect
to substrates and inhibitors. The fairly potent inhibition of glycogen phos-
phorolysis by the product glucose- 1-P in preparations from rabbit muscle
was reported by Cori et al. (1939), 7 mM inhibiting 93%, whereas glucose-
6-P at the same concentration inhibits only 17%. The reverse reaction of
glycogen synthesis from glucose-1-P is inhibited competitively by glucose,
and to a lesser extent by mannose, galactose, and maltose, but the ajffinity
of the enzyme for these sugars is low since 30% inhibition occurs when
glucose and substrate are approximately equimolar at 17 m.M (Cori and
Cori, 1940). However, the lobster muscle phosphorylase is inhibited only
25% by 250 mM glucose when glucose-1-P is 100 mM (Cowgill, 1959).
The only known substrate for rabbit muscle phosphorylase is a-D-glu-
cose-l-P with respect to glycogen synthesis, and Cori and Cori (1940) had
found that only a-D-glucose inhibits, /?-D-glucose having little if any ef-
fect, so the question of a, /5-specificity was studied in detail on the crystal-
line enzyme by Campbell et al. (1952). It was found that /?-D-glucose-l-P
is neither a substrate nor an inhibitor, and that whereas a-methyl glucoside
inhibits, /?-metliylglucoside is without activity. The /?-anomers appear to
have no affinity for the enzyme. Furthermore, a large number of sugars
and derivatives at 50 mM were found to be inactive. The configurations of
the hydroxyl groups on all the positions of glucose seem to be necessary
for combination with the enzyme. A pyranose structure seems also to be
a requirement for inhibition, since sorbitol and inositol are without effect,
and a primary alcohol group on C-5 is necessary since D-xylose is inactive.
The failure of fructose to inhibit might be explained in several ways: (1)
the more planar furanose form is dominant, (2) the configurations on the
406 2. ANALOGS OF ENZYME REACTION COMPONENTS
C-1 of D-glucopyranose and C-2 of D-fructopyranose are different, (3)
there is a primary alcohol group on the C-5 of glucose but not on the C-6
of fructose, and (4) the /?-anomer of fructose may predominate.
An interesting type of inhibition on the phosphorolysis of starch by
monofluorophosphate was observed by Rapp and Sliwinski (1956). The
0- o-
-O— P+— OH -0— P+— F
o- o-
Orthophosphate Monofluorophosphate
sizes and electronic configurations of orthophosphate and monofluoro-
phosphate are quite similar so that interference with many reactions in-
volving phosphate might be anticipated. The inhibition of potato phosphory-
lase is completely competitive {Kj„ = 3.2 mM and K, = 2.8 mM calculated
from their ijv-ijiS) plot) and the affinity of the enzyme for the two substances
corresponds to this tectonic resemblance. The inhibition is not due to the
release of fluoride since 20 mM fluoride inhibits only 6.4% and 2.1 mM
monofluorophosphate inhibits 50%.
Phosphoglucose Isomerase
This enzyme is important in regulating carbohydrate metabolism since
its activity, along with other factors, determines how much glucose-6-P
enters the glycolytic pathway; in other words, this enzyme represents a
branching point of metabolism in the terminology of Krebs. We have seen
that inhibition by 2-DG-6-P is probably an important component of the
mechanism of action of 2-DG. The potent inhibition by 6-phosphogluconate
is particularly interesting because this substance is formed from glucose-6-P
in the pentose phosphate pathway and could determine to some extent the
diversion at the branching point. Parr (1956, 1957) reported inhibition of the
enzymes from blood, liver, muscle, and potato and found it to be competi-
tive; in the reaction glucose-6-P -> fructose-6-P, 1 mM inhibits 75% and
fructose-6-P »^ glycolytic pathway
Glucose »- glucose-6-P
-phosphogluconate ■*- pentose- P pathway
2 mM 95% (glucose-6-P 2 mM). Similar competitive inhibitions of the en-
zymes from yeast (Noltmann and Bruns, 1959) and Trichinella spiralis
(Mancilla and Agosin, 1960) have been noted. Rabbit muscle phosphoglu-
cose isomerase may be even more sensitive to 6-phosphogluconate, since the
INHIBITORS OF CARBOHYDRATE METABOLISM 407
Ki is around 0.005 mM (Kahana et al., 1960). Another potent inhibitor that
is an intermediate in the pentose-P pathway is erythrose-4-P, for which
the K^ is 0.0007- 0.001 mM {K„, for fructose-6-P is 0.08 mM) (Grazi et al,
1960). Under conditions that would lead to an accumulation of erythrose-
4-P, more glucose-6-P might be diverted into the pentose-P pathway or,
if fructose-6-P is being metabolized, a negative feedback effect would be
exerted. A third potent inhibitor is glucosamine-6-P (Wolfe and Nakada,
1956), which on the T. spiralis enzyme is around 40 times more inhibitory
than 6-phosphogluconate, 0.06 mM inhibiting 64% when glucose-6-P
is 5 mM (Mancilla and Agosin, 1960). The deamination of glucosamine-6-P
in E. coli:
Glucosamine-6-P -> fructose-6-P :f± glucose-6-P
thus terminates at fructose-6-P initially because of the inhibition of the
second reaction, but eventually disappearance of glucosamine-6-P relieves
the inhibition and glucose-6-P is formed, another example of metabolic
regulation through inhibition.
Aldolase
This enz^Tne splitting fructose- 1,6-diP to glyceraldehyde-3-P and dihy-
droxyacetone-P has unfortunately been studied very little from the stand-
point of inhibition by hexose or triose phosphates. Yeast aldolase binds sev:
eral hexose phosphates quite tightly but splits them very slowly (Richards
and Rutter, 1961), as may be seen from the K/s and the relative reac-
tion rates (rate with fructose-l,6-diP as 1) in the accompanying tabulation.
Inhibitor , ', Pielative rate
(mM)
L-Sorbose-l,6-diP
0.13
0.0014
L-Sorbose-1-P
0.2
0.0002
D-Fructose-l-P
1.0
0.0004
D-Fructose-6-P
3.8
0.0001
Muscle aldolase splits these analogs, niueh more readily. The aldolase from
rabbit Ihuscle was found by Herbert et al. (1940) to be competitively
inhibited by fructose-6-P (31%), fructose (16.3%), and glucose (5.4%), the
percentages being for 20 mM inhibitor and fructose-l,6-diP. The difficulty
in interpreting these results lies in our ignorance of the preferred form of
the substrate (i. e., a or j3, p>Tanose, furanose, or linear) for the enzyme,
and the distribution of the substrates and inhibitors among these forms
under the experimental conditions. The values of K^ may be quite misleading
408 2. ANALOGS OF ENZYME REACTION COMPONENTS
because the inhibiting form could represent only a small fraction of the total
concentration of the inhibitor.
Glyceraldehyde-3-Phosphate Dehydrogenase
A new, potent, and apparently specific inhibitor of this enzyme has
been found and is likely to be useful as a blocking agent of this step in
glycolysis. Glycolaldehyde-2-P freshly prepared does not inhibit glyceral-
dehyde-3-P dehydrogenase, but either aging the preparation or allowing it
to react in 1 N NaOH for a short time leads to the formation of a potent
inhibitor, termed tetrose-diP by Racker et al., (1959) and isolated as d-
threose-2,4-diP by Fluharty and Ballou (1959). Both the d- and L-isomers
CHO
CHO
"OgP— 0— C— H
CHa— 0— P03= —
H— C— OH
CH^— 0— PO
ycolaldehyde-2-P
D-Threose-2,4-diP
of the inhibitor were synthesized by the latter workers and only the D-iso-
mer was found to inhibit strongly. The inhibition is reversible but non-
competitive with respect to glyceraldehyde-3-P; the inhibition may actually
increase with NAD concentration. The value of K, for rabbit muscle en-
zyme is 0.0001-0.0002 milf. D-Threose-2,4-diP is oxidized by the enzyme
but only as much as the NAD present. Fluharty and Ballou postulated
that it might react with a site other than the normal catalytic site for
oxidation of glyceraldehyde-3-P but Racker etal., showed spectroscopically
that a stable acyl-enzyme complex is formed, probably with the SH group
known to be involved in the catalysis and the binding of NAD. The sub-
strate reactions might be written as:
Enzyme-NAD -f Glyceraldehyde-3-P :f± phosphoglyceryl-enzyme-NADH
Phosphoglyceryl-enzyme-NADH + P 5± glycerate-l,3-diP + enzyme-NADH
whereas the reaction with the inhibitor is:
Enzyme-NAD + threose-2,4-diP :^ diphosphothreonyl-enzyme-NADH
Normally an inorganic phosphate is transferred from a second site to the
phosphoglyceryl-enzyme to form glycerate-l,3-diP, but the location of the
2-phosphate group on the inhibitor is such as to occupy this second site
so that no phosphate can enter the reaction. This is thus an example of
an inhibition in which a substance enters the reaction sequence in the same
INHIBITORS OF CARBOHYDRATE METABOLISM 409
manner as the substrate, forming a stable complex with the enzyme, but is
unable to complete the sequence.
The use of D-threose-2,4-diP to block glycolysis in intact cells or tissues
will probably not meet with general success due to the poor penetration.
Extracts of ascites tumor cells are inhibited readily, but glycolysis in intact
ascites or HeLa cells is unaffected (Racker et al., 1959). Preparations of
glycolaldehyde-2-P, containing the tetrose-diP, inhibit the growth of some
bacteria and not others, probably depending on the degree of penetration.
The possibility was considered that the inhibitor, can be formed intracellu-
larly but so far appropriate precursors have not been found.
If photosynthesis involves the reversal of the glycolytic sequence,
D-threose-2,4-diP should inhibit, and it has been found that the total C^^Og
fixation in sonically ruptured spinach chloroplasts is reduced 57% by this
analog at 0.1 mM (Park et al., 1960). The photoreduction of 3-phospho-
glycerate is inhibited and this substance accumulates in the presence of the
inhibitor. This inhibition is due primarily to an action on glyceraldehyde-3-P
dehydrogenase leading to a deficiency of ribulose-l,5-diP, the CO2 acceptor.
Carboxy dismutase is inhibited only slightly by 0.1 mM D-threose-2,4-diP
but higher concentrations inhibit appreciably. This inhibitor may thus play
a role in photosynthetic studies.
Keleti and Telegdi (1959) examined various glyceraldehyde-3-P dehy-
drogenases and found inorganic phosphate to stimulate the activity at low
concentrations but to inhibit progressively above 20-30 mM. The inhi-
bition of the yeast enzyme is competitive with both glyceraldehyde-3-P
and NAD. Taylor et al., (1963) also found phosphate inhibition of the
hydrolysis of p-nitrophenylacetate by glyceraldehyde-3-P dehydrogenase,
the muscle enzyme being much more sensitive than the yeast enzyme.
Indeed, the muscle enzyme is 56% inhibited already at 10 mM phosphate.
Phosphate inhibitions of the glycolytic enzymes have seldom been consid-
ered as playing a role in the regulation of carbohydrate metabolism.
Enolase
Enolase catalyzes the reaction
D-2-phosphoglycerate :^ phosphoenolpyruvate
and various analogs of these substances inhibit competitively (see tabula-
tion) (Wold and Ballou, 1957). The following substances do not inhibit:
D-lactate, D-glyceraldehyde-3-P, dihydroxyacetone-P, and glycerol-2-P. An
inhibitor must have a carboxyl and a phosphate group, and the distance
between them is of some importance, although not critical. The 3-methyla-
tion of 2-phosphoglycerate does not lead to much reduction in binding, but
3-methylation of the 3-phosphoglycerate reduces the afiinity by approxi-
410
2. ANALOGS OF ENZYME REACTION COMPONENTS
Inhibitor
{mM)
D-Phospholactate
0.35
^-Hydroxypropionate-P
0.45
D - 3 - Phosphogly cerate
0.45
D-er;/<Aro-2,3-Dihydroxybutyrate
-2-P
0.60
D-er2/<7»ro-2,3-Dihydroxybutyrate-3-P
3.3
COO-
HCOP03H-
H2COH
D-2-Phosphoglycerate
coo-
I
CH,
H2COPO3H-
/9-Hydroxypropionate-P
coo-
HCOH
H2COPO3H-
D -3 - Phosphogly cerate
coo-
HCOPO3H-
HCOH
CH3
D-er7/<A/-o-2,3-Dihydroxy-
butvrate-2-P
COO-
HCOP03H-
CH3
D-Phospholactate
coo-
HCOH
HCOPO3H-
CH3
T>-erythro-2,Z-T)'\\ry-
droxybutyrate-3-P
mately 1.2 kcal/mole. This would indicate that 2- and 3-phosphates fit
comparably as long as there is no bulky group on the 3-position, but when
there is it sterically interferes with the bending of the 3-phosphate to fit the
active site.
Glucose and Glucose-6-P Dehydrogenases
Beef liver glucose dehydrogenase is inhibited strongly and competitively
by glucose-6-P (Strecker and Korkes, 1952) and fructose- 1,6-diP (Brink,
1953 a), the K-b being 0.0025 mM and 0.062 mM, respectively. The K^
for glucose is around 31 mM at pH 7, so that if this represents a dissociation
constant the phosphorylated compounds are bound much more tightly
(around 5.8 kcal/mole). The rat liver enzyme is similarly inhibited: glucose-
6-P at 0.015 mM inhibits 78% when glucose is 200 mM (Metzger et al,
1964). Glucose-1-P also inhibits but is about one-tenth as effective as glu-
cose-6-P. Ribose-5-P and fructose-6-P are also less inhibitory (Brink, 1953 a).
The potency of the glucose-6-P inhibition is surprising and it is quite likely
that it may be important in metabolic regulation or conserving glucose for
phosphorylation. Not enough is known about this enzyme or the reaction
mechanism to speculate on the nature of the interaction of these inhibitors.
INHIBITORS OF CAEBOHYDRATE METABOLISM 411
Yeast glucose-6-P dehydrogenase is inhibited competitively by glucos-
amine-6-P but in this case the affinity for the inhibitor is not as great
as for the substrate {K,,, = 0.058 mM, and K^ = 0.72 mM) (Glaser and
Brown, 1955). Neither mannose-6-P nor N-acetyl-D-gkicosamine-6-P inhi-
bits. Phosphate inhibits the yeast enzyme rather weakly but it is competitive
with respect to NADP. The enzyme from Prototheca zopfii, however, is inhi-
bited 13% by 0.07 mM phosphate, 43% by 0.14 mM, and 70% by 0.7
mM (glucose-6-P = 3 mM and NADP = 0.67 mM) (Ciferri, 1962). There
needs to be much more study of the inhibition of such enzymes if we are
to understand the controlling factors of carbohydrate metabolism.
Phosphopentose Isomerases
The phosphoribose isomerase of alfalfa is j^otently inhibited by 5-phospho-
ribonate and much less so by a variety of related substances (as shown in
the accompanying tabulation) (Axelrod and Jang, 1954). Since ribose-5-P
Inhibitor
Concentration
{mM)
% Inhibition
5-Phosphoribonate
0.13
50
Glucose-6-P
11.3
32
Phosphate
25
49
Ribose
11.5
0
Ribose-3-P
12.5
0
was 2.5 mM, the 5-phosphoribonate is bound more tightly. However, even
19 mM 5-phosphoribonate does not inhibit the growth of Leuconostoc mes-
enteroides or Lactobacillus arabinosus in glucose medium; this could mean
that the pentose phosphate pathway is not necessary for growth of these
organisms or that the inhibitor does not penetrate readily. The Ophiodon
elongatus muscle enzyme is also inhibited by 5-phosphoribonate but less
potently (28% inhibition at 0.5 mM and 45% at 5 mM) (Tarr, 1959).
The phosphoarabinose isomerase of Propionibacterium pentosaceum is not
inhibited by D-ribose, D-xylose, D-arabinose, L-arabinose, ribose-5-P, and
glucose-6-P at 10 times the concentration of arabinose-5-P (Volk, 1960),
and the phosphopentose isomerase of EcJiinococcus granvlosiis hydatid cysts
is not inhibited by glucose, fructose, glucosamine, glucose-6-P, fructose-
6-P, and mannose-6-P at 4 mM (Agosin and Aravena, 1960). However, the
latter enzyme is inhibited 51% by 1.2 mM dihydroxyacetone-P and 54% by
4 mM dihydroxyacetone, so that conditions favoring accumulation of these
substances might suppress the alternate pentose-P pathway, which could
be important in the results obtained with iodoacetate or D-threose-2,4-diP.
412 2. ANALOGS OF ENZYME REACTION COMPONENTS
Glucose-6-Phosphatase
The hydrolysis of glucose-6-P in rat liver preparations (probably microso-
mal) is inhibited by glucose with a K^ around 29 mM (Langdon and Weak-
ley, 1957). This inhibition is noncompetitive with respect to glucose-6-P,
and it was postulated by Segal (1959) that the glucose competes with the
second substrate, water, for its site; the transfer of phosphate could occur
either to water or another molecule of glucose. If this is true, incorporation
(1) Enzyme + glucose-6-P -^ enzyme-gIucose-6-P
(2) Enzyme-glucose-6-P :^ enzyme-P + glucose
(3) Enzyme-P + water ^ enzyme + Pt
(4) Enzyme-P + glucose 5=^ enzyme-glucose-6-P
of C^* from glucose-C^* into glucose-C^'*-6-P should occur and this was dem-
onstrated. It was shown that with the appropriate rate constants the re-
actions (l)-(4) (reaction (4) is of course the reverse of (2) and is included to
visualize competition between water and glucose) lead to noncompetitive
kinetics. A study of the inhibition of this incorporation by Hass and Byrne
(1960) showed that glucose is the most potent inhibitor of various sugars
tested.
Miscellaneous Inhibitions
Numerous other analog inhibitions of enzymes involved in carbohydrate
utilization have been reported, some of which have been summarized in
Table 2-21. These inhibitions along with those previously discussed point
to many mechanisms for the control of glucose metabolism. The complex
interplay between all the sugars and their phosphorylated derivatives with
respect to the inhibition of various enzymes in the different available path-
ways must always be borne in mind in work on intact cells. Many of these
enzymes are also inhibited to varying degrees by inorganic phosphate; hence
the level of phosphate can also be a regulating factor. Enzymes inhibited
by phosphate include phosphodeoxyribomutase, phosphoribose isomerase,
phosphoglucose isomerase, triosephosphate isomerase, glucose-6-P dehydro-
genase, enolase, glyceraldehyde-3-P dehydrogenase, phosphorylase, trans-
aldolase, transketolase, glucose-6-phosphatase, and ribulose-P carboxylase.
In many instances appreciable inhibition is exerted by 5-20 niM phosphate.
An interesting study of phosphate inhibition of transaldolase was made by
Bonsignore et al. (1960) in which the following reactions were examined:
Fructose-6-P + glyceraldehyde -> glyceraldehyde-3-P + fructose
Sedoheptulose-7-P + glyceraldehyde-3-P -> fructose-6-P + erythrose-4-P
Phosphate inhibits the first reaction competitively with respect to fruc-
INHIBITORS OF CARBOHYDRATE METABOLISM
413
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414 2. ANALOGS OF ENZYME REACTION COMPONENTS
tose-6-P but inhibits the second reaction noncompetitively with respect to
sedoheptulose-7-P or glyceraldehyde-3-P. Since the ^/s are roughly the
same for all inhibitions (around 60 vnM), a single binding site for phosphate
was postulated. Phosphate prevents the formation of the transaldolase-
dihydroxyacetone complex but does not interfere with the transfer of the
dihydroxyacetone to its acceptor.
Multivalent anions in general are inhibitors of glycolysis. The anaerobic
formation of lactate from glucose in pigeon hemoly sates is depressed 84%
by 40 mM sulfate, 64% by 20 mM phosphate, 100% by 4.2 mM oxalate,
and 48% by 0.1% ribonucleate (Dische and Ashwell, 1955). The reactions
from glucose -^ lactate are inhibited more strongly than from fructose-1,6-
diP — > lactate; therefore there is inhibition previous to fructose-l,6-diP. The
sequence from glucose -^ glyceraldehyde-3-P is inhibited strongly by sulfate
and ribonucleate. Aldolase is inhibited about 20% by 40 mM sulfate but is
not affected by oxalate. It was concluded that there are three sites of action,
the strongest on hexokinase, next on glyceraldehyde-3-P dehydrogenase,
and the last possibly on pyruvate kinase, the inhibitions being competitive
with ATP or NAD. Actually little positive evidence was provided for these
sites and other possibilities are just as likely; furthermore, competition with
hexose phosphate and glycerol phosphate is also possible. In addition the
complexing of Mg++ by these anions must be considered since several gly-
colytic enzymes are activated by this ion.
It will suffice to mention four additional interesting examples of analog
inhibition on this phase of metabolism. Cataractogenic sugars and polyols
inhibit lens mutarotase while noncataractogenic sugars do not; the K,'s are
4 mM for galactose, 15 mM for xylose, and 100 mM for sorbitol (Keston,
1963). Despite the weak inhibitory activity of sorbitol, there is a large
amount in the lens in certain conditions, such as diabetes (perhaps around
30 mM). Mannose is quite toxic to honeybees; of bees offered 1 M mannose
solution, 50% were dead in 90 min and over 90% in 3 hr (Sols et al., 1960 a).
It was found that bees have a hexokinase very active toward mannose
coupled with a negligible amount of phosphomannose isomerase, so that
mannose not only may interfere with phosphorylation of glucose and fruc-
tose, but many accumulate as mannose-6-P, which could disturb glycolysis
in a number of ways. Xylose appears to be in some manner a specific inhibi-
tor of photosynthesis, since Chlorella propagation is not inhibited by 0.5-
1.5% xylose when glucose, fructose, or mannose is present (thus it is not
inherently toxic), but under photosynthetic conditions the cells rapidly lose
their color and ability to divide, an effect that can be reversed by glucose
(Hassall, 1958). It was postulated that xylose may compete with xylulose-
5-P for an enzyme in the transketolase pathway and block photosynthesis;
on the other hand, a phosphorylated product may be the active inhibitor.
Analogs without the usual hexose structure may also inhibit glycolysis and
GLYCOSIDASES 415
the pentose-P pathway. Sahasrabudhe et al. (1960) in looking for carcino-
static analogs found that thiophene-2,5-dicarboxylate, which might be con-
sidered as an analog of substances such as ribose-5-P, inhibits the formation
of C^^Og from glucose- 1-C^* and glucose-6-C^'* around 43% (concentration
unspecified) in tumor tissue, and suppresses the growth in vivo of rat sar-
coma. No evidence was given as to the site of action and so the assumption
is tenuous, but the concept of using heterocyclic substances analogous to
the furanose and pyranose structures may be important.
GLYCOSIDASES
This large group of enzymes hydrolyzing the glycosidic bonds of simple
glycosides, oligosaccharides, and polysaccharides has been studied for many
years and it is not surprising that numerous instances of analog inhibition
have been observed. Most of the reports, although important in themselves,
do not lend themselves to interpretations on the molecular level; some of
the data have been briefly summarized in Table 2-22. Most of the inhibi-
tions are relatively weak but a few could definitely be significant in meta-
bolic regulation. The a, (3 configuration of the inhibitor is seen to be im-
portant in some instances. However, the enzymes are seldom completely
specific for the a- or /?-forms and the affinities often diff"er very little, as
with maltose transglucosylase (amylomaltase) from E. coli where (see ac-
companying tabulation) the binding difference between the u- and /5-glu-
Inhibitor
(mM)
Relative
(k
- AF of binding
cal/mole)
/3-Methylmaltoside
2.5
3.70
a-Methylglucoside
8
2.98
^-Methylglucoside
10
2.84
a-Phenylglucoside
10
2.84
/S-Phenylglucoside
30
2.16
cosides is 0.14-0.68 kcal/mole (Wiesmeyer and Cohn, 1960). It appears that
the hydroxyl groups on C-2, C-4, and C-6 of ring A are involved in the
CH,OH CH,OH
-Ott XT J O
ring A
416
2. ANALOGS OF ENZYME REACTION COMPONENTS
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GLYCOSIDASES
417
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418
2. ANALOGS OF ENZYME REACTION COMPONENTS
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GLYCOSIDASES 419
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420
2. ANALOGS OF ENZYME REACTION COMPONENTS
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GLYCOSIDASES
421
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422 2. ANALOGS OF ENZYME REACTION COMPONENTS
binding since alteration of the positions, as in a-methylmannoside or a-
methylgalactoside, abolishes the inhibition, and omission, as in a-methyl-
xyloside, also prevents binding. The substitution of a methyl group on C-1
of ring B (/?-methylmaltoside) does not interfere with the binding, since
this analog has approximately the same affinity for the enzyme as the sub-
strate maltose. Alteration of the glycosidic link from the o;-l,4 in maltose
to the /?-l,4 in cellobiose does not result in an inhibitor, and other changes
(as in trehalose, sucrose, dextran, or lactose) likewise reduce the affinity.
The requirements for binding may be summarized as (1) a glucose-like con-
figuration of hydroxyl groups in ring A, (2) a glycosidic link of the a- 1,4
type, and (3) a widely variable glycosidic group.
An a-mannosidase of Streptomyces griseus with a-phenylmannoside as the
substrate is inhibited competitively (probably) by several sugars and gly-
cosides (Hockenhull et al., 1954 c) and the relative binding energies have
been estimated on this basis (see tabulation), although the reliability of
Inhibitor
% Inhibition at
50 mM
Relative — Zli^ of binding
(kcal/mole)
a-Methylmannoside
96
5.23
Mannose
83
4.25
Cellobiose
82
4.20
Maltose
73
3.88
a-Methylglucoside
14
2.15
Xylose
11
1.98
Arabinose
10
1.91
Sucrose
7.5
1.72
Glucose
5
1.45
Fructose
5
1.45
Ribose
4
1.31
Mannitol
2
0.87
Rhamnose
0
—
these figures is quite low due to variations between experiments. The man-
nose configuration seems to confer strong binding, as expected, but the high
inhibitory activities of maltose and cellobiose {a and /5 glucosides, respec-
tively), especially in view of the weak inhibitions produced by a-methyl-
glucoside and glucose, are unexpected and perhaps indicate that these sub-
stances are oriented on the enzyme surface in a different way than the
substrate, although there is no necessity to postulate an irreversible com-
plex as did Hockenhull and his co-workers. The active centers for such
enzymes probably possess several binding groups for the hydroxyls arrang-
ed in a certain pattern; glycosides other than the substrate could conceivably
GLYCOSIDASES 423
interact satisfactorily with this pattern by appropriate translation or ro-
tation of the molecules (indeed, the other sides of these roughly planar
molecules could possibly fit the pattern in some cases).
One of the most interesting studies of analog inhibition was reported
by Halvorson and Ellias (1958) for the a-glucosidase of Saccharomyces
italicus, the data from which are given in the accompanying tabulation.
Inhibitor
{mM)
Relative — AF oi binding
(kcal/mole)
a-Phenylgliicopyranoside*
Glucose
0.3
1.2
5.00
4.15
a-Butylglucopyranoside
a-Ethylthioglucopyranoside
Turanose*
1.9
4
9.5
3.86
3.40
2.87
Isomaltose
11
2.78
Xylose
Sucrose*
12
23
2.72
2.32
/S-Methylmaltoside*
a-Ethylglucopyranoside
Maltose*
37
45
81
2.04
1.91
1.55
a-Ethylthioglucofuranoside
86
1.51
Arabinose
110
1.36
a-Methylglucopyranoside
400
0.56
Several of the substances (marked by *) are also substrates; maltose, in
fact, is probably the natural substrate. Inversion of hydroxyl groups on
C-2 and C-4, and substitution or oxidation at C-6, abolish the affinity.
Increasing the size of the aglycone groups increases the affinity. The rather
potent inhibition by glucose is surprising and indicates that the alkyl agly-
cone groups actually reduce the affinity. This might mean that the C-1
hydroxyl group can be involved in the binding; a small substituent prevents
this but the binding increases as the alkyl group is lengthened.
Inhibition of (3-Glucuronidases by D-Glucaro-1,4-lactone
and Related Compounds
These widely distributed enzymes catalyze the hydrolysis of /5-glucur-
onides occurring naturally in plants and animals, and in addition may play
a role in mucopolysaccharide metabolism. They are not involved in the
glucuronide syntheses of detoxification, these reactions being catalyzed by
glucuronyl transferases. There is evidence that /^-glucuronidases are related
in some manner to growth and the activities of certain tissues, and it was
424
2. ANALOGS OF ENZYME REACTION COMPONENTS
to establish their role in metabolism that Karunairatnam and Levvy (1949)
searched for specific inhibitors. The most potent of the analogs tested is
glucarate (K^ = 0.06 mM, and K,^, = 3.5 for phenyl-/?-glucuronide) but
subsequent studies in different laboratories showed great variability in ac-
tivity. The problem was solved by Levvy (1952), who found that the ac-
tive inhibitor is glucaro-l,4-lactone,* which is formed from glucarate and
occurs to varying extents in different preparations. The tabulation shows
Inhibitor
(mM)
Relative — AF oi binding
(kcal/mole)
Glucaro- 1 ,4-lactone
0.00054
8.87
3-Methylglucaro- 1 ,4-lactone
0.13
5.50
Glucarate
0.17
5.33
Glucaro-3,6-lactone
0.48
4.70
Glucuronate
1.6
3.96
Galacturonate
6.0
3.14
Galactarate (mucate)
6.0
3.14
the inhibitor constants obtained on mouse liver /^-glucuronidase hydrolyz-
ing phenolphthalein-/?-glucuronide, and the particular potency of the glu-
caro-1,4-lactone is evident."^ No inhibition at 1 mM is observed with man-
narate, mannaro-l,4-3,6-dilactone, 4-methylglucuronate, and ascorbate, or
at 10 mM with 3-methylglucarate, glucurone, 3-methylglucuronate, man-
nuronate, mannurone, and 2-keto-L-gulonate. Heat and acid treatments of
glucarate and galactarate increase the inhibitory activity greatly and it is
possible that the dicarboxylates are completely inactive. The inhibition by
glucaro- 1,4-lactone is competitive and the high potency probably due to
the structural similarity between the inhibitor and the /5-glucofuranuronide
form of the substrate. The hydroxyl configuration in the furan ring is very
important since the glucaro-3,6-lactone is bound much less tightly to the
enzyme, and the 3-OH must also be involved since methylation reduces
the binding by over 3 kcal/mole. These rather large energy differences argue
* This substance has also been called saccharo- 1,4-lactone, 1 ,4-saccharolactone,
glucosaccharo- 1,4-lactone, and other names. The generic name for the dicarboxylic
acids derived from sugars is saccharic acid and that specifically from glucose is glu-
cosaccharic acid. However, it seems that glucaric acid is used most commonly today
and this terminology is more consistent with the naming of the monocarboxylic acids,
so that the lactones will here be named accordingly.
t The figures in this and other reports for inhibitors such as glucarate, galactarate,
glucuronate, and related acids may reflect to varying degrees the presence of lactones,
since, due to the high potency of the lactones, only small amounts need be present.
GLYCOSIDASES
425
5
1
o
o-
/
'^
SB
-o
murom
tone
one)
/
^
r-O
O— A.
O-
fur;
lac
cur
O
\
-^o
-K
-Gluco
3,6-
(glu
\
o-
-K
1
B3
oa
k"
s
o
2
s
a
o
as
u-
s
o
1
-u-
EC
1
-o-
a
o
1
-u-
O
1
-o-
8
-o
1
1
o
1
1
3 4)
1
'o O X O O O
5 I I I I x"
o-u-u-u-u— u
I I I I
X O X X
X
-22
8
o-
X
o
1
-u-
X
X o
1 1
-u-u-
?8
-u-u
3 <U
5^
1
X
1 i
O X
X
1
X
*
2
'?
o
a
rt
u
0)
3
v
OiS.
426 2. ANALOGS OF ENZYME REACTION COMPONENTS
against van der Waals' forces being a major factor in the hydroxyl inter-
actions, and suggest hydrogen bonding. The terminal carboxylate group
also participates, since glucaro-3,6-lactone is much more inhibitory than
glucurone. (See formulas on page 425).
Some inhibitor constants for various /^-glucuronidases are shown in Table
2-23. Not enough /^-glucuronidases have been tested with glucaro-l,4-lac-
tone to determine the variations in susceptibility, but from the limited
data it appears that the animal enzymes are quite sensitive whereas the
plant enzyme baicalinase is much less readily inhibited. It is clear that
glucaro-l,4-lactone is one of the most potent inhibitors known and that
none of the other analogs so far examined on /^-glucuronidase is comparable,
although galactaro-l,4-lactone is undoubtedly a very effective inhibitor. It
is interesting that the /?-galacturonidases of limpet and preputial gland are
approximately as susceptible to these lactones as are the /^-glucuronidases
(Marsh and Levvy, 1958).
Limpet or-glucuronidase shows quite a different pattern of inhibition (see
accompanying tabulation for inhibitions at 5 mM with phenyl-a-glucur-
Inhibitor
% Inhibition
Relative — AF of binding
(kcal/mole)
Glucuronate
68
3.73
Mannuronate
43
3.09
Galacturonate
28
2.68
Glucurone
9
1.84
Xylono- 1 ,4-lactone
22
2.48
Glucono- 1 ,5-lactone
7
1.67
Arabono- 1 ,4-lactone
4
1.31
Glucono- 1 ,4-lactone
3
1.13
a-Glucuronate- 1 -phosphate
55
3.39
/?-Glucuronate- 1 -phosphate
22
2.48
Menthyl-a-glucuronide
55
3.39
Menthyl-/S-glucuronide
6
1.57
Borneol-a-glucuronide
41
3.04
Veratroyl-/3-glucuronide
9
1.84
onide at 1 raM with relative binding energies calculated on the basis of
competitive inhibition). a-Glucuronides are more inhibitory than /5-glucur-
onides, as expected, but no particularly potent inhibitors were found {Kj^
for glucuronate is 0.54 mM and for galacturonate 4.7 mM). The following
do not inhibit: mannurone, glucaro-l,4-lactone, mannono-l,4-lactone, and
galactarate. The difference between the a and /? enzymes with respect to
glucaro-l,4-lactone is especially striking.
GLYCOSIDASES
427
A
00 -^ (M — O -*
• c
O -H CD — O O
Id
_^ CO <M C5 05
rZT o c-i t^ ^ CO
c
c c c c
ooooco-*-^-^ —
'H :2
O GO CO .,
O Oi o (^i
o o o «^
O O O O >0 CO
o o o o ^ -<t
P -:, ci:i
>» = -^ ^
■*^ Si A lA
03 03 eS ^^ -*^
o o o o o o
> a
a O
o ^ p
> :§ o
3 S O ^
T3 7:=
« o
-r TTn =5 3 -T _. -Q.
^ Is « C3 5
^ 6 rt "ci 5 g P
2 ^ "3 ^ le -2 Is
S O a O O O O
0)
-r)
r2
'6
'S
*2
"S
o
o
o
^
tH
«H
3
3
3
O
CJ
o
_3
'S
"3i
bC
tiC
'S
^^
oi.
^
**?-
%.
o
3
o
s
'S
-a
o
u
>>
^
N
"^
"eg
'm
c
j3
<ai.
■>*
^
Bf
T.
ec
'o
"o
3
a
3
<»
3
3
4)
a
Ph
Pl,
PlH
M
M
oi.
<ai.
^
r-
r2
«
'3
13
jil
43
-►i
4i
^
J3
a.
ft
"o
o
3
3
05
(»
43
^
Ph
M
GQ
is
^
+3
OJ
^
3
'^^
<u
-ti
^
e
cS
^
05
^
3 C
03
^
(C
3
"^ .^
^
O
ai
3
0)
-tj
-S
-0
CJ
c
<A
^
03
.s
OJ
3
' — V
Q
"3
-3 CO
2 "o
^H
cS o
f* cS
eS 73
c
<1)
o
ft
cS
3
frt
o
4J
Tl
t^
tS
ft
428 2. ANALOGS OF ENZYME REACTION COMPONENTS
It was previously stated that /^-glucuronidases do not seem to partici-
pate in glucuronide synthesis in tissues and the evidence is mainly the
lack of inhibition by glucaro-l,4-lactone usually observed (Levvy and
Marsh, 1960). For example, Lathe and Walker (1958) found no effects of
2 mM glucaro-l,4-lactone on the formation of o-aminophenylglucuronide
or bilirubin-glucuronide in liver suspensions. However, Fishman and Green
(1957) reported that glucarate potently inhibits glucuronyl transfer (50%
inhibition by 0.05 mM), and Sie and Fishman (1954) found glucarate and
glucurone to inhibit glucuronide synthesis in liver slices. It is not definitely
known if these inhibitions relate to a /^-glucuronidase or a transferase, but
the question of the role of the /^-glucuronidases in glucuronide synthesis
should probably be left open.
The urinary bladder cancer that occurs in people employed in certain
industries may be related to the formation of glucuronides of aromatic
amines in the body and their subsequent hydrolysis by urinary /5-glucur-
onidase. The oral administration of inhibitory analogs to patients is a
possible approach to the prevention of such cancers. Boy land et al. (1957)
found that the urinary enzyme can be inhibited by administration of glu-
conate and glucarate at a dose of 10 g/day, but the most potent inhibition
is produced by glucaro-l,4-lactone, 73% inhibition occurring from 1.5-2
g/day and 90% inhibition from 4 g/day. Nevertheless, these inhibitions are
less than expected and the urinary pH was reported later to be an important
factor, the inhibition decreasing as the pH rises above 6 (Boyland et al.,
1959). Inhibition of liver /^-glucuronidase in mice by the administration
of glucaro-l,4-lactone at doses from 50 to 800 mg/kg (17% to 76%) was
found by Akamatsu et al. (1961). The maximal inhibition occurs at 30-60
min and after 2-4 hr most of the activity has returned. Similar results
were found in rat liver and kidney. Since /^-glucuronidase and /?-iV-acetyl-
glucosaminidase hydrolyze products from chondroitin and hyaluronate and
may participate in the metabolism of connective tissue mucoproteins, and
since certain tumors have relatively large amounts of these enzymes, Carr
(1963) administered the two inhibitors — glucaro-l,4-lactone and 2-acet-
amido-2-deoxygluconolactone — to mice bearing Tumor 2146. At 150 mg/kg
these substances are nontoxic and cause regression of the tumors. It was
suggested that the inhibitors prevent the penetration of the tumor cells
through the intercellular cement substance, but this is admittedly only a
tenuous hypothesis. Whatever the explanation, it represents an interesting
approach to tumor chemotherapy.
Inhibition of Various Glycosidases by Glucono- and Glucaro Lactones
The inhibition of the /^-glucuronidases by the saccharo-l,4-lactones sug-
gested that the /^-glycosidases might be inhibited by the aldonolactones,
and this was found to be so by Conchie (1953, 1954). A sheep rumen /?-
PYRUVATE METABOLISM 429
glucosidase, possibly involved in the digestion of cellulose, is strongly inhi-
bited by glucono-l,4-lactone (^, = 0.094 mM) and glucono-l,5-lactone
(^. = 0.091 milf ) in a competitive manner, the affinity of the enzyme for
these analogs being about 10 times that for the substrate o-nitrophenyl-
/?-glucoside, whereas gluconate itself has no action. Ox liver /?-galactosidase
is inhibited much more potently by the galactono- and fucono-l,5-lactones
than the corresponding 1,4-lactones, and this can be readily explained on
the basis of the relationship to substrate configuration (Lewy et al., 1962).
A remarkable degree of specificity is exhibited by the a and /? glycosidases,
inhibition usually resulting only from the corresponding aldonolactone in
a number of enzymes from different sources (Conchie and Lewy, 1957).
For example, limpet a-mannosidase is inhibited markedly by mannono-
1,4-lactone but not by the glucono-, galactono-, arabono-, or xylonolactones.
On the other hand, galactono- 1,4-lactone is specific for /?-galactosidase
(Conchie and Hay, 1959). Cellulytic rumen enzymes are inhibited to vary-
ing degrees by glucono- 1,4-lactone, depending on the substrate chain length;
hydrolysis of cellobiose is inhibited 99%, of cellotriose 90%, and of cellote-
traose 75% by 0.5 mM (Festenstein, 1959). Glucono- 1,4-lactone also inhibits
a yeast debranching isoamylase (Gunja et al., 1961) and a mammalian
thioglycosidase (Goodman et al., 1959), so that this type of inhibition ap-
pears to be widespread.
Very potent and specific inhibitions are exerted on N-acetyl-/?-glucos-
aminidase and N-acetyl-/?-galactosaminidase by the corresponding lactones
(Marsh and Lewy, 1957; Findlay et al, 1958). The former enzyme from
epididymis is inhibited by N-acetylglucosaminolactone competitively, with
K- = 0.000072 mil/, but the limpet enzyme is less sensitive, the K^ being
0.027 mM. The N-acetyl-a-glucosaminidase, on the other hand, is inhibited
much less. The acetyl group is essential for the inhibition and cannot be
replaced by other acyl groups. Certainly the use of the corresponding lac-
tones for specific and potent inhibition of the glycosidases has been one of
the most successful endeavors in the application of analogs.
PYRUVATE METABOLISM
The usefulness of a direct and specific inhibitor of pyruvate utilization
is obvious and the natural occurrence of certain pyruvate analogs makes
this field of inhibitor study an important one, but relatively little work
has been done. Fluoropyruvate will be taken up with other fluorinated
compounds, such as fluoroacetate, in a separate chapter. Phenylpyruvate
is the best studied of the other pyruvate analogs. It is readily formed from
phenylalanine and is usually decarboxylated to phenylacetate, although
some may be reduced to phenyllactate. In phenylketonuria (phenylpyruvic
oligophrenia), the accumulation of phenylpyruvate could be responsible for
430 2. ANALOGS OF ENZYME REACTION COMPONENTS
some of the disturbances by interfering with pyruvate metabolism, but no
work has been done on the metabolic changes in tissues resulting from
such levels of the analog. Hydroxyphenylpyruvate is similarly formed from
tyrosine.
(. ))— CH2— CH- COO"
Phenylalanine
The formation of acetoin from pyruvate in Streptococcus fecalis is inhibited
13% by 1 mM and 75% by 10 mM phenylpyruvate, whereas the formation
of acetoin from acetoacetate is unaffected by 10 mM (Dolin and Gunsalus,
1951). Pyruvate oxidase is inhibited to about the same degree. Several
anaerobic pyruvate pathways are inhibited by phenylpyruvate in several
bacteria and yeast, including the formation of acetoin, the phosphoroclastic
reaction, and decarboxylation, whereas the oxidative metabolism of pyru-
vate is not so readily affected (Watt and Werkman, 1954). The concentra-
tions of pyruvate and phenylpyruvate used (120 mM) are unfortunately
too high to be physiologically significant, but further study on extracts
of Aerobacter aerogenes showed competitive inhibition of acetoin formation
with K,„ = 123-197 mM, and K^ = 0.59-1.1 mM, so that in this case phen-
ylpyruvate is bound to the enzyme much more tightly than pyruvate.
The reduction of hydroxypyruvate to glycerate by glycerate dehydrogenase
from spinach is inhibited by phenylpyruvate (5%), pyruvate (34%), and
bromopyruvate (52%) at 10 mM (Holzer and Holldorf, 1957). These inhi-
bitions are competitive but rather weak.
The most pertinent study with respect to blocking an important pyruvate
pathway is that of Gale (1961) on yeast pyruvate decarboxylase, which
reports the inhibitions given in Table 2-24. The following compounds are
inactive: pyruvic ethyl ester, oxalacetate, propionate, phenyllactate, phen-
ylalanine, acetamide, oxamate, and oxalate. One might infer that (1) the
C=0 group is necessary for inhibition (reduction or substitution abolishes
activity), and (2) the COO" group is necessary for strong inhibition (amides
and esters inactive). The nature of the R group in R — CO — COO" can vary
quite widely and it is difficult to correlate structure with activity; for
example, it is surprising that ketomalonate is bound so well and chloro-
pyruvate relatively poorly, and that oxanilate is bound so very weakly.
PYRUVATE METABOLISM
431
Table 2-24
Inhibition of Yeast Pyruvate Decarboxylase by Analogs"
Inhibitor
Structure
Concen-
tration
(mM)
Inhibition
Relative
activity
Glyoxylate
o-Nitrophenyl-
pyruvate
Ketomalonate
/)-Hydroxyphe-
nylpyruvate
Phenylpyruvate
Chloropyruvate
2, 3-Butanedione
O
II
H-C-COO
i>
CH,
O
II
c-coo
o
II
ooc-c-coo
HO
CHj— C-COO'
O
II
CH,— C-COO
O
II
CI— CH,— C-COO
O O
II II
HoC C C CHo
0.023
33
0.45
95
0.09
61
0.14
70
0.23
71
0.34
82
0.45
62
1.1
71
1.1
68
2.3
80
0.45
27
2.3
73
32
17
12
2.9
56
1.0
0.14
Oxanilate
a-Ketoglutarate
O
II
NH- C-COO'
OOC-CH,— CH,— C— COO
27
31
24
0.05
0.012
o Pyruvate concentration 4.5 mW and preincubation with inhibitor 15 mln. Relative inhibitory
activity calculated from the formula t/(l - i)(I) and roughly would be inversely proportional
to Kf for noncompetitive Inhibition. (From Gale, 1961.)
The inhibition by ketomalonate is more surprising when one considers that
oxalate and oxalacetate are inactive. The kinetics of the inhibitions pro-
duced by the five most potent substances were studied in greater detail
and a typical noncompetitive mechanism was established. However, pre-
sence of the substrate prevents development of the inhibitions, suggesting
that the inhibitors combine at the substrate site. These results indicate an
irreversible or pseudoirreversible type of inhibition, and it was indeed de-
monstrated that the inhibitions are all progressive with time. Phenylpy-
ruvate is the only inhibitor whose effects are even partially reversible by
432 2. ANALOGS OF ENZYME REACTION COMPONENTS
dialysis. It was considered that the true inhibitors are the aldehydes cor-
responding to the keto acids, and it was shown that COg is evolved from
phenylpyruvate and p-hydroxyphenylpyruvate but not from the other
three inhibitors. Although this does not completely invalidate the aldehyde
proposal, it makes it unlikely. On the other hand, it is possible that the
tight complex is formed with some intermediate prior to decarboxylation.
Hydroxypyruvate is decarboxylated by yeast decarboxylase at a rate
1/76 that of pyruvate and strongly inhibits the pyruvate reaction (Holzer
et al., 1955 d). It was thought that hydroxypyruvate might exert some
regulatory function on pyruvate metabolism in yeast. Formaldehyde and
acetaldehyde inhibit pyruvate decarboxylase but the mechanism may not
be competitive with substrate (Bauchop and Dawes, 1959). The oxidation
of pyruvate in rat kidney slices is inhibited around 70% by 20 mM meso-
tartrate, but not at all by d- and DL-tartrates (after correction for the inhi-
bition of endogenous respiration) (Quastel and Scholefield, 1955). This
inhibition is completely reversed by fumarate and malate. However, when
mitochondria were examined it was found that there is little direct effect on
pyruvate oxidation but a rather strong inhibition of or-ketoglutarate oxida-
tion, which is progressive with time. In the presence of bicarbonate, pyru-
vate is oxidized and this is inhibited by meso-tartrate. It was assumed
that the inhibition is upon the incorporation of CO.2, and it was stated that
meso-tartrate is a specific inhibitor of pyruvate oxidation in slices of rat
kidney cortex, a conclusion that would seem to be unjustified since so few
systems were tested.
LACTATE METABOLISM
A specific inhibitor of lactate dehydrogenase would be useful in studying
the effects of a block of glycolytic lactate formation and for determining
the role of this enzyme in the functioning of tissues. One of the most in-
teresting lactate dehydrogenase inhibitors is oxamate, first reported by
Hakala et al. (1953), who stated that it is the most potent inhibitor of many
lactate and pyruvate analogs examined. The inhibition is competitive with
O
+H3N— C— coo-
Oxamate
respect to pyruvate,* noncompetitive with respect to lactate and NAD,
and uncompetitive with respect to NADH (Papaconstantinou and Colo-
* Examination of the double reciprocal plot reveals that the situation is not purely
competitive but mainly so.
LACTATE METABOLISM 433
wick, 1957; Novoa et al., 1959). The substrate and inhibitor constants have
been summarized by Papaconstantinou and Colo wick (1961 a), as shown
in the accompanying tabulation; if the KJs, are dissociation constants,
Source
A',„ (pyruvate) K^ (oxamate)
{mM) (mif)
Beef heart 0.137 0.0374
Ascites carcinoma 0.212 0.0563
Rabbit muscle 0.302 0.10
oxamate is bound around 0.77 kcal/mole more tightly than pyruvate. The
difficulty in interpreting K^ is that oxamate complexes with the apode-
hydrogenase, E, with E-NAD, and with E-NADH, the dissociation con-
stants being different for each (Novoa et al., 1959). The values for three
different pH's are shown in the accompanying tabulation. Such ternary
K
i (mM) at:
Complex
pH 6.40
pH 8.45
pH 9.70
E-oxamate
0.10
0.78
20
E-NAD-oxamate
0.069
0.57
11
E-NADH-oxamate
0.026
0.17
1.6
complexes have recently been demonstrated by ultracentrifugal separation,
1 mole of oxamate being bound for each mole of NAD or NADH (Novoa
and Schwert, 1961). The dissociation constant for the E-NADH-oxamate
complex determined ultracentrifu gaily is 0.011 mM at pH 7.4. No evidence
could be found for a complex with the apoenzyme alone; whether such a
complex occurs or not, the inhibition is mainly due to the ternary complexes.
The lactate dehydrogenase from human liver and heart is also inhibited by
oxamate (Plummer and Wilkinson, 1963). The reduction of 2-ketobutyrate
is inhibited more strongly than pyruvate reduction, presumably because
2-ketobutyrate is bound to the enzyme less tightly, but the reduction of
succinic semialdehyde by a rat brain lactate dehydrogenase is inhibited
to the same degree as the reduction of pyruvate, namely, 88% by 0.1 mM
(Fishbein and Bessman, 1964). Not all lactate dehydrogenases are sensitive
to oxamate; that from L. mesenteroides is inhibited only 50% by 7 mM
(Papaconstantinou and Colowick, 1961 a). Oxamate specifically inhibits the
L( + )-lactate dehydrogenase and does not affect d( — )-lactate dehydroge-
^-
434 2. ANALOGS OF ENZYME REACTION COMPONENTS
nase up to 8 raM (Dennis and Kaplan, 1960). The active site of the l( + )-
lactate dehydrogenase was represented as containing a cationic group for
electrostatic interaction with the C00~ group, and a hydrogen bonding
group which interacts with the OH group of lactate and the NHg group of
oxamate (it might also bond to the CO or enolized COH group of oxamate).
The role of the amino group of oxamate on the binding is not known, nor
can the importance of the enolic tautomer of oxamate be evaluated. The
marked decrease in inhibition between pH 8.45 and 9.70 (see tabulation
above) could indicate the deprotonation of an amino group, but it could
be on the enzyme as well as the inhibitor.
The aerobic lactate production in human leucocytes is inhibited less than
25% by 10 mM oxamate, even in broken cell suspensions, suggesting that
lactate dehydrogenase is not solely responsible for lactate formation, al-
though the sensitivity of the leucocytic enz^^me to oxamate has not been
examined (McKinney et al., 1955). The effects of oxamate on tumor cell
metabolism and growth have been studied thoroughly by Papaconstantinou
and Colowick (1957, 1961 a, b). Anaerobic glycolysis in ascites carcinoma
cells is inhibited 50% by 8 mM oxamate and aerobic glycolysis is similarly
depressed. This may indicate that oxamate does not penetrate into cells
readily, since the K^ of 0.0563 mM for ascites cell lactate dehydrogenase
would lead one to expect a greater effect at this concentration. The inhi-
bition of anaerobic glycolysis decreases with time due to the accumulation
of pyruvate, whereas no accumulation of pyruvate occurs aerobically, in-
dicating that oxamate has little effect on pyruvate oxidase (an 18% inhi-
bition of pyruvate oxidation by 10 mM oxamate was observed). A decrease
in the inhibition anaerobically with time was also noted in Tetrahymena
pyriformis (Warnock and van Eys, 1963). The growth of HeLa cells is
completely inhibited by 40-80 mM oxamate and this is paralleled by de-
creases in glucose uptake and lactate formation, so that lactate dehydro-
genase appears in some manner to be essential for the growth of these cells
(assuming that oxamate acts specifically on lactate dehydrogenase). It was
proposed that oxamate might be a useful inhibitor for selectively blocking
glycolysis in mammalian cells. Mice can tolerate quite large doses (1 g/kg),
however. A block of glycolysis, of course, refers here only to an inhibition
of lactate formation, and the formation or utilization of pyruvate should
not be significantly affected, so it would seem that aerobic glucose meta-
bolism, at least with respect to the generation of energy, would be resistant
to oxamate.
A number of disturbing observations have appeared which cast some
doubt on the simple concept that oxamate specifically inhibits lactate de-
hydrogenase. Leached HeLa cells restored to normal conditions actively
extrude Na+ and accumulate K+; these processes are inhibited 50% and
77%, respectively, by 38 mM oxamate (Wickson-Ginzburg and Solomon,
LACTATE METABOLISM 435
1963). Inasmuch as the conditions were aerobic here, it is difficult to un-
derstand how an inhibition of lactate dehydrogenase would account for the
marked effects observed, unless there is an elevation of the NADH/NAD
ratio due to the prevention of pyruvate reduction, this slowing the oxidation
of glyceraldehyde-3-P and reducing the generation of ATP. This does not
seem to occur in Ehrlich ascites carcinoma cells inasmuch as oxamate stim-
ulates the formation of C^^Og from glucose-6-C^* without affecting that
from glucose- 1-C^'* (Christensen and Wick, 1963). The stimulation is thus
associated only with oxidation through the cycle and there is no effect on
the fraction going through the pentose-P pathway. More pyruvate enters
the cycle since less goes to lactate. The effects of oxamate on glucose utiliza-
tion will depend for one thing on how rapidly pyruvate can be oxidized.
The Crabtree effect is abolished almost completely by 40 uiM oxamate, i.e.,
in the presence of glucose, oxamate stimulates the respiration in ascites
cells (Papaconstantinou and Colowick, 1961 a). Simultaneously, glucose up-
take is depressed 40% and lactate formation 70%. In view of the conclusion
about the nature of the Crabtree effect in the section on 2-DG, it would
seem that inhibition of lactate dehydrogenase could not be responsible for
this effect of oxamate. It is possible that oxamate diverts more pyruvate
into the cycle and hence stimulates respiration under these conditions, but
this would not be a true abolition of the Crabtree effect. One would not
expect glucose respiration to be depressed by oxamate in any case if the
only action is on lactate dehydrogenase, but 31% respiratory depression
is produced by 10 mM oxamate in guinea pig alveolar macrophages (Oren
et at., 1963). Despite the statements relative to the specificity of oxamate,
it must be admitted that very few enzymes or metabolic pathways have
been studied. It is quite likely that certain phases of amino acid metabolism
might also be inhibited, since oxamate could be considered as an amino
acid analog. It will also be noted that all the effects discussed in this para-
graph were produced by oxamate at the high concentration of 10 raM
or above.
Oxalate often inhibits lactate-metabolizing enzymes as potently as does
oxamate and similarly forms ternary complexes with lactate dehydrogenase
and NAD. However, it differs from oxamate in being competitive with
lactate instead of pyruvate; this has been shown on beef heart lactate
dehydrogenase (^, = 0.015 mM at pH 6.7) (Novoa et al, 1959), yeast
D-lactate dehydrogenase (^, = 0.007) (Labeyrie and Stachiewicz, 1961),
yeast D-lactate cytochrome c reductase {K^ = 0.0016 mM) (Nygaard, 1961
b), and yeast D-hydroxy acid dehydrogenase (/if, = 0.0025 mM) (Boeri et
al., 1960), although the inhibition seems to be uncompetitive on the lactate
dehydrogenase oi Propionihacterium pentosaceum (Molinari and Lara, 1960).
The complex kinetics have been treated in detail by Novoa et al. (1959),
but it is still rather puzzling that oxalate inhibits all these enzymes so
436 2. ANALOGS OF ENZYME REACTION COMPONENTS
potently and competes with lactate rather than pyruvate. Tartronate and
malonate also inhibit competitively with lactate and noncompetitively with
respect to pyruvate, although by no means as strongly as oxalate, and
Ottolenghi and Denstedt (1958) concluded that pyruvate and lactate react
with different sites on the enzyme surface, but this is not necessary, as
Novoa et al. (1959) have shown, since the configurations of the active sites
on E-NAD and E-NADH are different. The five lactate dehydrogenase
(LD) isoenzymes from human tissues are inhibited to different degrees by
oxalate at 0.02 vaM (see accompanying tabulation) (Emerson et al., 1964).
Isoenzyme
0/
/o
Inhibition
LDi
70
LD^
64
LD3
56
LD4
46
LD,
32
There are A and B monomers, the B monomer being more sensitive to
oxalate. LD^ is a pure B tetramer, LD5 is a pure A tetramer, and the others
are intermediate in the proportion of A to B. This is an illuminating example
of how enzymes may respond differently to inhibitors by reason of varied
composition.
There are several interesting studies on lactate dehydrogenases from
which deductions on the nature of the active site and certain interaction
energies may be derived. Dikstein (1959) found that yeast lactate dehydro-
genase is not inhibited potently by monocarboxylates: the concentrations
for 50% inhibition are 6000 vaM for formate, 2300 mM for acetate, 800 milf
for propionate, 300 mM for butyrate, 120 mM for valerate, and 10 mM for
caprylate. By plotting log (1)50 against the number of carbon atoms in the
aliphatic chains he obtained a straight line, from the slope of which it was
possible to calculate that the transfer of a methylene group from the solvent
to the enzyme surface involves an over-all energy change of 0.5 kcal/mole.
It was assumed that the interaction energy between the COO" group and
an enzyme cationic group could be calculated from the intercept of this
line, but it is doubtful that at the high concentrations used the residual
energy can be attributed with certainty to ion-ion interactions, since non-
specific effects cannot be excluded.
The D-of-hydroxy acid dehydrogenase of yeast is competitively inhibited
by several dicarboxylates (see accompanying tabulation) (Cremona, 1964).
Monocarboxylates inhibit much more weakly and the inhibitions are usually
not purely competitive. Oxalate is bound approximately 3.7 kcal/mole more
LACTATE METABOLISM 437
tightly than malonate, suggesting that there are two cationic groups quite
close on the enzyme.
Inhibitor Ki (mil/)
Oxalate
0.0025
Tartronate
0.84
Malonate
0.95
L-Malate
1.05
a-Ketoglutarate
1.4
The D-lactate dehydrogenase of yeast studied by Boeri et al. (1960) is
inhibited potently by oxalate {K^ = 0.0025 mM), moderately by malonate
(^ . = 0.9 mM), and not at all by 16 mM succinate or fumarate, indicating
the importance of the position of the second C00~ group. The enzyme is
inactivated gradually by EDTA and reactivated with Zn++. Although this
does not prove that Zn++ is the normal metal ion involved, it points to the
possibility of chelation between a-hydroxy carboxylates, or dicarboxylates,
with an enzyme-bound metal ion. The configuration around the a-carbon
atom is important since L-lactate binds very weakly to the enzyme {K^ =
= 62 mM).
The lactate cytochrome c reductases of yeast are flavoproteins and are
competitively inhibited by a variety of lactate analogs (Nygaard, 1961 a,
b, c). Fatty acid inhibitions lead to the calculation of 0.37 kcal/mole for
the interaction of methylene groups with the enzyme and 2.80 kcal/mole
for the C00~ group in the case of the D-lactate cytochrome c reductase,
and of 0.37 kcal/mole and 0.88 kcal/mole, respectively, for the L-lactate
cytochrome c reductase. Neither eftzyme is inhibited by dicarboxylates,
with the exception of oxalate, and from the differences in inhibition patterns
it is probably safe to assume that the active sites of the two enzymes are
quite different.
Some analog inhibitors of lactate dehydrogenases need only be listed for
reference: pyruvate (Green and Brosteaux, 1936; Das, 1937 b; Neilands,
1952; Labeyrie and Stachiewicz, 1961), bromopyruvate, chloropyruvate,
hydroxypyruvate (Busch and Nair, 1957), phenylpyruvate (Dikstein, 1959),
a-ketoglutarate (Boeri et al., 1960), malate (Boeri et al., 1960; Busch and
Nair, 1957), tartronate (Green and Brosteaux, 1936; Lehmann, 1938), tar-
trate (Labeyrie and Stachiewicz, 1961), mandelate (Lehmann, 1938), a,
y-diketovalerate (Meister, 1950), benzenesuLfonate (Baptist and Vestling,
1957), mercaptoacetate, mercaptosuccinate, or-mercaptopropionate, of-mer-
captobutyrate, and a-mercaptovalerate (Chaffee and Bartlett, 1960). The
benzenesulfonates, where the COO" group is replaced by a SO3" group.
438 2. ANALOGS OF ENZYME REACTION COMPONENTS
and the mercapto fatty acids, where the a-OH is replaced by an a-SH
group, are particularly interesting and deserve further study to determine
their specificity on lactate metabolism.
The oxidation of glycolate is catalyzed by glycolate oxidase and the
CHO-CHO- + NADH + H+ ;fi CH^OH-COO- + NAD+
Glyoxylate Glycolate
reverse reaction by glyoxylate reductase, both enzymes being found in
plants. This reaction is similar to the pyruvate ±5 lactate interconversion
and, indeed, L-lactate is slowly oxidized by the oxidase. Since these enzymes
bear some relationship to those involved in lactate metabolism due to this
similarity, it is not inappropriate to discuss them at this point, particularly
as Zelitch at the Connecticut Agricultural Experiment Station has reported
some very interesting inhibitions by analogs. Glyoxylate reductase is ap-
parently not especially susceptible to analogs: the following inhibitions were
observed with 16.5 mM glyoxylate and 10 mM analogs — phenylglyoxy-
late 21%, oxalacetate 22%, oxamate 31%, and pyruvate 56% (Zelitch,
1955). Glycolate oxidase, on the other hand, is well inhibited by a-hydrox-
ysulfonates. Competitive inhibition was found with hydroxymethanesul-
fonate {K, = 0.0018 mM), a-hydroxyethanesulfonate (^,; = 0.0023 mM),
and sulfoglycolate {K^ = 0.0021 mM); since K,,^ = 0.38 mM, these analogs
are reasonably potent (Zelitch, 1957). Rabbit muscle lactate dehydrogenase
is inhibited comparably, D-glycerate dehydrogenase less strongly, and ma-
late dehydrogenase not at all, so that some specificity toward enzymes
oxidizing a-hydroxy acids is evident. The inhibition of lactate oxidation
by the glycolate oxidase is inhibited very strongly because lactate is bound
less tightly to the enzyme. Another competitive inhibitor of comparable
potency is of-hydroxy-2-pyridinemethanesulfonate (Zelitch, 1959) which
has been used in most of the in vivo work apparently because it is more
effective in cells, although sulfoglycolate w^ould seem to act very similarly
(Zelitch, 1958).
OH
I
CH3— CH— SO3 H0-CH2— so;
Q-Hydroxyethanesulfonate HydroxymethanesuKonate
9" ^N OH
I /^ \ I
'OOC — CH-SO; \\ A>— CH-SO3
Sulfoglycolate ■^^'^^^!'u''^'K .
pyridinemethanesulfonate
PHOSPHATASES 439
Normal tobacco leaves contain around 0.5-1 //mole/g wet weight glyco-
late and this level can be increased as much as 10-fold by placing them in
solutions containing the a-hydroxysuLfonates (Zelitch, 1959). Since glyco-
late is formed photosyntheticaUy, marked accumulation occurs only in the
light. The concentration of a-hydroxy-2-pyridinemethanesulfonate for max-
imal inhibition is near 10 m.M, indicating that penetration into the cells
is quite poor. At higher concentrations of inhibitor the glycolate level falls,
due presumably to inhibition of glycolate formation; in fact, even at 10 mM
there must be some inhibition since photosynthetic incorporation of C^Og
is inhibited about 33%. The pattern of C^* distribution is, however, more
markedly altered; in controls, glycolate-C^^ accounts for 5.5% of the labeling
but in inhibited leaves it is almost 50%. This is a good example of a spe-
cific analog inhibitor useful in studying the importance of an intermediate
in a complex metabolic pathway, and valuable information may result
from more detailed studies on photosynthesis.
PHOSPHATASES
These enzjines are commonly inhibited by the products of the hydroly-
sis. We have already noted the inhibitions of phospho-L-histidinol phos-
phatase, O-phosphoserine phosphatase (page 270), and glucose-6-phos-
phatase (page 442) by dephosphorylated products, and there are other
examples related to analog inhibition. Orthophosphate also frequently in-
hibits: The following may be cited as instances of well-marked inhibition
of different types of enzyme — calf intestinal phosphatase (Schmidt and
Thannhauser, 1943), mouse liver acid phosphatase (Macdonald, 1961),
mouse liver pyrophosphatase (Rafter, 1958), calf brain carbamyl and acyl
phosphatases (Grisolia et al., 1958), sweet potato phosphatase (Ito et al.,
1955), and E. coli alkaline phosphatase (Garen and Levinthal, 1960). An-
other class of inhibitor comprises the phosphates that are substrates. Thus
sweet potato phosphatase with phenyl phosphate as substrate is competi-
tively inhibited by /^-glycerophosphate {Kf = 2 mM), pyrophosphate (K, =
= 0.33 mM), metaphosphate (^, = 3.2 mM), and ATP {K^ = 0.67 mM),
all of which are also substrates (Ito et al., 1955). Likewise the E. coli
phosphatase with p-nitrophenyl phosphate as substrate is competitively
inhibited by uridine phosphate {K^ = 0.044 raM), guanosine phosphate
{K^ = 0.046 mM), /^-glycerophosphate {K^ = 0.05 vnM), glucose- 1 -phos-
phate {K^ = 0.063 mM), and adenosine-5'-phosphate (Z,; = 0.093 mM)
(Garen and Levinthal, 1960).
More interesting are the inhibitions by various anions that may be con-
sidered as analogs of either phosphate or the substrate phosphates. Thus
arsenate (Garen and Levinthal, 1960; Ito et al., 1955; Macdonald, 1961),
borate (Ito et al., 1955), and silicate (Umemura et al., 1961) inhibit various
440 2. ANALOGS OF ENZYME REACTION COMPONENTS
phosphatases about as potently as phosphate and probably combine with
the enzymes in a similar manner. Certain carboxylates (oxalate, malonate,
malate, citrate, glucarate, gluconate, lactate, and others) have been found
to be inhibitory, but most of these are not remarkably effective, probably
binding to enzyme cationic groups to various degrees or chelating with
metal ions. However, ( + )-tartrate* is a very potent inhibitor of certain
phosphatases, as first shown by Abul-Fadl and King (1949) and confirmed
by Anagnostopoulos (1953), and is so much more active than other anions
that the mechanism has been investigated in several excellent studies.
The phylogenetic relationships of ( + )-tartrate inhibition and the vari-
able susceptibilities of phosphatases from different tissues are quite inter-
esting. Abul-Fadl and King (1949) found that although prostatic acid phos-
phatase is very sensitive, no effects are exerted on the acid phosphatases of
erythrocytes or plasma, and Anagnostopoulos (1953) noted no inhibition
with the phosphatases of mustard, wheat germ, or Aspergillus. Kilsheimer
and Axelrod (1958) investigated the effects of ( + )-tartrate on the phospha-
tases from many sources and found that at 20 mM the inhibitions vary
from 0 to 93%. They concluded that animal phosphatases are more suscep-
tible than plant phosphatases, and that bacterial phosphatases are generally
resistant. It was suggested that ( + )-tartrate may be of taxonomic use in
those more primitive organisms where it is difficult to decide whether they
are plants or animals.
The earliest work demonstrated that the inhibition is stereospecific, nei-
ther ( — )-tartrate nor meso-tartrate exerting appreciable effects on prostatic
phosphatase, and this has been confirmed in all the recent studies. In the
series of phosphatases tested by Kilsheimer and Axelrod (1958) it was ob-
served that very few of the enzymes are affected by (— )-tartrate and
these are inhibited only slightly. The iiT/s were determined by London et
at. (1958) as 0.13 mM for ( + )-tartrate and 93 mlf for (-)-tartrate on
prostatic acid phosphatase. It is strange that they found meso-tartrate to
be an effective inhibitor {K^ = 0.4 mM) in contrast to all other work.
The acid phosphatase from Neurospora crassa is inhibited completely by
12 mM ( + )-tartrate but is unaffected by 50 mM ( — )-tartrate or meso-
tartrate (Kuo and Blumenthal, 1961). In all cases the inhibition by ( + )-
tartrate is competitive with respect to substrate. The inhibition is thus more
marked with /^-glycerophosphate as the substrate than with phenyl phos-
phate or p-nitrophenyl phosphate, since the former substrate is bound less
tightly (Nigam at al, 1959; Kuo and Blumenthal, 1961).
Tartrate and related inhibitors have been used to map the active site
of prostatic phosphatase (London et al., 1958). The K/s and estimated
* There has been confusion in the nomenclature of the tartrates and the dextro-
rotatory ( + )-tartrate has been designated as l- or D-, depending on the system used.
To avoid ambiguity I shall indicate the isomers by the signs of their rotation.
PHOSPHATASES 441
relative binding energies are shown in Table 2-25. It was assumed that ( + )-
tartrate is bound to the enzyme at four points; the two carboxylate groups
interact with two enzyme cationic groups and the two hydroxyl groups
Table 2-25
Competitive Anionic Inhibitors or Prostatic Phosphatase"
Inhibitor
Fluoride dimer (HF^
( + )-Tartrate
DL-Glycerate
meso-Tartrate
Arsenate
D-Malate
Sulfamate
Nitrate
D- Alanine
L-Leucine
Diphenylphosphate
D-Lactate
L-Serine
L-Glutamate
L-Aspartate
( — )-Tartrate
L- Lactate
L-Malate
L- Alanine
Glycine
Ki
Relative — AF oi binding
{mM)
(kcal/mole)
<0.1
>5.68
0.13
5.51
0.2
5.25
0.4
4.82
1
4.25
1.8"
3.90
8
2.98
16
2.55
27
2.23
30
2.16
32
2.12
50
1.85
50
1.85
60
1.74
64
1.69
93
1.47
110
1.36
280
0.79
470
0.47
470
0.47
" The substrate is /S-glycerophosphate {K,^ — 16 mM). Experiments with amino
acids at pH 7.2 and with the rest at pH 5. (From London et al., 1958.)
" Kj for D-malate estimated from value for DL-malate since L-malate is a weak in-
hibitor.
form hydrogen bonds with the enzyme (Fig. 2-14). On the basis of this
model it is possible to explain most of the variations in inhibitor binding.
( — )-Tartrate can make contact at only two points while meso-tartrate can
make a three-point attachment in two ways. Only D-malate can attach at
three points like ( + ) -tartrate but the second hydroxyl group is missing.
The most important requirement for binding is a negative group separated
from the nucleophilic group by around 2.9 A. (-f )-Tartrate has two such
units. In the substrates the oxygen atom of phosphate is the nucleophilic
atom. The amino acids are rather poor inhibitors, probably because at
442
2. ANALOGS OF ENZYME REACTION COMPONENTS
pH 7.2 most of the amino groups are protonated and interfere with the
binding. HFj" may bind across two attachment points and its potency may
be related to the strong hydrogen bonds formed by fluorine. The nitrate
Fig. 2-14. Scheme for the active site of
prostatic acid phosphatase showing two
cationic groups and two hydrogen -bonding
groups separated by a seam in the enzyme.
The unmarked circles represent various
enzyme groups and the stippled area
between them a lipophilic region. (Modi-
fied from London et al., 1958.)
ion also can bridge two of the attachment points, one oxygen atom being
negatively charged and the other nucleophilic.
Some of the results of Nigam et al. (1959), shown in the accompanying
tabulation, differ from those of London et al. (1958). These inhibitions were
Inhibitor (10 mikf )
% Inhibition
( -f )-Tartrate
100
Glucarate
91
Pyruvate
80
Oxalate
72
Malonate
66
Maleate
45
Glutamate
42
Glucuronate
40
Lactate
0
SULFATASES 443
observed with 3 mM /^-glycerophosphate as substrate. The potency of (-(-)-
tartrate is not evident in the tabulation since 50% inhibition is produced
by 0.07 mM. The failure of lactate to inhibit is surprising, especially as
pyruvate is fairly potent. The susceptibility to glucarate observed here has
not been observed with Neurospora phosphatase, 12.5 mM having no effect
(Kuo and Blumenthal, 1961 ), and Jeffree (1957) found only a moderate inhi-
bition {K^= 10 mM) on prostatic phosphatase, it being definitely less potent
than oxalate in contrast to the results above. Whether the lactone plays a role
in this inhibition is not known. Two anionic polymers have been found to
be more potent inhibitors than (-f) -tartrate: polyxenyl phosphate inhibits
the Neurospora phosphatase completely at 0.16 mM (Kuo and Blumenthal,
1961), and alginate (556 residues) inhibits the prostatic enzyme with a K^
of 0.0054 mM (Jeffree, 1957). These polymer inhibitions are only partially
competitive.
SULFATASES
Arylsulfatases of type II (liver enzymes A and B and molluscan enzymes)
are generally inhibited by sulfate and phosphate, but type I arylsulfatases
(liver enzyme C and bacterial enzymes) are resistant. The product inhibi-
tion by sulfate was first reported by Tanaka (1938) and shown to be com-
petitive with the substrate nitrocatechol sulfate by Roy (1953). The sus-
ceptibilities of ox liver arylsulfatases to sulfate vary markedly: The K^ for
sulfatase A is 0.75 mM, for sulfatase B is 70 mM, and sulfatase C is not
affected (Roy, 1953, 1954 b, 1956). The sulfatase A is inhibited also by
several organic sulfates, although not very potently (see accompanying tab-
ulation). The following do not inhibit at 25-50 mM: methyl sulfate, glu-
cose-3-suLfate, glucose-6-sulfate, and phenyl sulfite. The fact that methyl
sulfate does not inhibit and the affinity for the enzyme increases with size
of the ring system esterified might indicate that the sulfate group is not
Inhibitor
{mM)
Relative — AF o{ binding
(kcal/mole)
Sulfite
0.002
8.07
Phosphate
<0.21
>5.21
Sulfate
0.75
4.42
2-Phenanthryl sulfate
A
3.40
Phenyl phosphate
.^.3
3.23
2-Naphthyl sulfate /
30
2.15
1-Naphthyl sulfate
40
1.98
m-Tolyl sulfate
100
1.42
Phenyl sulfate
300
0.74
Benzyl sulfate
300
0.74
444 2. ANALOGS OF ENZYME REACTION COMPONENTS
involved in the binding of these substances. The inhibition by these organic
sulfates was stated to be noncompetitive but since some are slowly hydro-
lyzed substrates, the inhibition may be mixed. Phosphate seems generally
to be a more potent inhibitor than sulfate; this has been observed on the
arylsulfatases of rabbit liver (Maengwyn-Davies and Friedenwald, 1954),
ox liver (Webb and Morrow, 1959), Helix pomatia (Dodgson and Powell,
1959), Charonia lampas (Takahashi, 1960 a), and beef and rabbit cornea
(Wortman, 1962). The inhibitions, where examined, are competitive. Weak
inhibitions by xylenesulfonate (Maengwyn-Davies and Friedenwald, 1954)
and benzenesulfonate (Dodgson et al., 1955; Dodgson and Powell, 1959)
have been observed.
The potent inhibition by sulfite is interesting but the mechanism is not
yet understood, although it would appear to be competitive (Roy, 1953).
For all the ox liver arylsulfatases, sulfite is bound 3-4 kcal/mole more tightly
than is sulfate; indeed, sulfatase C, although resistant to sulfate, is inhibited
55% by 0.1 mM sulfite (Roy, 1956). Sulfite is also more inhibitory than
sulfate on sulfatases from bacteria and molluscs (Dodgson et al., 1955; Dodg-
son and Powell, 1959; Dodgson, 1959), although in most cases the concen-
trations used were too high to evaluate the potency readily.
Choline sulfatase of Pseitdomonas nitroredvcences is inhibited very strongly
by sulfite (33% by 0.01 mM and 83% by 0.1 mM when choline sulfate is
10 mM), but is not affected or somewhat stimulated by sulfate and phos-
phate (Takebe, 1961). The steroid sulfatase and glucosulfatase of Patella
vulgata are inhibited by sulfate and even more potently by phosphate
(Roy, 1954 a). Since the sulfatases comprise so heterogeneous a group of
enzymes and relatively few have been adequately studied, it is difficult to
draw general conclusions or make valid correlations, but it is at least evident
that analogs are occasionally effective inhibitors. It is also likely that phos-
phate must exert a regulatory action on sulfatase activity in vivo.
ADENOSINETRIPHOSPHATASES
AND TRANSPHOSPHORYLASES
Enzymes hydrolyzing ATP or transferring its terminal phosphate to
various acceptors are frequently inhibited by other nucleotides. Competi-
tive product inhibition by ADP has been noted for ATPases from several
sources; the inhibition is never marked, since ADP is usually bound some-
what less tightly than ATP to the enzyme, but is sufficient to slow progres-
sively the rate of ATPase reactions. The ATP-P^ and ATP- ADP exchange
reactions catalyzed by mitochondria and digitonin particles are also inhib-
ited by ADP (Low et al., 1958; Cooper and Kulka, 1961). Some results with
ADP and other related substances are shown in Table 2-26. The inhibitions
however, depend on the pH and the concentrations of Ca++ and Mg++, as
ADENOSINETRIPHOSPHATASES , TRANSPHOSPHORYLASES
445
Table 2-26
Inhibition of ATPases by Analogs
ATP
Analog
Source , , .
centration
Analog
con-
centration
0/
T ,.,.,. Reference
Inhibition
(mM)
(milf)
Liver
4
ADP
2
23
Kielley and KieUey
mitochondria
4
42
(1953)
IDP
2.2
4
0
16
ITP
4
13
15
AMP
15
22
Low et al. (1958)
ADP
15
45
3.5
ADP
3.5
50
Cooper and Kulka
(1961)
Muscle myosin
1.07
ADP
0.4
2.4
7
12
Blum (1955)
0.21
ADP
2.4
29
Spinach
4
AMP
4
37
VVessels and
chloroplasts
ADP
4
29
Baltscheffsky (1960)
Brain
—
Adenine
5
23
Gore (1951)
Adenosine
20
16
Guanine
5
2
shown by Green and Mommaerts (1954). Addition of Ca++ decreases ADP
binding at pH 6.4 and increases it at pH 9, whereas Mg++ has no effect at
the lower pH but decreases affinity at the higher pH. The K^ for ADP is
around 0.13 roM in the absence of Ca++ and Mg++, but around 0.5 mM
at pH 6.4 and 40 mM Ca++. Kielley and Kielley (1953) had shown with
liver mitochondrial ATPase that ADP does not alter the optimal Mg++
concentration for ATPase activity, indicating that the inhibition is not by
the binding of Mg"*""*", but Nanninga (1958) reported that part of the inhi-
bition of myosin ATPase by ADP is due to chelation of Ca++. In the pre-
sence of excess Ca++ this chelation can be neglected and the true K^ for
the enzyme- ADP complex is found to be 4.6 mM at pH 7.
An interesting phosphonic analog of ATP, adenylmethylenediphospho-
nate:
0 0 0
II II II
Adenine-ribose— O— P— O— P— CH— P— O-
O- O- OH
446 2. ANALOGS OF ENZYME REACTION COMPONENTS
was found by Moos et al. (1960) to be unable to replace ATP in contracting
glycerinated muscle and not to be hydrolyzed by myosin ATPase. However,
some inhibition on ATPase is exerted (the affinities of the enzyme for ATP
and its analog appear to be roughly the same), although this is not com-
petitive. Mg++ is able to overcome the inhibition at a concentration lower
than that of the analog, indicating that Mg++ is not simply complexing
with and removing free analog. The mechanism of the inhibition was re-
presented by the following reactions:
EMg
Mg
+
E
+
+
I
I
w
w
Mgl
EI
where E is low-activity enzyme and EMg is high-activity enzyme. The inhi-
bition has a dual basis: (1) removal of Mg++, thus decreasing the fraction
of the enzyme in the high-activity form, and (2) reaction of the low-activity
enzyme directly with the analog. This situation may be fairly common in
inhibitions on enzymes with activating metal ions.
A few other nucleotidase inhibitions may be mentioned. ITPase is inhib-
ited by IDP and ADP (Blum, 1955; Kielley and Kielley, 1953). Indeed,
ADP inhibits ITPase more strongly than ATPase. The ITPase of fly muscle
is strongly inhibited by ADP (K^ = 0.0165 mM) and much less readily by
IDP (Kj = 1.59 mM), the inhibition being competitive at low but noncom-
petitive at higher concentrations (Sacktor and Cochran, 1957). GTPase is
likewise inhibited but UTPase is unaffected by either ADP or IDP. In
phage-infected E. coli the hydrolysis of deoxycytidine diphosphate (deoxy-
CDP) is inhibited by deoxyCMP and deoxyCTP, and the hydrolysis of
deoxyCTP is inhibited by deoxyCMP and deoxyCDP, in both cases the
deoxyCMP being relatively less active (Zimmerman and Romberg, 1961).
AMP- ATP transphosphorylase (myokinase) from rabbit muscle is inhi-
bited by ADP {K^ = 0.33 mM) and this is competitive with respect to
both AMP and ATP (Noda, 1958). The reverse reaction from 2ADP->AMP
-f ATP is inhibited by AMP {K, = 0.5 mM) and ATP (A', = 0.32 mM),
the K/s being the same as the K„^s for these substances (Callaghan and
Weber, 1959). A much more effective analog is adenosine monosulfate
{K^ = 0.0186 mM). Creatine kinase is inhibited competitively by ADP
(Ki = 0.27 mM), AMP {K^ = 7 mM), adenosine (Z, = 7 mM), tripoly-
phosphate {K, = 8 mM), orthophosphate (A, = 13 mM), sulfate (A, =
6 mM), and nitrate {K^ = 22 mM) (Noda et al, 1960). The substrate here
is MgATP= and it is possible that the most effective inhibitors form Mg
complexes. Most of the anions inhibit the forward reaction competitively
with respect to MgATP= and the reverse reaction competitively with re-
HYDROXYSTEROID DEHYDROGENASES
447
spect to creatine phosphate (Nihei et al., 1961). However, ADP competes
with MgADP" in the reverse reaction.
Since oxidative phosphorylation may in some ways be related to ATPase
activity and transphosphorylations, it is not out of place to discuss the ef-
fects of various inorganic phosphorus compounds on this process. The phos-
phorylation in a rat liver mitochondrial suspension oxidizing fumarate, and
with glucose and hexokinase to trap the phosphate, was studied by Thom-
son and Sato (1960) (Table 2-27). Some of the analogs investigated reduce
the P : 0 ratio by depressing phosphate uptake more than oxygen uptake,
but the only compound that can be considered as a true uncoupler is thio-
phosphate. In such a complex system a number of sites for inhibition are
evident. It is known that anions can inhibit fumarase and it is possible that
other enzymes attacking dicarboxylates might be inhibited. Hexokinase is
inhibited at concentrations interfering with oxidative phosphorylation only
by triphosphate. Some of these compounds might deplete Mg++ but it was
shown that thiophosphate does not. It is not known if any of these sub-
stances can enter into the phosphorylative reaction but fail to form ATP.
Further study of thiophosphate would seem warranted by these preliminary
results.
HYDROXYSTEROID DEHYDROGENASES
The /?-hydroxysteroid dehydrogenase of Pseudomonas testosteroni cata-
lyzes the oxidations of 3/?- and 17/?-hydroxysteroids to their respective ke-
tones with NAD as acceptor. The oxidations of testosterone and 17/5-estra-
diol are competitively inhibited by 17cif-estradiol (Talalay and Dobson, 1953).
H,C
H,C
OH
Androstane
Testosterone
Androst-l-ene-3, 17-dione
HO
Estra-l, 3, 5-triene
Estrone
Progesterone
448 2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-27
Effects of Inorganic Puosi'HORUs Compounds on Oxidative Phosphorylation "
Structu
re
Concen-
tration
(mA/)
7,
Change
Inhibitor
O3
uptake
Pi
uptake
P:0
Pyrophosphate
-O-P-O-P-
o o
O'
1.5
3
- 22
- 20
- 36
- 51
- 16
- 38
Triphosphate
o" o'
O—P-O— I'-
ll II
o o
o' o'
1 1
0-
0
1
-P-O
II
0
0.6
3
+ 8
- 30
- 11
- 48
- 17
- 25
Pyrophosphite
"O-P-O-P-
1 1
H H
o'
1
- 4
- 4
0
Diphosphite
O' O'
1 1
0-P-P=0
II 1
0 H
?' ?'
3
15
- 12
- 11
- 4
- 18
f 10
- 16
Hypophosphate
1 1
O-P— P-O
a 5
1
- 25
- 38
- 20
Isohypophos-
phate
'O— P— O— P-
II II
o o
-H
3
15
t^ 10
- 25
- 4
- 15
Trimetaphos-
phimate
O^ /O'
p
o^ 1 1 /O'
1
H
o' o'
0'
3
+ 12
+ 8
- 4
Diimidotri-
phosphate
1 1
"O— P— N— P-
Tl 1 II
OHO
-t
t
-P-O
11
0
3
- 9
- 24
- 19
0.6
- 4
- 2
+ 2
Thiophosphate
PSO33-
3
5
+ 18
+ 26
0
- 28
- 15
- 42
15
- 59
- 91
- 78
"Rat liver mitochondria oxidizing fumarate with phosphate at 15 mAf. (From Thomson and
Sato, 1960.)
HYDROXYSTEROID DEHYDROGENASES 449
This observation was extended to a number of estra-l,3,5-trienes, most of
which are inhibitory to the oxidation of testosterone and are not themselves
attacked (Marcus and Talalay, 1955). The most potent inhibitors are the
following derivatives of estratriene: -3,17a-diol (a-estradiol), -3,16«-diol;
-3,16a,17/5-triol (estriol); -3-ol; -3,17/?-diol-16-one; and -3,16/3-diol. It ap-
pears that the aromaticity of ring A combined with the 3-OH group re-
sults in strong binding. The 17-ols are not inhibitory although there are
enzyme regions for the oxidation of either 3-OH or 17-OH groups in other
ring systems. The total ring system is not necessary since dieth/lstilbestrol
and hexestrol are potent inhibitors. The K/s for most of the effective in-
hibitors are around 0.001-0.01 mM corresponding to over-all interaction
energies of 7-8.5 kcal/mole, implying a rather close fit over the surface of
the molecules and a summation of dispersion and polarization forces. It is
possible that the high polarizability of the aromatic ring A is important in
augmenting binding, this ring overlying some ionic group on the enzyme,
and the 3-OH interacts to form a hydrogen bond. The a,^ specificity indi-
cates that the steroids attach to the enzyme by their "rear" surfaces. The
cf-hydroxysteroid dehydrogenase is not inhibited so readily by the estra-
trienes as is the (3 enzyme (Talalay and Marcus, 1956). It was stated that
the inhibitions are neither exactly competitive nor noncompetitive, but no
data or plots were given, nor was the exact experimental procedure des-
cribed, so that it is impossible evaluate the nature of the inhibitions.
The most potent inhibitor of the Pseudomonas /?-hydroxysteroid dehydro-
genase yet found is the noncompetitive 2-hydroxymethylene-17a-methylan-
drostan-17/5-ol-3-one {K, = 0.0003 mM), although the competitive 4,4-di-
methyl-17/?-hydroxyandrost-5-eno(3,2-c)pyrazole {K^ = 0.0005 mM) is al-
most as active (Ferrari and Arnold, 1963 a, b). The inhibitions by these and
simpler steroids are dependent on the pH; e.g.. diethylstilbestrol is approxi-
mately 20 times as effective at pH 8.5 than at pH 5.5. Since it is unlikely
that the phenolic groups would ionize in this range (pK^ for diethylstilbes-
trol is 12.2), this implies the ionization of enzyme groups at or near the
active site. This emphasizes the importance of polarization of the aromatic
rings by anionic groups on the enzyme in determining the tightness of
binding.
The J^- and J*-dehydrogenases which introduce unsaturation into ring
A of the 3-ketosteroids at the 1- and 4-positions, respectively, are inducible
enzymes in P. testosteroni, and both are inhibited quite potently by estrone
(Levy and Talalay, 1959). The J*-3-ketosteroid reductase (5a) of rat liver
microsomes, which catalyzes the hydrogenation of the 4-5 double bond, is
a NADPH-requiring enzyme acting on cortisone, Cortisol, desoxycorticoste-
rone, and related steroids (McGuire et al., 1960). It is competitively inhi-
bited by a variety of less substituted steroids, such as androst-l-ene-3,17-
dione and 5a-androstane-3,17-dione; the 5/?-androstane-3,17-dione is, how-
450
2. ANALOGS OF ENZYME REACTION COMPONENTS
ever, inactive, indicating very specific attachment to the enzyme. The K^
for 17/5-hydroxyandrosta-l,4-diene-3-one is 0.146 mM when the substrate
is cortisone (iiC,„ = 0.14 xaM), and the other inhibitors presumably have
similar values for K^. It may also be noted that 6of- and 6y5-methyl-ll-keto-
progesterone are inhibitors.
The normal roles of these enzymes in bacteria or mammals are not as
yet well understood, although the steroid dehydrogenases in Pseudomonas
enable this organism to grow with the appropriate steroids as the sole
source of carbon. There are many other enzymes involved in steroid syn-
thesis and catabolism which have not been studied with respect to analog
inhibition. When such studies are made it may well be found that this is
an important regulating mechanism in controlling the levels of the steroid
hormones.
NITRITE AND SULFITE METABOLISM
Nitrobacter is a soil autotroph that can obtain its energy from the oxida-
tion of nitrite to nitrate. Chlorate at low concentrations (0.01-0.1 mM)
inhibits growth without affecting nitrite oxidation, whereas at higher con-
centrations (above 1 mM) nitrite oxidation is progressively depressed (Lees
and Simpson, 1955). The inhibition is completely reversible and it was pos-
tulated that chlorate combines with a nitrite-oxidizing enzyme at a stage
when it is bound to nitrate; however, the inhibition seems to be unrelated
to the concentrations of either nitrite or nitrate, and is not reversed by
increasing the nitrite concentration (Lees and Simpson, 1957). It was then
postulated that chlorate does not inhibit directly but is first converted to
some substance, perhaps chlorite, which inhibits rapidly and irreversibly.
Nitrite
o
0"
11-
1 V \- ■
0/ V
'o-ci^—
Nitrate
Chlorate
Chlorite
Inhibitor (2 mM)
0/
/o
Inhibition
CIO3-
80
BrOs
8
IO3-
1
PO3-
0
o
N=C=0
'N— C=0^
'^N=C— O'
Cyanate
Related anions do not inhibit comparably with chlorate (see tabulation,
where nitrite is 8mM), but cyanate is apparently even more inhibitory
NITRITE AND SULFITE METABOLISM 451
(Butt and Lees, 1960). However, the effect of cyanate is markedly depen-
dent on the oxygen concentration (see following tabulation), so that at
low oxygen levels inhibition is reversed to stimulation. It was suggested
Cyanate
%
Change
(mikf)
0^ = 2.5%
O2 = 20%
0.2
+30
-50
0.3
+45
-66
0.5
+ 18
-76
0.7
- 3
-82
1.0
-46
-92
that a membrane transport system brings nitrite into the cells, and at mod-
erate nitrite concentrations and normal oxygen pressures the rates of
transport and oxidation are comparable. At low oxygen concentrations
there is accumulation of nitrite in the cells with consequent inhibition of the
nitrite-oxidizing enzyme (since substrate inhibition occurs when nitrite is
above 1 vaM). Cyanate may interfere with the transport so that at normal
oxygen concentrations the accumulation of nitrite is suppressed and the
substrate inhibition released. It would be interesting to know what effect,
if any, chlorate has on the nitrite transport.
The oxidation of sulfite by the liver is inhibited by thiosulfate and this
action is probably exerted on sulfite oxidase (Fridovich and Handler, 1954,
1956). Thiosulfate inhibits both aerobically and anaerobically (methylene
blue reduction). Although it was originally stated that the inhibition is
competitive, it has been found more recently (MacLeod et al., 1961) that
it is not, at least with respect to sulfite. The following reaction scheme was
suggested:
-S
-S + HSO3
-S— H
-S— H
(+ H2O)
-s— SO3- > h-s— H + SOr + H^
and it was proposed that thiosulfate competes with the active thiosulfonate
in the hydrolytic step of the sequence. Methanesulfonate, ethanesulfonate,
benzenesulfonate, and pyridine-3-sulfonate are noninhibitory.
Wild-type Neurospora crassa can utilize sulfate as the sole source of sulfur
but certain mutants require more reduced forms for growth (Ragland and
Liverman, 1958). Some strains can use sulfite and others only thiosulfate.
Since sulfate inhibits competitively the utilization of thiosulfate but does
not interfere with the utilization of sulfite, and inasmuch as previously it
452 2. ANALOGS OF ENZYME REACTION COMPONENTS
had been postulated that sulfite must pass through thiosulfate to be utiliz-
ed, a shunt around thiosulfate was proposed:
I I
Active sulfate -> active sulfite -> active thiosulfate -► growth
t t t
sulfate sulfite thiosulfate
Sulfate would thus block the formation of active thiosulfate from exogenous
thiosulfate, or it could block the utilization of active thiosulfate. Another
possibility is that thiosulfate transport into the cell is depressed by sulfate,
in which case there is no necessity to assume a shunt as above.
SIMPLE ION ANTAGONISMS
Many examples of the inhibition of biological, metabolic, or enzymic
processes by simple inorganic ions are known and in certain instances it is
likely that the mechanism involves a competition between the inhibiting
ion and a necessary ionic cofactor. The inhibiting ions may be considered
as analogs of the normally functional ions. It is impossible here to treat
this subject completely but it may be worthwhile to mention briefly some
specific examples of competitive enzyme inhibition to illustrate the phe-
nomenon. MacLeod and Snell (1950) emphasized the possible importance
of such competitions in the effects of certain ions on the growth of bacteria.
The growth of Lactobacillus arabinosus is suppressed by various ions and it
was concluded that some of these effects are due to the fact that the inhib-
itory ions are structural analogs of the metal ions which are cofactors in
metabolism. The following antagonisms, among others, were demonstrated:
K+ reverses the inhibitory effects of Na+ and NH4+; K+ reverses the in-
hibitory effects of Rb+; Zn++ reverses the inhibitory effects of Mn++; and
Mg++, Ca++, and Sr++ reverse the inhibitory effects of Zn++. Actually it is
not certain that these antagonisms relate to metabolic events, since it is
possible that physiological ions play nonmetabolic roles in bacteria, and it
would be interesting to investigate these antagonisms on a metabohc level.
Inasmuch as so many enzymes either involve ionic cofactors or are affected
by physiological ions, it is likely that such competitions are very common
and important in metabolic regulation.
The phosphotransacetylase of Clostridium kluyveri is active only in the
presence of K+ or NH4+ ions. Inhibition is exerted by Na+ and Li+ ions,
and this can be overcome by increasing the K+ or NH4+ concentration, so
that competition for an enzyme site would seem likely (Stadtman, 1955).
Yeast acid phosphatase requires Mg++ and is inhibited by Ca++, the inhi-
bition being strictly competitive, as shown by double reciprocal plots (Tsuboi
and Hudson, 1956). Ba++ does not inhibit at all, but Mn++ and Zn++ depress
INHIBITION BY MACROIONS 453
the activity, although less than Ca++, and this may be competitive also.
Ca++ is a competitive inhibitor of rabbit muscle phosphorylase 6 kinase
with respect to the activator Mg++, but is noncompetitive with respect to
ATP {K,„ = 1.9 mM for Mg++, and K^ = 0.3 vaM for Ca++) (Krebs et al,
1959). Most ATPase are activated by Mg++ and some are inhibited by Ca++,
but L-myosin ATPase is activated by Ca++ and inhibited by Mg++; in both
cases competition may occur. One might speculate that the uncoupling of
oxidative phosphorylation by Ca++ may involve displacement of other ions
such as Mg++ or Mn++. The question of metal ion competition will be taken
up in greater detail in the chapters on inhibitions produced by Zn++, Pb++,
Cd++, and other metal cations. Some new ideas on the regulatory role of
simple cations in metabolism, especially glycolysis, may be found in the
interesting discussion of Wyatt (1964). One example of ion inhibition in an
enzyme system in which an ion is the substrate will be mentioned. This is
the formation of ^-chlorolevulinate from /5-ketoadipate by an enzyme, /?-
ketoadipate chlorinase, from Caldariomyces fumago (Shaw and Hager, 1959).
F~, Br-, and I" inhibit this reaction around 85% when they are present in
equimolar concentration with Ch (10 mM). The nature of the inhibition
is not known but it is rather surprising that the three inhibiting ions are of
the same degree of potency.
INHIBITION BY MACROIONS
The inhibition by high molecular weight substances of enzymes attacking
related high molecular weight substrates may be thought of as analog inhi-
bition in certain instances, especially where competitive kinetics has been
demonstrated (the type of inhibition has seldom been studied in work on
polymers). On the other hand, these inhibitions are probably often nonspe-
cific in the sense that the polymers interact with any or all regions of the
enzyme surface rather than just the active center. Spensley and Rogers
(1954) reviewed inhibitions of this type and suggested the terms macrocat-
ionic and macroanionic as applicable to effects exerted by positively charged
and negatively charged polymers, respectively. Early work was mainly with
naturally occurring macroions, such as heparin or protamine, but the pre-
paration and use of more homogeneous and physically characterized syn-
thetic poljoners and copolymers have enabled the nature of the interactions
to be better understood. Some of these inhibitions may well be physiologi-
cally important, for example, the intracellular effects of the various types of
nucleic acids on enzymes involved in nucleic acid metabolism or their role
in protein synthesis. The suggestion by Jones and Gutfreund (1964), that
certain enzymes participating in metabolic sequences may interact speci-
fically with each other to form complexes facilitating flow along the path-
way, brings up the possibility that macroions can interfere in this interaction
454 2. ANALOGS OF ENZYME REACTION COMPONENTS
and thereby depress metabolism by a mechanism unrelated to direct action
on the individual enzymes. General macroion inhibition will be briefly dis-
cussed in this section, whether specific or nonspecific, and that these are all
instances of analog inhibition is not implied. Bernfeld (1963) has recently
reviewed some aspects of macroanionic inhibition.
General Nature of the Interactions between Macroions
Inasmuch as enzymes are macroions, the interactions involved, whether
specific or nonspecific, will be mainly electrostatic and will depend primarily
on the total charges and the distributions of charges on the enzymes and
the inhibitory polymers. It is likely that enzymes at pH's removed from
their isoelectric points will interact to varying extents with macroions of
opposite net charge irrespective of whether the enzymes attack high mo-
lecular weight or low molecular weight substrates, but we shall direct our
attention chiefly to enzymes whose substrates are polymeric. When inhi-
bition occurs under conditions in which the enzyme and the macroion are
carrying the same net charge, it has often been assumed that other than
electrostatic interactions are involved, but this is not necessary. Let us
picture an enzyme of isoelectric point pH 7 in a reaction medium of pH 6.5.
The net charge on the enzyme will be positive, but interspersed with the
cationic groups on the enzyme surface there will be many anionic carboxy-
late groups in most instances. If a macrocationic substance can orient itself
on the enzyme surface so as to react with these anionic groups, perhaps
because of complementary spacing of the oppositely charged groups, inhi-
bition may result, although it will undoubtedly be less than at pH's above
the isoelectric point. This is not said to minimize the importance of nonelec-
trostatic interactions, which certainly must occur, particularly when the
inhibitory macroions contain groups capable of forming hydrogen bonds or
regions contributing to the binding through van der Waals' forces.
The question of the specificity of macroionic inhibition is a difficult one
and the data available do not allow us to draw general conclusions. It has
been established, however, that a particular enzyme will be inhibited quite
differently by various macroions of the opposite net charge, and that a
particular macroion exerts very different effects on a group of enzymes.
There is probably sufficient evidence, to be discussed later, that enzymes
acting on macroionic substrates are likely to be inhibited rather strongly
by other macroions in which the charge distribution is similar to that of
the substrate, and in such cases the inhibition may indeed be competitive.
A quantitative treatment of the interactions must be on a statistical basis
and, as far as I know, this has not been undertaken, and would indeed be
very difficult since there are several factors about which there is inadequate
information.
The rate at which macroionic inhibition develops has not been studied
INHIBITION BY MACROIONS 455
extensively but one can imagine some interesting phenomena in this con-
nection. The initial binding of the macroion to the enzyme would not be
expected to be that for maximal interaction because the first contact be-
tween them would be random. Inasmuch as most macroionic inhibitions are
readily reversible, it is likely that the polymer would move on the enzyme
surface until near maximal or maximal interaction occurs. The inhibition
for this reason might increase with time, the rate depending on several
factors, such as the binding affinity of the individual group interactions
and the flexibility of the polymer, and progressive developments of inhi-
bition have been experimentally observed. Inasmuch as many configura-
tions of the macroion on the enzyme surface may be characterized by inter-
action energies very near the maximal, it is likely that even at equilibrium
each enzyme molecule will not be inhibited to the same degree, especially
for those enzymes acting on small substrates, since the inhibition will usual-
ly be due to a steric interference by a polymer chain passing over or near
an active site.
Factors Determining the Degree of Inhibition
The most important factors relating to the inhibitory macroion would be
(1) the molecular weight or polymer length, (2) the density of ionic groups
on the polymer, or the repeat distance between them, (3) the over-all con-
figuration of the polymer, i.e., linear, branched, or globular, and (4) the
flexibility of the polymer. The last factor is perhaps very important but
has been generally ignored. If one assumes an approximately globular en-
zyme, the net binding energy and the degree of inhibition may well depend
on the ability of the polymer to conform to the enzyme surface, specifically
to wrap around it so that interactions between many ionic groups can take
place. Some of the ionic polysaccharides must not be too flexible and this
may limit the effects they have on certain enzjTnes, while the synthetic
macroions vary in flexibility over a wide range. Entropy factors must be
very significant in the binding of macroions, and would to a great extent
depend on the deviation of the bound polymer from its statistical configu-
ration in solution. A polysaccharide or polypeptide macroion of molecular
weight 10,000 would contain roughly 35 units and the total extended length
would be 200-250 A if linear. Such a macroion might be able to encircle
an average enzyme 1 or 2 times or, if it is randomly distributed over the
enzyme surface, would cover very roughly about 10-15% of the enzyme.
Many macroions used to inhibit enzymes are, of course, much larger, often
being of molecular weights of 100,000 or over.
Two characteristics of the media used in inhibition studies are particu-
larly important, namely, the pH and the ionic strength, since the interac-
tions between enzyme and macroion are mainly of the ion- ion type. The
pH will determine the net charge on the enzyme and occasionally the ioni-
456 2, ANALOGS or enzyme reaction components
zation state of groups with which the inhibitor reacts, and in all cases in
which this has been studied a marked dependence on the pH has been
demonstrated. An increase in the ionic strength should reduce such inhibi-
tions because of the competition of the small ions for the enzyme and ma-
croion groups, and this has been repeatedly confirmed experimentally (data
for the inhibition of trypsin by polyglutamate are given in Table 1-15-6).
Hydration of the ionic groups must also be a significant factor in reducing
the inhibitions from what might be expected on the basis of interactions
in a vacuum, so that anything which modified the extent of hydration of
either enzyme or macroion might secondarily affect the inhibition. A final
factor which can markedly reduce such inhibitions is the presence of ma-
croionic impurities in the preparation if one is not working with pure en-
zymes. It has been shown many times that inhibitions by macroions can
be prevented or actually reversed by other macroions of opposite charge to
the inhibitor. Such results are not particularly significant since they imply
only that the inhibitor can also bind to nonenzyme macroions, a fact which
can be better demonstrated with other techniques, but they emphasize the
possible importance of such impurities in the studies on macroionic inhi-
bitions.
Trypsin and Chymotrypsin
The isoelectric point of trypsin is close to pH 11 and that of casein is
between 4 and 4.5; thus the hydrolysis of casein involves the interaction
of a macrocationic enzyme with a macroanionic substrate at pH values near
neutrality. Since heparin, a strongly negatively charged sulfated polysac-
charide, was known to form complexes with positively charged proteins,
Horwitt (1940) examined its action on trypsin and found a rather potent
inhibition at pH 7.3. Inhibition does not occur unless the enzyme is incubat-
ed with heparin before the addition of the casein, possibly indicating a
competitive type of interaction. Acidification to pH 3 leads to a dissociation
of the trypsin-heparin complex with restoration of full activity. The pH^pt
for trypsin is possibly shifted from 8.4 to lower values by heparin (Glazko
and Ferguson, 1940); it is not known if this means that enzyme combined
with heparin can act on casein — it is difficult enough to understand the
pHopt of proteolytic enzymes in the absence of inhibitors. The distance
between sulfate groups in heparin is 10.2 A, which is approximately equiv-
alent to 3 peptide residues in proteins, so it was suggested by Kornguth
and Stahmann (1960) that heparin may bridge the active site by combining
with cationic groups on either side. The active site appears to be covered,
since the hydrolysis of benzoylarginamide by trypsin is inhibited. Poly-or-
L-glutamate and polycysteate also inhibit trypsin, but poly-y-D-glutamate
does not, and this is probably correlated with the different distances between
C00~ groups in these macroanions. Poly-D-lysine inhibits the tryptic hy-
drolysis of poly-L-lysine, equimolar concentrations giving complete inhi-
INHIBITION BY MACROIONS 457
bition, showing the importance of the configuration of the polypeptide
chains (Tsuyuki et al., 1956). Polyacrylate at concentrations around 0.25-
0.5 mg/ml inhibits trypsin at high pH and accelerates catalysis at low pH
(Morawetz and Sage, 1955). Denatured hemoglobin, the substrate, has an
isoelectric point around 7.8, so that above pH 7.8 the polyacrylate can
combine only with the positively charged trypsin (the trypsin-polyacrylate
complex has some activity), whereas at lower pH's polyacrylate also binds
to the hemoglobin. Since the hemoglobin-polyacrylate complex is more sus-
ceptible to trypsin and since hemoglobin is much in excess of trypsin (thus
binding most of the polyacrylate), the rate is stimulated at lower pH values.
The inhibitions by polyglutamate and polyacrylate are reduced by increas-
ing the ionic strength, as expected (Table 1-15-6). It has been shown that
copolymers of glutamate with other amino acids (e.g., tyrosine, phenylala-
nine, or leucine) are more effective inhibitors than glutamate polymers,
but the copolymer of glutamate and alanine is less inhibitory (Rigbi and
Sela, 1964). Ornithine polymers or copolymers with ornithine are not inhi-
bitory and will reactivate trypsin inhibited by glutamate polymers. This
is one instance in which the inhibition produced by glutamate polymers or
copolymers is progressive and depends on the incubation time. From the
different degrees of inhibition brought about by the various copolymers
and the effects of the ionic strength, it was concluded that forces other
than electrostatic are involved in the binding. It is interesting that trypsin
seems to be particularly susceptible to macroanions, inasmuch as neither
chymotrypsin nor papain is inhibited by heparin, although both enzymes
are negatively charged at physiological pH.
The hydrolysis of acetyltyrosine ethyl ester and methylhippurate by chy-
motrypsin is inhibited 35-50% by various proteins (seralbumin, ovalbumin,
and /?-lactoglobulin ) at concentrations equivalent to the enzyme and in the
absence of salt (Hofstee, 1960). Addition of salts, particularly multiply
charged ions, reduces or abolishes the inhibition. These proteins are as po-
tent inhibitors as the naturally occurring chymotrypsin inhibitors, but differ
in not being so sensitive to salt concentration. Carboxymethylcellulose and
nucleic acids (both DNA and RNA) also inhibit chymotrypsin (Hofstee,
1961). The complexes are dissociated by 100 mM KCl. The inhibition is
not complete at maximal binding of nucleic acid, indicating that the active
center is not directly involved in the interaction. Methylhippurate was the
substrate and if protein substrates had been used it is likely that the active
center would not have been accessible.
Pepsin
The hydrolysis of hemoglobin by pepsin is rapidly inhibited by poly-L-
lysine and this is readily reversible by adding heparin, which complexes
with the inhibitor (Katchalski etal., 1954). Cationic polyornithine and poly-
458
2. ANALOGS OF ENZYME REACTION COMPONENTS
p-aminophenylalanine act similarly, but anionic polyalanine, poly aspartate,
and polyglutamate do not inhibit, indicating purely electrostatic binding.
The inhibition disappears at high poly-L-lysine concentrations (Dellert and
Stahmann, 1955). Changes in the optical transmittance also occur (Fig. 2-
15); that is, low concentrations of inhibitor complex with the enzyme to
decrease the solubility, but as the inhibitor concentration rises the com-
FiG. 2-15. Inhibition of pepsin by polylysine of mean molecular weight 2100
at pH 4.7. Transmittance determined at 400 m//. (Data from Dellert and
Stahmann, 1955.)
plexes become more soluble. However, the correlation between inhibition
and transmittance is such as to suggest that the depression of enzyme ac-
tivity is by no means directly related to the aggregate size of the complex.
Some have believed that pepsin plays a role in the genesis or maintenance
of gastric ulcers and hence have looked for inhibitors that might be affective
clinically. Strange to say, they have invariably used macroanions such as
heparin, chondroitin sulfate, polyhydromannuronic sulfate, and various pol-
ymers formed by condensation of aldehydes with hydroquinonesulfonate
(Levey and Sheinfeld, 1954; Marini and Levey, 1955; Heymann et al., 1959).
The isoelectric point of pepsin is around 2.8 so the relative effectiveness of
macroanions and macrocations might depend on the experimental or phys-
iological pH. Although the hydrolysis of casein is inhibited to some extent
by these polymers, it is possible that this is due in part or wholly to the
formation of complexes with the casein. It has been claimed that chon-
droitin sulfate and the polymeric sulfonates reduce the number of ulcers
in Shay rats, but it is doubtful if this is related to pepsin inhibition, even
if it occurs, and there are other explanations (for example, inhibition of
lysozyme, which has also been implicated in ulceration).
INHIBITION BY MACROIONS 459
Lysozyme
It is not surprising that this mucolj'lic enzyme is inhibited by a variety
of macroanions since its isoelectric point is above 10.5. Heparin is a fairly
potent inhibitor of lysozyme (Kaiser, 1953), and it has been stated that
this is competitive with substrate (which was a dried preparation of M.
lysodeikticus) (Kerby and Eadie, 1953). Inhibition is also exerted by hyalur-
onate, polysaccharide of Pneumococcus, polyglutamate, DNA, and RNA
(Skarnes and Watson, 1955). Various synthetic polymeric sulfonates also
inhibit to different degrees (Heymann et al., 1959). The most potent inhi-
bitor yet found for lysozyme is poly glucose sulfate (molecular weight around
20,000), although oxidized polyglucose (containing carboxylate groups)
is likewise very active; tetraglucose sulfate is without activity (Mora and
Young, 1959). These inhibitions are generally reversed by increasing salt
concentration or by the addition of a macrocation, such as protamine, to
bind the inhibitor. Copolymers of glutamate, tyrosine, phenylalanine, and
leucine are potent inhibitors of lysozyme, and the inhibitions can be com-
pletely reversed by polylysine (Sela and Steiner, 1963). The greater inhi-
bitory activity of the copolymers compared to the homopolymer of gluta-
mate is attributed to nonionic bonds; although this is the most likely ex-
planation, one must recognize that the charge distribution is quite different
in the copolymer relative to the homopolymer.
Hyaluronidase
It was noted by Pantlitschko and Kaiser (1951) that hyaluronidase is
not significantly inhibited by low molecular weight substances or by high
molecular weight substances unless they are esterified with sulfate or are
otherwise anionic, and, furthermore, that inhibitory macroanions must be
filiform and not globular. Heparin and artificially sulfurated polysaccharides
are inhibitory; sulfurated hyaluronate, which can with some justification
be thought of as a true analog of hyaluronate, inhibits well. Hyaluronate is
a high molecular weight polymer of i\'-acetylhyalobiuronate units and hence
contains free C00~ groups; however, sulfuration essentially doubles the neg-
ative charge on the molecules, and prevents degradation by the enzyme.
Nitrated and acetylated hyaluronates are also inhibitory. A few macro-
anionic inhibitors can be mentioned but require no discussion: chitin sul-
fates, polymers formed from formaldehyde and various phenolic sulfonates
(e.g., hydroquinone, catechol, and resorcinol), polymers formed from form-
aldehyde and various hydroxybenzoates, polyesters of phosphate with phe-
nols and aniline, polystyrenesulfonate, sulfated pectate, polymethacrylate,
amylopectin sulfate, and heparin (Rogers and Spensley, 1954; Bernfeld et
al, 1961).
Alburn and Whitley (1954) suggested that the inhibition of hyaluronidase
460 2. ANALOGS OF ENZYME REACTION COMPONENTS
by heparin is competitive and this was studied in detail by Houck (1957 a),
who found competitive behavior from l/v-l/(S) plots for both heparin and
chondroitin sulfate B. The inhibitor constants and their variation with
Ki {mM)
Temperature K^ (mlf)
Heparin Chondroitin sulfate B
22°
0.70
6.17
27°
0.74
6.45
32"
0.78
6.68
37°
0.81
7.00
6.58
6.85
7.10
7.41
temperature are tabulated, and from these values it was possible to cal-
culate the important thermodynamic quantities shown in the following
tabulation:
- AF<^ - AH° - zl<S°
(kcal/mole) (kcal/mole) (e. u.)
Heparin 3.07 5.0 6.8
Chondroitin sulfate B 3.0 4.9 6.2
The rather weak binding might indicate that only a fraction of the anionic
groups on heparin or chondroitin sulfate B interact at close range with
enzyme cation groups. It is difficult to predict the entropy changes in the
interactions of such complex molecules because several factors may be in-
volved, e.g., the restriction of polymer configuration, possible changes in
enzyme structure, and release of water of hj^dration. The rather small
changes in AS observed are probably the result of the balancing of larger
changes in different directions. The importance of ion-ion interactions is
shown by the marked reduction in the inhibitions when the ionic strength
rises above 0.3.
The differences in inhibitory activity between low molecular weight sub-
stances and polymers made from them are well illustrated by Hahn and
Fekete (1953). Various phenolic compounds inhibit testicular hyaluron-
idase weakly, but upon polymerizing these with formaldehyde it is possible
to obtain potent inhibitors. Their results are expressed in terms of the po-
tency relative to resorcinol. y-Resorcylate has an activity of 1.5 while its
polymer has an activity of 980. The most active inhibitor is the polymer
COO- COO- COO- coo-
— C— R— C— R— C— R— C— R— C—
INHIBITION BY MACROIONS 461
of gentisate with values around 2000-2500 (Hahn and Frank, 1953). Poly-
esters of phosphate with phloretin (or other polyphenols) are very inhibi-
tory to hyaluronidase, 0.005 mg/ml completely abolishing enzyme activity
(Diczfalusy et al., 1953). Phloroglucinol phosphate polymer may be even
more potent, 0.00013 mg/ml inhibiting 80% (Ferno et al, 1953). Such poly-
mers may be represented as:
o- o- o-
— 0— P— 0— R— 0— P— O— R— 0— P— O—
OH OH OH
The extent of the inhibition may depend primarily on the distance between
anionic groups and the ability of the polymer to assume the appropriate
configurations on the enzyme surface.
Ribonuclease
Pancreatic ribonuclease is strongly and competitively inhibited by hep-
arin (Zollner and Fellig, 1952, 1953) but the results by different investigators
vary quite widely, due perhaps to different experimental conditions (espe-
cially pH and ionic strength), different preparations of heparin, and differ-
ent techniques for measuring the enzyme activity. The competitive nature
of the inhibition (at least the reduction in inhibition upon increasing RNA
concentration) has been confirmed by Roth (1953) and Houck (1957 b).
Increase in ionic strength reduces the inhibition as expected (Houck, 1957 b;
Heymann et al, 1958; Fellig and Wiley, 1959; Lorenz et al, 1960), although
Houck found some deviation from this at very high NaCl concentrations.
The results of Lorenz et al (1960) are quite typical (see accompanying
tabulation):
NaCl (m
31)
0/
/o
Inhibition
0
97
30
50
50
33
100
0
Lowering the pH from around 7.5-8.0 to 5.0 progressively augments the
inhibition (Zollner and Fellig, 1953; Roth, 1953 b), which is perhaps a re-
flection of the increasing positive charge on the ribonuclease (isoelectric
point is 9.5). The ribonucleases of rat kidney and liver (Roth, 1953 b) and
rat and guinea pig serum (Rabinovitch and Dohi, 1957) are also inhibited
462 2. ANALOGS OF ENZYME REACTION COMPONENTS
by heparin. It has been known for many years that heparin depresses cell
division and it has been said to prevent gelation of the mitotic apparatus.
Paff et al. (1952) found that heparin inhibits mitosis in cultured chick heart
fibroblasts and after 24 hr there is a marked accumulation of granular ri-
bonucleoprotein in the cells. It was postulated that heparin might interfere
with the metabolism of nucleoproteins and thereby block mitosis.
The inhibition of ribonuclease by DNA was first clearly shown by Houck
(1957 b) and this could be thought of more reasonably as a true analog
inhibition. Likewise, deoxyribonuclease can be inhibited by RNA, the K^
being around 0.00001 vaM for the endonuclease of E. coli (Lehman et al.,
1962 a, b). Although inhibition occurs with RNA from various sources, the
most potent inhibitor is the amino acid acceptor RNA from E. coli, the
inhibition being competitive. The other DNA-cleaving enzymes tested are
not inhibited. The potency and specificity of this inhibition cannot but
stimulate thoughts on the possible regulatory relationships intracellularly.
Other macroanions have variable effects on ribonuclease. ZoUner and
Fellig (1952) reported no inhibition by chondroitin sulfate, hyaluronate,
and alginate, but Houck (1957 b) found chondroitin sulfate A and hyalur-
onate to inhibit equivalently with heparin. Synthetic polyglucose sulfate is
a competitive inhibitor, the effect decreasing with increase in the pH, the
net charge on the ribonuclease being reduced (Mora, 1962). Vandendriessche
(1956) studied the inhibitions by sulfonated pectin, poly-j9,'/)-dioxydibenzyl
phosphate, and poly-L-aspartate, and found them to be much weaker than
heparin, while Fellig and Wiley (1959) claimed that the sulfation of a va-
riety of polysaccharides (e.g., cellulose, amylose, amylopectin, dextran, pec-
tate, and nitrochitin) produces inhibitors often more potent than heparin
(although in this work the inhibition by heparin was unaccountably weak),
Ribonuclease is also inhibited by copolymers of glutamate and tyrosine (or
phenylalanine), which are more affective than poly aspartate or poly gluta-
mate (Sela, 1962). Interactions between the benzene rings of the aromatic
amino acids and certain groups on the enzyme were believed to occur in
addition to the electrostatic forces. Possibly the most potent inhibitors
were discovered by Heymann et al. (1958) in a survey of 66 macroanions
of synthetic origin, some inhibiting around 50% at 0.001 mg/ml. It was
noted that the inhibitory activity is markedly reduced in the presence of
proteins, a point worth considering in the use of such substances in cellular
preparations. A number of these polymers exhibit antiviral activity against
influenzal and vaccinial infections in eggs. Sulfate groups seem to be par-
ticularly able to confer inhibitory activity on polymers and in the sulfated
polysaccharides the carboxylate groups may be relatively unimportant,
since Dickman (1958) showed that sulfation of pectate, pectate methyl
ester, and pectic amide gives inhibitors roughly equiactive. It should finally
be noted that ribonucleases of different origins may not be equally suscep-
INHIBITION BY MACROIONS 463
tible to macroanions. A striking difference was demonstrated by Nishitnura
(1960), bovine ribonuclease being readily inhibited by polyvinyl sulfate to
which Bacillus subtilis ribonucleases are completely resistant.
Lipoprotein Lipase
Macroions inhibit lipoprotein lipase but affect other lipases little or not
at all. The enzyme from chicken fat is inhibited by macrocations in a purely
noncompetitive fashion and the inhibition is rapidly reversible either by
dialysis or the addition of a macroanion (Korn, 1962). Poly-L-lysines of
increasing chain length are progressively more effective, which is one of the
few observations relating polymer size to inhibition. The degree of inhibition
by various copolymers of tyrosine and lysine depends on the content of
lysine, indicating purely electrostatic binding. The enzyme is also inhibited
by macroanions, which may act noncompetitively or competitively (e.g.,
heparin and polyglucose sulfate). The lipoprotein from mouse heart is inhi-
bited by many polysaccharide sulfates as long as there is at least 0.6 sulfate
group per repeat unit (Bernfeld and Kelley, 1963). The potency of the inhi-
bition is independent of the configuration of the polysaccharide, whereas
in the case of the chicken enzyme the highly branched polymers are less
effective than the more linear ones.
Polynucleotide Phosphorylase
This enzyme, usually obtained from Azotobacter vinelandii or Micrococcus
lysodeikticus , catalyzes the synthesis of polynucleotides, such as polyade-
nylate:
ADP + (A.AIP)„ ^ P, + (AMP)„^i
The synthesis of a particular polynucleotide may be inhibited by another
polynucleotide; thus polyuridylate inhibits the synthesis of poly adenylate,
and polyuridylate, poly adenylate, and RNA inhibit the synthesis of poly-
cytidylate (Mii and Ochoa, 1957). The formation of polyadenylate has been
shown to be inhibited by variously degraded yeast RNA and polyadenylate
(Hendley and Beers, 1959, 1961; Beers, 1961). Increase in substrate con-
centration reduces the inhibition but not in a strictly competitive fashion.
Competition with primers or activators was considered to be the most likely
mechanism. If the polynucleotides are too extensively depolymerized, the
inhibitory activity falls, indicating a certain minimal chain length for op-
timal inhibition. It was also established that phosphate groups on or adja-
cent to the 3'-position of the terminal ribose units are necessary. It is im-
portant to remember in such systems that interactions between different
polynucleotide chains, perhaps through hydrogen bonding, can occur (Warn-
er, 1957); the possible role of such interactions in the inhibitions observed is
not yet completely understood.
464 2. ANALOGS OF ENZYME REACTION COMPONENTS
Amylases
Both a- and /^-amylases from barley are inhibited by heparin at concen-
trations around 0.1 mg/ml (Myrback and Persson, 1953 a, b). These inhi-
bitions show a striking pH dependence. At pH 5 or above there is no inhi-
bition whereas below pH 5 the inhibition increases rapidly and is very mark-
ed at pH 4.5. The isoelectric points of amylases usually lie between pH 5
and 6, so it is likely that the combination of heparin with the enzymes
below pH 5 is due to the positive charge arising in this range. The electro-
static nature of the binding is indicated by the protection afforded by high
concentrations of NaCl during the incubation of the enzyme and the heparin.
Once the inhibition is established, raising the NaCl concentration will not
reactivate. The presence of substrate also protects the enzyme, pointing to
a basically competitive mechanism. Human salivary amylase is also inhib-
ited by heparin but the critical pH is at least one unit higher, possibly
because of a higher isoelectric point than the barley enzymes (Astrup and
Thorsell, 1954).
Enzymes Acting on Substrates Which Are Not Macroions
Macroions interact with proteins generally when the conditions are favor-
able (e.g., when the total charges on protein and macroion are opposite,
although this is not a necessary condition, and when the ionic strength is
low), so it is not surprising that many enzymes whose substrates are small
molecules are inhibited. The combination is probably seldom at the active
center, but more often a bridging or covering of the active center by larger
molecules bound at many points and in no specific orientation. This type
of inhibition is, of course, independent of structural relations with the sub-
strate, but must always be considered in the use of macroionic inhibitors,
A few examples only will be mentioned.
Prostatic acid phosphatase is potently inhibited by polyphloretin-phos-
phate in a noncompetitive fashion (Diczfalusy et al., 1953; Beling and Dicz-
falusy, 1959). K^ is given as 1.55 //g/ml. Polyestradiol phosphate is even
more inhibitory {K^ = 0.55 //g/ml). The inhibitions increase as the pH is
lowered below the pH^pt. Alginate of 556 residues and molecular weight
of 92,000 is also strongly inhibitory (^, = 0.0054 mM); the mechanism is
partially competition with substrate and partially interference with stabi-
lizing or protective substances present (Jeffree, 1957). The variation of the
inhibition with chain length is complex: chains of 10-100 residues inhibit
less, but below 10 the inhibition rises (Jeffree, 1956). A third potent inhi-
bitor is polyxenyl phosphate, a polydisperse, small molecular weight poly-
mer of branched chains and random coils, 66% inhibition being given
by 0.001 mM (Hummel et al., 1958). Polyhydroquinone is almost as potent,
polyethylenesulfonate and sulfonated polystyrene inhibit moderately, and
INHIBITIONS BY NUCLEOTIDES 465
polyacrylate and chondroitin sulfate are relatively inactive. The inhibition
by polyxenylphosphate is noncompetitive, partially reversed by raising the
NaCl concentration, and maximal at pH 4.6, decreasing on either side.
/?-Fructofuranosidase of yeast is inhibited by heparin and chitin disiilfate
at low pH's (Astrup and Thorsell, 1954). The glucuronidases from several
rat tissues are inhibited by heparin and hyaluronate (Becker and Frieden-
wald, 1949). Fumarase is inhibited 92% by heparin at a concentration of
0.2 mg/ml in a pH range of 5.5-6.0, whereas nucleic acid and chondroitin
sulfate inhibit only 26% and 11%, respectively, at 2 mg/ml (Fischer and
Herrmann, 1937). These examples indicate that inhibitions of this sort are
widespread. There has been only one investigation of the effects of ma-
croions on a complex metabolic sequence, the study of Dische and Ash well
(1955) on the actions of ribonucleate and some smaller anions, such as sul-
fate, on anaerobic glycolysis in pigeon hemolysates. RNA inhibits lactate
formation 48% at 1 mg/ml and the formation of phosphoglj^ceraldehyde
30% at 3 mg/ml. There would thus appear to be at least two sites of
action, the major effect being on the transformation of 3-phosphoglyceral-
dehyde to lactate.
INHIBITIONS BY NUCLEOTIDES
AND RELATED SUBSTANCES
Enzymes acting on pyrimidines, purines, nucleosides, nucleotides, or poly-
nucleotides are frequently inhibited by analogs of these substrates. Some-
times the inhibitors are normally occurring substances and it is here that
some of the most clear-cut and important examples of feedback control
and metabolic regulation have been demonstrated. In other cases the inhi-
bitors are synthetically derived abnormal analogs, which are frequently
quite depressant to rapidly growing cells where nucleotide metabolism is
active and have for this reason been studied with regard to carcinostasis.
Many instances of inhibition have been reported, some of which are sum-
marized in Table 2-28, but thorough quantitative work and studies of the
mechanisms are rather uncommon.
Most of the inhibitions in Table 2-28 appear to be competitive and prob-
ably many of those in which the kinetics were not studied are competitive.
Although the inhibitory activity of most of these analogs is low or moderate,
a few analogs, particularly the abnormal aza and fluoro derivatives, are
quite potent. It seems that all parts of the nucleotide structure can con-
tribute to the binding. Where only the nature of the purine or pyrimidine
component is varied, the inhibitions may be very different, indicating that
the ring-substituted groups can be important. The pentose structure is also
a determinant since different activities are observed in nucleotides contain-
ing ribose or deoxyribose. Finally, the number of phosphate units in the
466
2. ANALOGS OF ENZYME REACTION COMPONENTS
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INHIBITIONS BY NUCLEOTIDES 473
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2. ANALOGS OF ENZYME REACTION COMPONENTS
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INHIBITIONS BY NUCLEOTIDES
475
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2. ANALOGS OF ENZYME REACTION COMPONENTS
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INHIBITIONS BY NUCLEOTIDES 477
nucleotide is a factor in the binding. However, the binding energy does
not always increase with increase in phosphate residues or total negative
charge; the potencies of the inhibition of pig kidney phosphodiesterase are
AMP > ADP > ATP. Little is known about the topography of enzyme
sites for nucleotides but it is clear that multiple binding sites must be in-
volved. The relative binding energies for competitive inhibitors of yeast
pyridoxal kinase (see tabulation) (Hurwitz, 1953) show that here the purine
Inhibitor
Relative
— AF oi binding
(kcal/mole)
Adenine
4.60
Adenosine
4.55
AMP
4.51
ADP
5.17
ITP
3.77
Pyrophosphate
3.90
component is of primary importance, addition of ribose or phosphate res-
idues having little effect. Yet ITP inhibits while inosine does not, and
pyrophosphate is bound fairly tightly to the enzyme. The phosphate resi-
dues in ADP must not be oriented for interaction as optimally as free
pyrophosphate.
Azaguanine and Azauracil
The 8-azapurines are usually inhibitory to growth and this has been gen-
erally attributed to the incorporation of these analogs to form abnormal
polynucleotides which are nonfunctional or inhibitory. However, it has more
recently been found that these analogs or their immediate metabolic prod-
ucts are potent inhibitors of certain enzymes involved in purine metab-
olism, and it is possible that such actions contribute to the growth de-
pression. 8- Azaguanine has been studied the most thoroughly and has been
shown to inhibit the growth of many bacteria, fungi, algae, viruses, tissue
culture cells, chick embryos, epithelium, and tumors. It is usually antag-
onized by guanine or guanidylate, and occasionally by other purines, nu-
cleosides, and nucleotides. The intention is not to discuss these azapurines
in detail since it is a very large subject but to mention only a few observa-
tions bearing on enzyme inhibition.
Adenosine deaminase is inhibited reversibly by 8-azaguanine {K, = 0.28
mM) and the inhibition was stated to be noncompetitive, although the
l/f-l/(S) plot appears to indicate uncompetitive or coupling inhibition
(Feigelson and Davidson, 1956 b). Xanthine oxidase is also strongly inhi-
bited (Table 2-2). Unfortunately, very few enzymes operative in purine
478 2. ANALOGS OF ENZYME REACTION COMPONENTS
metabolism have been examined with respect to 8-azaguanine inhibition,
or to inhibition by nucleosides and nucleotides of 8-azaguanine, but this
type of mechanism for the growth inhibition must be borne in mind. 8-
AzaGTP is formed from 8-azaguanine and can serve as a substrate for
adenylosuccinate synthetase; it also inhibits competitively with respect to
GTP (Cohen and Parks, 1963). Here one sees the interesting situation in
which 8-azaGTP stimulates the rate when GTP is low and inhibits the
rate when GTP is high. It was pointed out that in such cases one might
obtain selective inhibition of an enzyme in tissues having a relatively high
substrate concentration. This behavior is characterized in the double re-
ciprocal plot by the curve for the inhibited reaction crossing the curve (or
straight line) for the uninhibited reaction and being nonlinear. 8-Azaguanine
suppresses the induction of liver glucose-6-phosphatase, fructose-l,6-diphos-
phatase, and tryptophan pyrrolase in the rat (Kvam and Parks, 1960) and
the formation of catalase in yeast (Bhuvaneswaran et al., 1961). These ef-
fects on protein synthesis may relate to an interference with nucleic acid
metabolism, but whether it is a general depression or a more specific block
is not known. Studies on 8-azaguanine in animals are complicated by the
rapid deamination to the relatively noninhibitory 8-azaxanthine so that
only a fraction of the quantity fed is available for either inhibition or in-
corporation (Mandel, 1955). Thus the usefulness of 8-azaguanine in tumor-
istasis is limited.
6-Azauracil is also generally growth-inhibiting and tumoristatic. It is
possible that 6-azauridine-5'-P (6-azaUMP) is the true inhibitor, since it
has been shown that orotidylate decarboxylase is strongly inhibited by
6-azaUMP, leading to the accumulation of orotidylate (Handschumacher
and Pasternak, 1958; Pasternak and Handschumacher, 1959). 6-Azauridine
is metabolized to 6-azaUMP and inhibition of the decarboxylase after ad-
ministration of 6-azauracil was demonstrated, so it is likely that the block
in pyrimidine metabolism is at this point and that this is an important
mechanism in the tumoristatic action. The inhibition of the decarboxylase
is characterized by a K, of 0.00075 mM (Handschumacher, 1960). It is
interesting to speculate that a similar mechanism might be involved in the
action of 8-azaguanine.
Fluoropyrimidines and Feedback Inhibitions in Pyrimidine Pathways
The fluoropyrimidines are among the most potent inhibitors of nucleic
acid biosynthesis yet discovered but the sites of action have not been com-
pletely elucidated. The 5-halogen analogs of orotate inhibit the conversion
of orotate to the uridine phosphates, the most active being the fluoro com-
pound (Stone and Potter, 1957). It was suggested that some of the action
could be due to nucleotide analogs formed from these, and it was shown
that 5-fluoroorotate is converted to 5-FUMP in yeast (Dahl et al., 1959).
INHIBITIONS BY NUCLEOTIDES
479
However, 5-fluoroorotate blocks earlier in the sequence, an inhibition of
dihydroorotase, which forms dihydroorotate by cyclization of carbamylas-
partate, having been observed (Smith and Sullivan, 1960). Orotate also
inhibits but more weakly, this being an example of negative feedback.
Orotidylate decarboxylase which forms UMP is inhibited by UMP, although
not by uridine, UDP, or UTP, and it is possible that 5-FUMP would also
inhibit at this locus (Blair and Potter, 1961). The conversion of dUMP to
TMP by thymidylate synthetase, which is phage-induced in E. coli, is very
potently inhibited by 5-F-dUMP, K^ being around 0.00005 mM, and follow-
ing a competitive phase there is irreversible reaction with the enzyme (Ma-
thews and Cohen, 1963). This illustrates the principle that analogs often
simulate feedback inhibition if they are structurally similar to the com-
pound normally exerting the inhibition. Some of the inhibitions observed
in pyrimidine nucleotide metabolism are shown in the following scheme
modified from Smith and SuUivan (1960).
A + CP
(FCMP, CTP)
CA
(O, FO)
DHO
DNA-
(FdUMP)
TMP -« — K dUMP
)( (FO)
OMP
X (UMP)
UMP
-UDP
(FUMP,
FUDP)
X *
UTP
CTP
]<. (FUMP, FUDP)
RNA
(A = aspartate, CP = carbamyl-P, CA = carbamylaspartate, DHO = di-
hydroorotate, 0 = orotate, OMP = orotidine-5'-P, and fluorinated analogs
are indicated by an inital F).
The effects of 5-fluorouTacil on protein synthesis in E. coli and B. mega-
teriimi are very interesting because total protein synthesis is not altered
significantly but the proteins formed have abnormal amino acid compo-
sitions (Gros and Naono, 1961). For example, the proteins contain less
proline and tyrosine but more arginine. The alkaline phosphatase has nor-
mal catalytic activity but is less thermostable, whereas a-galactosidase
480 2. ANALOGS OF ENZYME REACTION COMPONENTS
seems to be synthesized, as shown serologically, but is catalytically inactive.
An RNA fraction into which 5-fluorouracil is rapidly incorporated was de-
tected and it is possible that this is responsible for the changes in protein
synthesis. The halopyrimidines are very potent growth inhibitors but, al-
though much is known of their fate and actions (Brockman and Anderson,
1963), the over-all effects produced are usually so complex that the primary
sites of block have not yet been determined. These analogs can be metabol-
ized into such a variety of abnormal substances which can inhibit at many
different sites that it is likely no single mechanism for the growth inhibition
will be found. More investigations of the changes in the steady-state con-
centrations of the intermediates in these pathways, as they are affected by
the analogs, would be valuable in determining the important loci attacked.
A few examples of enzyme inhibitions which may be involved in feed-
back regulation of pyrimidine and purine metabolism will be cited because
of the importance of this type of inhibition in the control of nucleic acid
and protein synthesis.* Gots and Gollub (1959) described the suppression of
the formation of purine precursors in E. coli whenever a purine which sup-
ports growth is added, and the accumulation of 5-amino-4-imidazolecar-
boxamide is suppressed in certain mutants. Various purine analogs (e.g., 6-
mercaptopurine and 2,6-diaminopurine) act like the normal feedback inhibi-
tors, only more potently. These analogs can thus act on at least two sites
in the biosynthetic sequence, the formation of purine precursors and the
eventual utilization of the purines, their therapeutic usefulness possibly be-
ing related to this type of sequential inhibition. It may be noted, however,
that Rubin et al. (1964) have recently shown that sequential inhibition in
pyrimidine biosynthesis is not synergistic. Combinations of 5-azaorotate,
which competitively inhibits the conversion of orotate to orotidylate, and
6-azauridine, which after its metabolism to 6-azauridylate competitively in-
hibits the conversion of orotidylate to uridylate, do not produce greater
inhibitions in either isolated enzyme systems or leucocytes than are seen
with the individual analogs. Almost every step in the nucleotide synthesis
has been shown to be inhibited by more distal intermediates. The PRPP-
amidotransf erase, which catalyzes the first irreversible and specific step in
purine synthesis, utilizing glutamine as the amino donor, is inhibited by
AMP, ADP, ATP, GMP, GTP, other nucleotides, and some analogs (Wyn-
gaarden and Ashton, 1959); adenylosuccinate synthetase (Wyngaarden and
Greenland, 1963), aspartate transcarbamylase (Bresnick, 1963; Gerhart and
Pardee, 1962), phosphoribosylformylglycineamidine synthetase (Henderson,
* Although an intermediate or product in a metabolic sequence is shown to be
an inhibitor of an enzyme catalyzing a previous step, it is perhaps not justified to
call it a feedback inhibitor, which implies that inhibition occurs during the in vivo
operation of the pathway. For various reasons the substance may not play a role in
regulating metabolism, even though it is a reasonably potent inhibitor.
INHIBITION BY NUCLEOTIDES 481
1962), and other enzymes are inhibited in similar manner but each exhibits
a unique pattern (Table 2-28); while TTP inhibits several steps in its for-
mation, including CDP -^ dCDP, deoxyuridine -^ dUMP, and deoxythy-
midine -> dTMP (Ives et al., 1963). All of these inhibitions and many more
constitute possible feedback situations, but in the cell probably only a few
are important, since the concentrations of some intermediates may never
rise sufficiently to exert an effect, and compartmentalization may limit the
actions of these inhibitors. We have mentioned that certain enzymes ap-
pear to contain sites specially evolved for feedback inhibition (Gerhart and
Pardee, 1962, 1964), the best documented case being aspartate transcarba-
mylase, which is inhibited particularly well by CMP, CDP, and CTP. This
enzyme is normally a tetramer and it may be that these inhibitors alter the
subunit interactions since the monomer is not inhibited. Another interesting
example of this phenomenon is the inhibition of xanthosine-5'-P aminase
by psicofuranine (9-D-psicofuranosyl-6-aminopurine), which occurs in two
steps, a reversible pyrophosphate-dependent reaction with the enzyme and
an irreversible xanthosine-5'-P-dependent reaction (Udaka and Moyed,
1963). The first step can be observed in a psicofuranine-resistant bacterial
strain and here the inhibition is noncompetitive. It would appear that the
inhibitor is bound to a different site than that at which the substrate reacts
and this second site could have regulatory function.
Some interesting results have been obtained in the analysis of the inhi-
bitions produced by the metabolites of 6-mercaptopurine, a few of which
will be mentioned briefly. One product into which 6-mercaptopurine is con-
verted is 6-thio-IMP, a potent competitive inhibitor of IMP dehydrogenase
(which is involved in the formation of GMP in certain cells) {K^^ = 0.0036
mM) (Atkinson et al., 1963). The inhibition proceeds rather slowly, requir-
ing 10-20 min for completion, and the enzyme is then inactivated (Hamp-
ton, 1963). Evidence was presented that reaction occurs with an SH group
on the enzyme, a stable disulfide bond being formed with the 6-thio-IMP.
This is one example where an analog turns out to be an SH reagent. On the
other hand, adenylosuccinate lyase is inhibited by 6-thio-IMP only if a
metal ion is present and it was postulated that the metal ion forms a bridge
between the SH groups (E-S-Me-S-IMP) (Bridger and Cohen, 1963). These
inhibitions create new possibilities by which analogs can inactivate enzymes.
Another product derived from 6-mercaptopurine is 6-mercaptopurine ribo-
side-5'-diphosphate, which inhibits polynucleotide phosphorylase quite po-
tently (50% inhibition by around 0.03 mM), rapidly, and competitively
(Carbon, 1962). The role this enzyme plays in vivo or the significance of
such inhibition is not known.
These few remarks on the effects of nucleotides and related substances
are made only to suggest certain interesting aspects of enzyme inhibition
which broaden our concepts of how analogs may act; adequate coverage of
this subject, young as it is, would require a volume of this size or more.
482
2. ANALOGS OF ENZYME REACTION COMPONENTS
INHIBITIONS BY COENZYME ANALOGS
This field in which interest was stimulated by the demonstration of the
mechanism of sulfonamide action is a large one because of the great amount
of work done on the growth inhibition of microorganisms by these analogs,
so that here the presentation will be restricted to those aspects directly re-
lated to enzyme inhibition and specific metabolic disturbance. The competi-
tion between a coenzyme analog and a coenzyme for combination with the
apoenzyme for which the coenzyme is essential is basically of the same
nature as the examples of substrate analogs discussed previously. However,
there are usually additional complexities due primarily to the greater num-
ber of sites for antagonism. Figure 2-16 indicates some of the reactions
(1)
PRECURSORS iSLC -i^^
X
PHOSPHATASE [(4) (3) ATP +
KINASE
\ / (5,
CP + E =
E-
■CP
E-CP
A ^ B
Fig. 2-16. General scheme for the formation and possible reactions
of coenzymes in cells. C = a unit of the active coenzyme (e.g.,
nicotinamide, adenine, or thiamine), CP = the active coenzyme,
E-CP = the active enzyme-coenzyme complex for the reaction
A -> B, and X = any derived substance from the unit C, which
may be inactive, or active to some degree after phosphorylation,
or capable of interfering with the formation or action of the coen-
zyme. Reaction 1 may be a simple diffusion into the cell or be
mediated by facilitated diffusion or active transport; reaction (2)
occurs in cells which synthesize the coenzyme from precursors; re-
action (3) is usually a phosphorylation; reaction (4) is a dephos-
phorylation; and reaction (5) represents the complexing of the
coenzyme with the apoenzyme.
involved in coenzyme formation, destruction, and function. Inasmuch as
the catalytically active forms of most coenzymes are formed within cells
from precursors, these reactions and the membrane processes responsible
for entrance of the precursors must be considered as possible loci for analog
interference. Furthermore, in many instances the analogs are metabolized
along the same pathways as the coenzymes to form inhibitory products.
Certain coenzymes are active in phosphorylated forms and the reaction
immediately forming the active coenzyme is often a phosphorylation in-
INHIBITIONS BY COENZYME ANALOGS 483
volving ATP and a kinase. The analogs are occasionally phosphorylated
and exert their major effects in this form. The inability of most phosphor-
ylated substances to enter cells readily makes it necessary to use the analog
of the coenzyme precursor if inhibition in cell preparations is to be obtained.
Thus the initial analog or any of its metabolic products may interfere in a
number of reactions involving the coenzyme, and it is this that militates
against facile interpretations from superficially simple results. It should also
be evident that when the reaction catalyzed by the coenzyme-dependent
enzyme (A ^ B in Fig. 2-16) is determined, the kinetics of inhibition by
an analog of the coenzyme precursor will generally not be simple and, al-
though the fundamental block may be strictly competitive, the quantita-
tive relationship between the analog and the precursor will not necessarily
be competitive.
The direct effect of a coenzyme analog on the enzyme reaction requiring
the cooperation of the coenzyme will depend on the tightness with which
the coenzyme is bound to the enzyme. Some coenzymes are so tightly bound
that they remain on the enzyme through numerous isolation procedures,
and in such cases the addition of an analog, even though it has a high
affinity for the enzyme, may not be able to replace the natural coenzyme
rapidly enough to induce inhibition. It must be remembered that the analog
does not actively displace the coenzyme (i.e., it does not force it from the
enzyme) but only binds to the free enzyme; if essentially all of the enzyme
is combined with coenzyme, there is little opportunity for the analog to act.
For this reason experiments on coenzyme analogs are frequently done with
reconstituted enzymes. In such cases the enzyme and coenzyme are disso-
ciated by some means and the effect of the analog on the reconstitution of
the active enzyme is investigated, this allowing the analog to act on the
free enzyme and to demonstrate competitive behavior. This technique is
not, of course, so applicable to cellular systems.
It has been frequently stated that coenzyme analogs are specific inhi-
bitors. This is true in one sense inasmuch as these analogs or their meta-
bolic derivatives appear to interfere only with those enzymes or reactions
involving the corresponding normal coenzymes, in most instances. On the
other hand, the coenzymes often participate in several different types of
metabolic activity so that the metabolic disturbances produced by the
analogs may not be specific with respect to a single reaction. For example,
analogs of pyridoxal seem to interfere specifically with pyridoxal metab-
olism or the functions of pyridoxal phosphate, but pyridoxal phosphate
plays a role in many reactions of amino acids — racemization, transamina-
tion, oxidative deamination, decarboxylation, hydrolytic cleavage — as well
as being an important component of other enzyme systems, such as muscle
phosphor ylase, so that a deficiency of pyridoxal phosphate can induce wide-
spread disturbances. In addition to this, a generalized depression of amino
484 2. ANALOGS OF ENZYME REACTION COMPONENTS
acid metabolism can secondarily bring about changes in systems not in-
volving pyridoxal phosphate through reduction in the concentrations of
amino donors or suppression of enzyme synthesis. Whether the coenzyme
analogs can be considered as specific or not will depend on the complexity
and general metabolic activity of the preparation being studied.
Most coenzymes are derivatives of vitamins and it has usually been anti-
cipated that analogs would induce vitamin-deficiency states. This has been
demonstrated in some cases; that is, effective analogs have been found to
produce a pattern of symptoms roughly similar to those seen in deficiency
of the corresponding vitamin. Nevertheless, it should be clearly understood
that the situations are basically different. A dietary restriction of a vitamin
leading to a generalized depletion in the tissues would not necessarily bring
about functional changes identical to those caused by an analog, which
could be much more effective in interfering with certain functions of the
coenzyme than simple depletion and possibly leave other functions untouch-
ed. All of the various enzymes binding a particular coenzyme do not have
the same affinities for an analog. Even though the analog primarily inter-
fered with the transport of the vitamin into the cell, or blocked its further
metabolism to the active coenzyme, it is not justifiable to conclude that a
state of generalized depletion will result, because these effects will pre-
sumably not be exerted equally on all tissues. The differential penetration
of the analog into the various tissues will perhaps be one important factor
in determining the response. Contrary to vitamin depletion, analogs often
cause a rise in the renal excretion of coenzyme or its metabolites, due to
the displacement of the normal coenzyme by the analog in the tissues and
its release from the cells. The analog might also alter the formation of the
coenzyme from its precursors, or inhibit the metabolism of the active co-
enzyme, or in some manner change the renal excretion or resorption of the
coenzyme or its precursors, so that a variety of effects on over-all excretion
is possible. If it is desired to demonstrate metabolic or functional defects
due to an analog in a short period of time, it is usually necessary to restrict
the intake or reduce the medium concentration of the coenzyme or its pre-
cursor, since the relationship between the analog and the coenzyme is usual-
ly competitive or pseudocompetitive, but in such cases one must use the
coenzyme-depleted preparation as a control to characterize the effects of
the analog.
ANALOGS OF NICOTINAMIDE
AND THE PYRIDINE NUCLEOTIDES
The importance of nicotinate and nicotinamide is as precursors of the
coenzymes NAD and NADP, and they do not, as far as is known, act di-
rectly in any metabolic system, nor do they usually occur in significant
concentrations in living cells. Some of the reactions involved in NAD syn-
ANALOGS OF NICOTINAMIDE 485
thesis and breakdown are shown in the accompanying diagram. The major
route of NAD formation, at least in mammalian tissues, is probably through
reactions (l)-(3) since the alternative pathway (8)-(10) is kinetically and
thermodynamically unfavorable. Analogs of nicotinate can thus either di-
rectly inhibit any of these reactions or enter into the reactions to form
abnormal intermediates, and perhaps analogs of NAD or NADP, which
are inhibitory. (See reactions on page 486).
Inhibition of NAD Nucleosidase (NADase) by Nicotinamide
and Related Compounds
It will be convenient to discuss first the direct inhibitions by simple
pyridine derivatives and then proceed to those substances incorporated into
NAD analogs. There is a constant turnover of NAD in tissues and at least
a fraction of the degradative process is attributable to NADase, and in
tissue extracts or homogenates the splitting of NAD may be an important
factor determining the dehydrogenase activity. Thus inhibitors of NADase
might be expected under certain circumstances to protect the coenzyme.
Furthermore, it will be evident later that the mechanisms of NADase inhi-
bition are involved in the formation of abnormal NAD analogs. Mann and
Quastel (1941) were the first to observe an inhibition of NAD breakdown
by nicotinamide. They worked with brain suspensions and determined NAD
by adding lactate dehydrogenase and lactate. Nicotinamide at 25 mM was
found to prevent the breakdown of NAD almost completely, and addition
of nicotinamide increases the respiration of various systems oxidizing lactate
by preventing the destruction of NAD. Nicotinate, on the other hand, is
completely inactive. Many investigators have subsequently used nicotin-
amide to preserve NAD in various preparations, often in very high concen-
trations and without regard for the other possible inhibitions it might exert.
Handler and Klein (1942) soon showed that NADP splitting is also inhib-
ited by nicotinamide.
Mcllwain and Rodnight (1949) pointed out that the indiscriminate use
of high nicotinamide concentrations to protect NAD in metabolic studies
is unnecessary, since almost complete inhibition of NADase is seen at con-
centrations from 2 to 10 mM (actually they showed that 2.67 mM inhibits
73%). The problem of the proper concentration of nicotinamide to use is a
difficult one because the NADases of various tissues and organisms show
marked differences in susceptibility to inhibition. The early work was all
done on brain NADase, which is quite sensitive, and it has been found that
some other NADases are also sensitive, e.g., from beef spleen (Zatman et
al., 1953). However, the enzymes from rabbit erythrocytes (Alivisatos and
Denstedt, 1952; Rubinstein et al., 1956; Malkin and Denstedt, 1956), mouse
mammary gland and tumor (Branster and Morton, 1956), and lupine seed-
lings (Hasse and Schleyer, 1961) are only moderately sensitive to nicotin-
486
2. ANALOGS OF ENZYME REACTION COMPONENTS
x:
a.
o
o <D
>. d
a c
2 b£
«
a>
2; c g
^ is
t-i ^ o
e s
ni a!
M ^ Q i! Q
rt -^ _ _ _
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K rt — -- --
2 xi
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9-K -S.
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CM CO in oj
ANALOGS OF NICOTINAMIDE 487
amide {K^s usually between 20 and 100 mM). The NADase from Neuro-
spora crassa is quite resistant to nicotinamide (Kaplan et al., 1951).
The inhibition by nicotinamide is competitive with respect to NAD for
the weakly inhibited NADases of rabbit erythrocytes (Alivisatos et al., 1956;
Hofmann, 1955), lupine seedlings (Hasse and Schleyer, 1961), and Neuro-
spora. However, the inhibition of the sensitive mammalian NADases from
brain and spleen is noncompetitive and the elucidation of the mechanism
by Zatman et al. (1953) has provided important information on NAD me-
tabolism and its inhibition by a variety of agents. The binding of nicotin-
amide to the enzyme is, however, readily reversible upon dilution or dialysis.
The following reaction mechanism was suggested as a working hypothesis:
+NRPPRA + enzyme ^ enzyme - +RPPRA + N
l+H.O
enzyme + RPPRA + H+
An intermediate enzyme complex which is subsequently hydrolyzed is as-
sumed. The hydrolysis is irreversible and N + RPPRA will not form NAD.
The inhibition by nicotinamide is thus a competition with water for the
enzyme-+RPPRA complex. This complex is not a Michaelis-Menten ES
complex but a covalent-linked compound in which the energy of the ni-
cotinamide— riboside bond is conserved. The free energy for the hydrolysis
of this bond is — 8.2 kcal/mole, and its conservation in the complex is very
important for the exchange reactions catalyzed by this enzyme. If this
mechanism is valid, one should observe exchange between free nicotinamide
and the nicotinamide in NAD, and this was demonstrated by using nicotin-
amide-C^*. These NADases might be considered as transglycosidases and
able to transfer the RPPRA group to compounds structurally related to
nicotinamide to form NAD analogs (Zatman et al., 1954 a). The NADases
which are weakly inhibited do not operate by such a mechanism and do
not catalyze exchange reactions.
Another enzyme which is nicotinamide-sensitive and catalyzes a similar
exchange reaction is the nicotinamide riboside phosphorylase of human
erythrocytes (Grossman and Kaplan, 1958 a, b). l/w-l/(S) plots showed the
inhibition to be uncompetitive, which is usually interpreted as a combina-
tion of the inhibitor with the ES complex, but in this case is perhaps due
to the transfer nature of the reaction. In the scheme:
ERN v=^ E + RN
'>
ER '^
ERP — > E + RP
where nicotinamide riboside is the substrate and — d(Rl^)ldt = /-(ERP), it
is seen that nicotinamide will slow the reaction by shifting the equilibrium
488
2. ANALOGS OF ENZYME REACTION COMPONENTS
in favor of ERN, and that actually competition with phosphate rather than
with nicotinamide riboside might be expected. A further complication is
the finding that the exchange reaction and the sensitivity to nicotinamide
depend on a cofactor, which was isolated and shown to be either ergothio-
neine or a closely related compound. It is possible that ergothioneine acts
as a ribosyl acceptor and this would modify the kinetics of the nicotin-
amide inhibition. It is interesting that ergothioneine will make the Neuro-
spora NADase inhibitable by nicotinamide.
A variety of substances related to nicotinamide or other portions of the
NAD molecule are inhibitory to NADases (Table 2-29). There is a good
deal of variation in susceptibility between the different enzymes. In this
connection it may be mentioned that Handler and Klein (1942) found that
rabbit brain NADase is readily inhibited by 5-10 mM nicotinamide but
not inhibited at all by 160 mM picolinate, quinolinate, benzamide, a-amino-
nicotinate, trigonelline, adenine, adenosine, or pyridine. These inhibitions
probably involve different mechanisms. Some are not competitive and the
inhibitors probably participate in the transfer reaction as does nicotinamide,
while others are competitive and the inhibitors are bound reversibly to sites
at the active center, thereby preventing the binding of NAD. The inhibition
Nicotine
€f
Nikethamide
CON'
.C2H5
COO
^N
CH3
Trigonelline
,CH,
4(5)-3'-Pyridyl-
glyoxaline
CONHNH,
CONHNH-CH
Isoniazid
CH,
of beef spleen NADase by isoniazid exhibits unique kinetics inasmuch as
an increase in the NAD concentration actually increases the inhibition, this
suggesting that some interaction between NAD and isoniazid occurs, the an-
alog formed being the active inhibitor. Isonicotinamide inhibits similarly to
isoniazid. 3-Substituted pyridines generally inhibit somewhat more strong-
ly than the 4-substituted compounds. One of the most potent inhibitors is
ANALOGS OF NICOTINAMIDE 489
4(5)-3'-pyridylglyoxaline but the mechanism is unknown; it is quite pos-
sible that NADases other than from brain may not be so potently inhibited
by this substance since nicotine does not inhibit the beef spleen enzjTne.
Another surprisingly potent inhibitor is theobromine, which is bound to
rabbit erythrocyte NADase around 1 kcal/mole more tightly than the other
purines tested, the inhibition being competitive. Malkin and Denstedt
(1956) concluded from the inhibition data that NAD is attached to the
enzyme surface at the quaternary nitrogen and the pyrophosphate group.
It is rather strange that adenine is a reasonably effective inhibitor, whereas
adenosine or the adenine nucleotides are much weaker or completely with-
out action, since, if adenine were bound in the same position as it is when
part of the NAD molecule, one might expect ribose and phosphate groups
to augment the binding. Thus the inhibition by adenine and other purines
may involve interaction with the enzyme surface in a manner unrelated to
the normal binding of the purine component of NAD. This is further borne
out by the studies on multiple inhibition by Hofmann and Rapoport (1957)
(see accompanying tabulation), inasmuch as adenine does not add to the
inhibitions produced by inhibitors presumably interacting with enzyme
groups binding NAD.
Inhibitors
(I)/(S)
% Inhibition
Nicotinamide
50
64
Adenine
9
20
Both
65
NMN
10
44
Adenine
9
20
Both
43
NADP
4.5
40
Adenine
9
20
Both
39
3-Acetylpyridine and the Formation of Analogs of NAD
In a search for pjTidine derivatives which might have vitamin activity
against black tongue in dogs, Woolley et al. (1938) observed that 3-acetyl-
pyridine is not only ineffective but kills nicotinamide-deficient animals in
1 day, normal dogs being unaffected. 3-Acetylpyridine rapidly produces
signs of nicotinate deficiency in mice and at the LDjq (around 3 mg per day)
the animals succumb in 3 to 4 days (Woolley, 1945 b). The effects produced
are: rapid respiration, motor incoordination followed by complete paralysis,
emaciation, and inflamed skin and tongue. The mice can be completely
490
2. ANALOGS OF ENZYME REACTION COMPONENTS
P5
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ANALOGS OF NICOTINAMIDE 491
o ^-
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492
2. ANALOGS OF ENZYME REACTION COMPONENTS
P5
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ANALOGS OF NICOTINAMIDE 493
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494 2. ANALOGS OF ENZYME REACTION COMPONENTS
protected by providing nicotinate or nicotinamide in the diet. On the other
hand, yeast and most bacteria seem to be quite resistant to 3-acetylpyri-
dine, although the growth of Lactobacillus casei in nicotinate-free medium
is inhibited around 50% at 16.5 mM, a depression that can be reversed by
nicotinate but not by nicotinamide. Chick embryos are killed by 450-600 //g
3-acetylpyridine injected into the eggs and sublethal doses cause distur-
bances in embryogenesis (Ackermann and Taylor, 1948), These effects can
be completely reversed by nicotinamide; even 6000 jugjegg of the analog
can be counteracted by 380 //g of nicotinamide, indicating a competitive
relationship. Changes in the heart, characteristic of nicotinate deficiency,
are produced by perfusion of 1.6-8 mM 3-acetylpyridine through the iso-
lated rabbit heart, dysrhythmias and a-v block occurring within 30 min
(Braun, 1949). Subsequent perfusion with nicotinamide reverses these ef-
fects but the concentration must be around 100 times that of the analog.
These early observations all point to the interference by 3-acetylpyridine
in the metabolism or function of nicotinate or nicotinamide. If it is assumed
that the primary role of these metabolites is the formation of the NAD and
NADP coenzymes, the following possible mechanisms for inhibition by 3-
acetylpyridine might be imagined. (1) Inhibition of some step in the syn-
thesis of NAD [especially reactions (1) to (3) in the scheme on page 486],
(2) inhibition of the interconversion of nicotinate and nicotinamide, (3) en-
trance into one of the pathways of nicotinate metabolism to form inhibitory
intermediates, (4) formation of an NAD analog, either through the normal
pathway or by the exchange reaction catalyzed by NADase, (5) inhibition
of NADases and related enzymes, and (6) direct interference with NAD or
NADP to inhibit dehydrogenase activity.
It will be well to consider certain aspects of the metabolism of 3-acetyl-
pyridine before taking up the problem of how this analog induces its inhi-
bitory effects. 3-Acetylpyridine at doses around 0.5 g/day increases the urin-
ary excretion of iV-methylnicotinamide in both normal and nicotinate-de-
ficient dogs (Gaebler and Beher, 1951). The iV-methylnicotinamide could
arise either from a disturbance of nicotinamide metabolism, since iV-meth-
ylation is an important reaction in the elimination of nicotinamide, or
directly from the 3-acetylpyridine. The oxidation of 3-acetylpyridine to
nicotinate might be anticipated because benzoate is formed from acetophe-
none in the tissues, the entire sequence being
3-acetylpyridine ->■ nicotinate -> nicotinamide -> iV-methylnicotinamide.
It was found that the latter explanation is correct by determining labeled
iV-methylnicotinamide formed from labeled 3-acetylpyridine (Beher et al.,
1952). It was pointed out that in the course of its oxidation and methylation
the analog might also interfere with nicotinate metabolism. Since iV-meth-
ylnicotinamide accounts for only about 10% of the administered 3-acetyl-
ANALOGS OF NICOTINAMIDE 495
pyridine, other excretory products were investigated and increased urinary
nicotinate and various glucuronides were found (Beher and Anthony, 1953).
No urinary 3-acetylpyridine could be detected. An interesting suggestion
that the oxidation of 3-acetylpyridine may involve NAD(P) enzymes was
made; this might mean that in nicotinate-deficient animals, where NAD(P)
levels are low, the oxidation of 3-acetylpyridine would be impaired and the
analog would be more toxic. 3-Acetylpyridine presents the strange situation
wherein the analog is detoxified to the normal metabolite, and this would
presumably tend to counteract the inhibitory effects. In low dosage (25-
60 mg/day), 3-acetylpyridine can protect against black tongue in dogs but
at higher dosage it can create a nicotinate deficiency (McDaniel et al., 1955).
Animals may have a limited ability to oxidize 3-acetylpyridine; small
amounts are mainly oxidized and little 3-acetylpyridine is left to inhibit,
whereas the larger doses exceed the metabolic capacity of the system. This
is indicated by the results of Guggenheim and Diamant (1958), who deter-
mined the excretion of iV-methylnicotinamide in rats given comparable doses
of nicotinamide and 3-acetylpyridine (see tabulation). Beyond a dose of 50
Dose
iV-Methylnicotinamide excretion
from:
(mg/kg)
Nicotinamide
3 - Acetylpyridine
Ratio "
0
23
23
10
65
40
2.5
20
182
105
1.9
50
225
120
2.1
100
700
218
3.5
200
1950
376
5.5
" Ratio calculated after substracting endogenous excretion.
mg/kg there seems to be relatively less oxidation of the analog. Adminis-
tration of 3-acetylpyridine-C^*H3 to rats leads to 30% of the activity ex-
pired as CO2 and 44% eliminated in the urine during 24 hr (Beher et al.,
1959); since the total dosage was probably around 100 mg/kg, smaller doses
might be even more efficiently oxidized. 3- Acetylpyridine is also partially
metabolized to NAD and the 3-acetylpyridine analog of NAD, as will be
discussed shortly. Finally, nicotinamide mononucleotide excretion is aug-
mented by 3-acetylpyridine and it is possible that this mainly originates
directly from the analog (McDaniel et al., 1955). The metabolism of 3-acetyl-
pyridine and the compounds derived from it thus depend on the species,
the dose, and whether the animals are normal or nicotinate-deficient.
We shall now examine the effects of 3-acetylpyridine on the tissue levels
of NAD, the formation of NAD analogs, and the enzymic activities of these
496 2. ANALOGS OF ENZYME REACTION COMPONENTS
analogs. Most of this work has been done by Kaplan and his associates at
Johns Hopkins, and a summary of their most important results will be
given. It was first demonstrated that the incubation of NAD, brain NADase,
and isonicotinyl hydrazide (isoniazid, IHN) leads to the formation of the
INH analog of NAD, which was isolated in good yield, and it was postulated
that the antitubercular activity of isoniazid may be related to the appear-
ance of this nonfunctional or inhibitory analog (Zatman et al., 1954 b).
It was soon shown that a variety of pyridine derivatives can exchange with
nicotinamide in the presence of certain NADases to form NAD analogs;
these include isonicotinamide, iproniazid, ethylnicotinate, and S-acetylpyri-
dine (N. 0. Kaplan c^oi., 1954). The formation of 3-AcPyr-NAD* in tissue
homogenates and whole animals is inhibited by nicotinamide. The exchange
reaction and hydrolysis may be represented as:
NRPPRA + E
^
E-NRPPRA
E-RPPRA ->
E + RPPRA
ti-'^
XRPPRA + E
:i±
E-XRPPRA
where NRPPRA is NAD, XRPPRA is the NAD analog, E-RPPRA is the
relatively stable ribosyl enzyme complex, and X is the pyridine derivative
exchangeable with nicotinamide. The over-all exchange reaction would be:
NRPPRA + X ^ XRPPRA + N
Injection of 3-acetylpyridine leads to a rise in total pyridine nucleotides in
most tissues; in the liver this is NAD and none of the analog is demon-
strable, due presumably to the oxidation of 3-acetylpyridine to nicotinate,
whereas in brain, spleen, and tumors 3-AcPyr-NAD appears. In tumors the
NAD content actually decreases as 3-AcPyr-NAD increases. The equilibrium
between NAD and any of its analogs, and the ratio of their concentrations
in a particular tissue, will depend on (1) the AF between NAD and the
analog, (2) the concentrations of N and X, (3) the rate of transformation
of X to nicotinate, if it occurs, and (4) the relative bindings of NAD and
its analogs to the dehydrogenases. The time courses for the formation of
3-AcPyr-NAD from 3-acetylpyridine and the toxic reactions led to the sug-
gestion that the toxic and lethal actions are related to the NAD analog;
whether the toxicity depends on a reduction of NAD or a rise in 3-AcPyr-
NAD was undecided.
In order to determine the nature of the effects of 3-acetylpyridine on
tissue metabolism, it will be necessary to consider the ability of 3-AcPyr-
* The analogs of NAD will be designated by prefixes of this type, following Kaplan,
since this is convenient if not exactly accurate.
ANALOGS OF NICOTINAMIDE 497
NAD to replace NAD as the coenzyme for the various dehydrogenases.
3-AcPyr-NAD can function in most NAD-dependent dehydrogenase reac-
tions. In some cases it can be reduced more rapidly than NAD (horse liver
alcohol dehydrogenase, beef liver glutamic dehydrogenase, Lactobacillvs d-
and L-lactate dehydrogenases) and in other cases proceeds more slowly
(yeast alcohol dehydrogenase, beef heart lactate dehydrogenase, yeast gly-
ceraldehyde-3-P dehydrogenase), while in a few instances the rates are ap-
proximately equivalent (rabbit muscle lactate dehydrogenase) (N. 0. Kap-
lan et al, 1956; van Eys et al, 1958; N. 0. Kaplan, 1959; Stockell, 1959).
3-AcPyr-NADP is reduced about one fifth as fast as NADP in the pig
heart isocitrate dehydrogenase system and is inactive in erythrocyte glu-
cose-6-P dehydrogenase (N. 0. Kaplan et al., 1956; Marks et al., 1961). The
relative rates do not necessarily reflect the relative bindings to the dehy-
drogenases. In those cas^s where coenzyme activity is low but binding is
appreciable, the NAD or NADP analogs can inhibit the dehydrogenases;
thus 3-AcPyr-NAD inhibits glucose-6-P dehydrogenase quite strongly {K,=
0.03 mM) and this is competitive. NAD analogs other than 3-AcPyr-NAD
are usually less active and tend to be more inhibitory. Thionicotinamide-
NAD, nicotinyl-hydroxamate-NAD, and nicotinyl-hydrazide-NAD com-
petitively inhibit lactate and alcohol dehydrogenases, whereas 3-benzoyl-
pyridine-NAD inhibits beef heart lactate dehydrogenase uncompetitively
(Anderson and Kaplan, 1959). The introduction of 3-acetylpyridine, or
other pyridine analogs, can thus produce several effects on tissue dehydro-
genase activity, and in the general case will bring about an imbalance of
the normal relative substrate oxidations, due to altering the rates of the
various dehydrogenases in different ways. Unfortunately there has not yet
been sufficient study of the oxidative abilities of tissues isolated from ani-
mals treated with 3-acetylpyridine. However, it is probably safe to assume
that at least a major cause of the toxic effects is the inhibition of certain
dehydrogenases by the 3-AcPyr-NAD formed.
The various NAD analogs have been very useful in demonstrating differ-
ences between dehydrogenases from different tissues or species. For exam-
ple, beef heart and rabbit muscle lactate dehydrogenases react better with
NAD than with 3-AcPyr-NAD, but the lactate dehydrogenases from lob-
ster heart and thorax muscle react better with the analog (N. 0. Kaplan,
1959). Kaplan et al. (1960) have pointed out that the molecular heterogeneity
of enzyme active centers has phylogenetic significance. It is possible to
classify animals with respect to the affinities of their dehydrogenases for
the coenzymes or their analogs, and it is hoped that further investigation
along these lines will elucidade some of the evohitionary problems relative
to the changes in the active center configurations.
We must now examine the evidence for other sites of action for 3-acetyl-
pyridine and related analogs, Mcllwain (1950) reported that 3-acetylpyri-
498
2. ANALOGS OF ENZYME REACTION COMPONENTS
dine inhibits spinal cord NADase 65% at 11 laM. Such inhibition could be
due to (1) direct competition with NAD, (2) reaction with E-RPPRA to
form a relatively stable complex, thereby depleting free enzyme, or (3) in-
hibition by a 3-AcPyr-NAD analog formed. In the case of the INH-NAD
analog, the inhibition seems to be mainly of the third type (Zatman et al.,
1954 b). Nicotinamide deaminase is inhibited quite well by 3-acetylpyri-
dine; the inhibition is competitive and 50% at (I)/(S) = 20 (Grossowicz
and Halpern, 1956 b). Yeast alcohol dehydrogenase is inhibited directly
by substituted pyridines (van Eys, 1956; van Eys and Kaplan, 1957 a).
The inhibitions are related to the pK„'s of the analogs (Table 2-30) and a
Table 2-30
Inhibition of Yeast Alcohol Dehydrogenase by Substituted Pyridines"
Substituted
pyridine
Concentration for 50% inhibition (mil/)
pA'a
Total base
Pyridinium ion
iV -methyl
derivatives
4-CH3
6.11
20
0.013
3-CH3
5.82
40
0.013
5.8
None
5.27
70
0.0066
5.5
3-CONH2
3.40
230
0.00029
5.0
3-COCH3
3.39
—
—
4.0
3-CHO
3.37
—
—
4.2
3-COOC2H5
2.24
300
0.000026
3.6
3-CN
1.45
600
0.0000085
3.2
3-SO3-
2.9
13
0.0000051
—
" From van Eys and Kaplan (1957 a).
straight line is obtained by plotting pK^ against p^^. The pyridinium ions
are presumably the active inhibitors. The iV-methyl derivatives are rela-
tively weak inhibitors. The pyridine N must be important for the binding,
its properties being altered by the substituents (the stronger the electro-
negativity of the substituent, the greater the inhibition). It is thus evident
from these data that the pyridine analogs can inhibit various enzymes di-
rectly; it is likely that these effects are not as important as those arising
from the corresponding NAD analogs in whole animals.
If the major actions of 3-acetylpyridine are mediated through 3-AcPyr-
NAD, the susceptibility of microorganisms or animals to 3-acetylpyridine
ANALOGS OF NICOTINAMIDE 499
will depend primarily on the exchange activity of the NADases present,
and perhaps secondarily on the ability to oxidize the 3-acetylpyridine to
nicotinate.
Some of the effects of 3-acetylpyridine on tissue functions and whole
animals were mentioned at the beginning of this section, and some of the
more recently studied actions will now be discussed. The LDjq for the in-
traperitoneal route is 300-350 mg/kg in mice and 80 mg/kg in rats (Cogge-
shall and MacLean, 1958). Hicks (1955) found that the administration of
3-acetylpyridine to rats and mice at doses around the LD50 produces ne-
crosis of adrenal medulla, of certain neurons in the supraoptic nucleus of
the hypothalamus, and of the pyramidal layer of the hippocampus. No
effects on the cerebral cortex were observed, contrary to the action of most
metabolic inhibitors. These effects are not seen in nicotinate deficiency,
but the picture may represent a more accelerated and acute deficiency;
it is possible that the regions affected are more dependent on the pyridine
nucleotides, but differential penetration might also be a factor. Coggeshall
and MacLean (1958) found that single LD50 doses to rats lead to weakness
of the extremities, inspiratory rhonchi, urinary incontinence, and other
symptoms, but gross pathological examination of the organs showed no-
thing remarkably abnormal. Surviving mice show motor incoordination and
a slight to complete loss of neurons in the hippocampal areas CA3 and CA4;
some damage to other hippocampal areas and the dentate gyrus may occur,
but no changes in other brain areas were detected. It was concluded that
the hippocampus must be metabolically different from the rest of the brain.
Rats given 100 mg/kg of 3-acetylpyridine develop ataxia, hyperkinesis,
and convulsions and it was found that 5-10% of the total brain pyridine
dinucleotides is 3-AcPyr-NAD (Brunnemann et al., 1962). The maximal
levels of the abnormal analogs occur at 6-8 hr and various regions of the
brain differ in the fraction incorporated, the highest levels being found in
the hippocampus (Herken and Neuhoff, 1963; Willing et al., i964). The ad-
ministration of 4-acetylpyridine does not lead to incorporation or to toxic
symptoms. This Berlin group of workers favors the concept that the cen-
tral neurological effects of 3-acetylp>Tidine are due primarily to interference
in electron transport as a result of the inhibitions produced by the 3- AcPyr-
NAD(P) formed in the brain.
Hollander and I have studied the effects of the acetylpyridines on the
isolated rat atrium and, although the work is not yet complete, the basic
actions are clear.* 3-Acetylpyridine at 1 mM increases the contractile ten-
sion of the atria 10% and simultaneously the resting and action potential
magnitudes are increased 4-5%. The action potential duration and con-
* We have found that most commercial samples of the acetylpyridines are quite
impure and redistillation under reduced pressure is necessary to obtain reliable pre-
parations.
500 2. ANALOGS OF ENZYME REACTION COMPONENTS
duction rate are either not affected or slightly decreased. The effects are
somewhat greater at 5 niM, contractile tension increasing 25%. 2-Acetyl-
pyridine and 4-acetylpyridine produce very much the same effects, and
nicotinamide itself quite potently stimulates atrial contractility, although
in this case the resting and action potentials tend to decrease. In view of
these effects by substances not giving rise to NAD analogs, the acute ac-
tion of 3-acetylpyridine on the atria must be attributed to some other me-
chanism. The lack of depressant activity at these concentrations is also
interesting, since metabolic disturbances invariably produce certain char-
acteristic changes. The contractile stimulation by nicotinamide increases
with concentration and at 100 mM is around 100%. The mechanism is not
understood but seems to be unrelated to membrane potential changes.
Inhibition of Dehydrogenases by Nicotinamide and Related Compounds
The inhibition of NADases and inhibitions dependent on the NADase-
catalyzed exchange reactions have been discussed. We now turn to the
inhibitions of NADP-dependent dehydrogenases by nicotinamide and other
substituted pyridines. The groups and interactions involved in the binding
of the pyridine coenzymes to the dehydrogenases have been discussed by
Shifrin and Kaplan (1960). Sulfhydryl groups are often important and have
frequently been thought to react with the pyridine N of the coenzymes,
while in some dehydrogenases Zn++ is involved and perhaps reacts with
the phosphate or adenine residues. It is apparent that the coenzymes are
bound in different ways to different dehydrogenases and this will determine
to some extent the ability of analogs to inhibit. The nature of the very
tightly bound intramitochondrial NAD in unknown, but possibly it is much
more difficult to inhibit intact mitochondrial dehydrogenases than the iso-
lated and often reconstituted enzymes usually studied.
The first report of an inhibition of metabolism by nicotinamide was made
by Baker et al. (1938) in connection with a study of the action of nicotine
on cerebral respiration. However, the inhibition of brain slice respiration is
slight, 30 mM depressing the oxidation of glucose 8% and lactate 15%.
A greater differential effect on glucose and lactate oxidation is exerted by
nicotinate, the inhibitions at 30 mM being 9% and 57%, respectively. Von
Euler (1942) made the initial investigation of dehydrogenase inhibition and
found inhibitions of both beef liver glucose dehydrogenase and heart lactate
dehydrogenase (see accompanying tabulation). The generally weak inhi-
bitory activity and the greater potencies of nicotinate and pyridine-3-sulf-
onate, compared to nicotinamide and pyridine-3-sulfonamide, may indicate
that these substances do not actually combine with the pyridine-binding
site on the enzymes, and it is even doubtful if they may be classed as NAD
or NADP analogs. However, the inhibition by the pyridine-3-sulfonate ap-
pears to be competitive. Brink (1953 b) continued von Euler's work in
ANALOGS OF NICOTINAMIDE
501
Inhibitor
Relative "KT "
Lactate DH
Glucose DH
Salicylate
p-Aminobenzoate
Salicylamide
Pyridine-3-sulfonate
Nicotinate
Benzenesulfonate
m - Aminobenzoate
Benzoate
Pyridine-3-sulfonamide
Nicotinamide
Benzamide
Trigonelline
Adenosine
Adenosine-3'-P
ATP
7.7
—
14.1
8.6
14.2
—
22.1
27.6
32.6
12.0
43.4
34.1
51.7
—
54.2
75.2
60.0
135
105
41.5
—
46.3
inhibition
No inhibition
7.2
12.6
—
21.0
" The relative "K/' values were calculated from the inhibitions at varying con-
centrations in order that the relative potencies of the inhibitors could be more readily
compared; they are not absolute values.
Stockholm but determined the inhibitor constants by plotting, so that the
values in the accompanying tabulation for beef liver glucose dehydrogenase
Inhibitor
Ki Relative — ZiF of binding
(mM) (kcal/mole)
4-Pyridoxate
Pyridoxal
3-Hydroxypyridine
Nicotinate
2-Methylnicotinat«
Nicotinamide
Isonicotinyl hydrazide
Pyridine
Isonicotinate
Trigonelline
ATP
Adenosine-3'-P
Adenosine
Adenine
Phosphate
0.3
4.98
0.8
4.39
9.2
2.89
21.5
2.36
25
2.26
23
2.32
200
0.99
300
0.74
inhibition
inhibition
1.75
3.92
3.5
3.48
9.25
2.89
12
2.73
810
0.13
502
2. ANALOGS OF ENZYME EEACTION COMPONENTS
are more reliable than in the tabulation above. The K^^^ for NAD is 0.00428
mM (relative —AF would be 7.63 kcal/mole) so that none of these inhi-
bitors are bound nearly so tightly. Substitution in the 3-position of the pyri-
coo
CONH,
Nicotin-
amide
SOpNH,
Pyridine-3-
sulfonamide
CONHNH,
Isonicotinyl
hydrazide
OH
CHO
COO
COO
3 -Hydroxy -
pyridine
Salicylate
CHoOH
CH,OH
dine ring is necessary for significant inhibition, but the nature of this group
can vary considerably and certainly no marked electrostatic interaction is
involved. The pyridine N would not seem to be of much importance in the
binding, since benzamide is about as inhibitory as nicotinamide, and ben-
zenesulfonate almost as potent as pyridine-3-sulfonate, and yet A"-methyl-
ation (to form trigonelline) abolishes the inhibition. The extra 2.1 kcal/mole
provided by the 3-hydroxy group suggests the possibility of hydrogen bond-
ing to the enzyme from this position, but dipolar and dispersion interactions
could also account for this. It may be noticed that the results with alcohol
dehydrogenase (Table 2-30) are in certain respects different than those
with glucose dehydrogenase. In this connection one must remember that
the ionization constants of these analogs should be considered, and it may
be that the major effect of the substituent groups is by modification of the
p^a of the pyridine N. Until these problems have been treated quantita-
tively, it is impossible to evaluate accurately the relationship between
structure and inhibitory activity. In any event, it is clear that the major
binding energy of NAD is contributed by the adenine nucleotide portion
of the molecule, so that pyridine derivatives might not be expected to be
potent inhibitors of dehydrogenases. The relatively strong inhibitions pro-
duced by pyridoxal and 4-pyridoxate may be significant for cellular meta-
bolic regulation and further study of the effects of these substances on
various dehydrogenases is probably warranted.
Nicotinamide has been frequently used in homogenates to inhibit the
splitting of NAD by NADases, as discussed above, and often at concentra-
ANALOGS OF NICOTINAMIDE 503
tions sufficiently high to interfere with dehydrogenases. Feigelson et al.
(1951) investigated this problem in liver homogenates and noted first that
nicotinamide reduces endogenous respiration, an effect reversed by NAD.
At 50-100 mM, nicotinamide stimulates the endogenous respiration some-
what, perhaps due to protection of NAD, but at higher concentrations in-
hibits quite potently. Malate dehydrogenase was partially purified and ni-
cotinamide inhibited competitively with respect to NAD with K^ — 0.00367
mM, and K, = 113 raM, corresponding to about 5 kcal/mole tighter bind-
ing for the NAD. Care must thus be used in the choice of nicotinamide con-
centration when NADase inhibition is desired. Results with lactate and
glucose-6-P dehydrogenases from rabbit erythrocytes are verj' similar (Ali-
visatos and Denstedt, 1952). Nicotinamide inhibits competitively with K^
around 100 mM. It was also shown that incubation of the apoenzyme with
nicotinamide in the absence of NADP leads to progressive irreversible inac-
tivation of the dehydrogenases; this may be related to the possible location
of binding sites for the coenzymes on adjacent helices of the apoenzyme,
separation of these sites occurring unless they are held together by the
coenzymes. Nicotinamide also inhibits 6-phosphogluconate dehydrogenase
competitively (Dickens and Glock, 1951). The NADH oxidases from pigeon
liver microsomes and mitochondria are inhibited 40% and 23% by 20 mM
and 80% and 78% by 200 mM nicotinamide, respectively (Jacobson and
Kaplan, 1957 a).
A unique nicotinamide derivative, A'-piperidinomethylnicotinamide,
which is claimed to be a specific dehydrogenase inhibitor has been reported
by Matkovics et al. (1961). Inhibition of methylene blue reduction in liver
homogenates in the presence of various substrates was studied. Unfortu-
nately the inhibitor concentrations are not given (only the milligrams added)
CONH-CH.— N
N-Piperidinomethylnicotinamide
and no control experiments on endogenous activity are included. Inhibition
of the oxidation of glucose, malate, lactate, and glutamate was observed.
However, succinate oxidation is also inhibited, indicating that this sub-
stance is not specific for the NAD(P) dehydrogenases. Furthermore, no
evidence for competition with the coenzymes was provided. Much more
work must be done before this inhibitor can be accepted as having specific
anticoenzyme activity.
504 2. ANALOGS OF ENZYME REACTION COMPONENTS
Pyridine-3-sulfonate and Pyridine-3-sulfonamide
These analogs might be expected to inhibit nicotinate and nicotinamide
metaboHsm. Both inhibit the growth of various bacteria and the inhibitions
can be overcome by nicotinamide (Mcllwain, 1940). Reference to the tab-
ulation on page 501 shows that lactate dehydrogenase is inhibited more
by these analogs than the corresponding nicotinic compounds, whereas glu-
cose dehydrogenase behaves in the opposite fashion (von Euler, 1942).
Feeding pyridine-3-sulfonate at 5% in the diet to mice produces no
signs of nicotinate deficiency, but the mouse does not require exogenous
nicotinate (Woolley and White, 1943 a). However, nicotinate-deficient dogs
are made worse by administration of the analog (Woolley et al., 1938),
although Gaebler and Beher (1951) observed no effect of 0.5-2 g/day of
pyridine-3-sulfonate on the excretion of A^-methylnicotinamide, erythrocyte
coenzyme level, or general health of either normal or nicotinate-deficient
dogs. Hicks (1955) found hippocampal necrosis in only one animal given
pyridine-3-sulfonate, so that it is presumably not as effective as 3-acetyl-
pyridine. Brain NADase is not inhibited by pyridine-3-sulfonamide (Mcll-
wain, 1950), and the sulfonate does not significantly inhibit either beef
spleen NADase (Zatman et al., 1954 a) or nicotinamide deaminase (Grosso-
wicz and Halpern, 1956 b). The most potently inhibited enzyme examined
seems to be yeast alcohol dehydrogenase, the sulfonate being the most po-
tent inhibitor of all the substituted pyridines (Table 2-30). There is no
evidence that NAD analogs can be formed from these substances. The
respiration of resting Mycobacterium phlei, either without substrate or with
glycerol, is not inhibited by 1 mM pyridine-3-sulfonate, but proliferating
bacterial respiration is inhibited 52-85% (almost completely by 10 niM)
(Miiller et al., 1960). These scattered observations do not arouse much in-
terest in these analogs, but perhaps the proper systems have not been
studied.
6-Aminonicotinamide
This analog has been called the most potent nicotinamide antagonist
available (Johnson and McCoU, 1955). The acute LD50 in mice is 35 mg/kg,
although 2 mg/kg/day leads to 50% mortality by the eleventh day (John-
son and McColl, 1956). Simultaneous administration of 50 mg/kg nicotina-
mide raises the LD50 8-fold. It is very toxic to rabbits, producing loss of
motor control and paralysis, and in rats it produces these and other signs
of nicotinate deficiency (Halliday et al., 1957). The endogenous respiration
of liver homogenates from treated mice is depressed 70% and lactate oxi-
dation is depressed 49%; addition of NAD counteracts these depressions.
However, no effect is observed when the analog is added directly to liver
slices. The possibility of the formation of a NAD analog was entertained
and such an analog was soon isolated following incubation of NAD, 6-ami-
ANALOGS OF NICOTINAMIDE 505
nonicotinamide, and NADase (Johnson and McCoU, 1956). The NAD ana-
log was also detected spectroscopically in the livers and kidneys of treated
mice. This NAD analog is completely inactive with yeast alcohol dehydro-
genase. If the analyses of Shapiro et at. (1957) actually represent true NAD,
the small decreases observed following administration of 6-aminonicotina-
mide at 30 mg/kg for 3 days (14% in liver, 17% in adenocarcinoma, and
none in brain) would point to the NAD analog as being inhibitory to dehy-
drogenases. This explanation was accepted by Friedland et al. (1958) on
the basis of decreases in tissue ADP and ATP, as well as oxidative inhi-
bition. Although it has generally been assumed that the central effects of
6-aminonicotinamide are due to the formation of an abnormal NAD(P) ana-
log and to reduction in normal NAD(P), Redetzki and Alvarez- O'Bourke
(1962) found that the NAD level in the brain is only slightly depressed,
despite the rather marked decrease of liver NAD, and obtained no evidence
for the occurrence of an abnormal analog. The 6-aminonicotinamide analog
of NAD inhibits creatine kinase and pyruvate kinase noncompetitively,
about 40% depression occurring at 1 mM (von Bruchhausen 1964). It is
unlikely that these actions can be important in vivo unless these enzymes
are much more sensitive in intact cells.
Administration of the analog to adenocarcinoma-bearing mice leads to
inhibition of certain enzymes determined in homogenates of the tumor:
lactate dehydrogenase is not affected, glyceraldehyde-3-P dehydrogenase is
inhibited 44%, the conversion of /?-hydroxybut>Tate to acetoacetate is in-
hibited 69%, and a-ketoglutarate oxidase is inhibited 83% (Dietrich et at.,
1958). It was believed that the NAD analog is quite tightly bound to the
apoenzymes and prevents the combination with NAD.
6-Aminonicotinamide exerts a depressing action against the growth of
certain lymphosarcomas and adenocarcinomas, and this is reversed by nic-
otinamide (Halliday et al., 1957). Tumor regression occurs at 3-4 mg/kg/
day but some weight loss also occurs; at lower doses the weight loss can
be minimized with some reduction in carcinostatic activity, but combined
at these lower doses with 8-azaguanine it is reasonably effective (Shapiro
et al., 1957). It was considered to represent a new class of potentially useful
carcinostatic agents. It is interesting that 6-aminonicotinate is one-seventh
to one-fifteenth as toxic as the amide, suggesting either that penetration
of the acid is Kmiting or that conversion to the amide is slow.
Inhibition of NAD(P) Enzymes by Various Nucleotides
and Related Substances
The study of these inhibitions has three major purposes: (1) to obtain
information on the nature of the active centers and the binding groups of
the coenzymes, (2) to understand better the mutual relationships between
these naturally occurring substances and the possible regulatory effects
506 2. ANALOGS OF ENZYME REACTION COMPONENTS
exerted in cellular metabolism, and (3) to find useful inhibitors that may
specifically inhibit particular reactions in complex systems. Some of the
results on different types of enzyme involving NAD or NADP are summa-
rized in Table 2-31. It is unfortunate that in very few instances have the
types of inhibition been determined and it is seldom possible to calculate
accurately the K/s or even relative K/s, from which interesting binding
energy information might be obtained.
One may first ask: Does the inhibitory activity generally increase as ri-
bose and phosphate groups are added? The answer is roughly in the ajBfir-
mative for NAD kinase, NADH pyrophosphatase, NADH oxidase, NAD :
NADP transhydrogenase, alcohol dehydrogenase, and malate dehydroge-
nase, but in a few enzymes there appears to be no definite trend, while in
some the addition of a group may reduce the binding. The addition of a
phosphate to adenosine to form 5'-adenylate (AR — »■ 5'-ARP) leads to only
0.1 kcal/mole extra binding to the alcohol dehydrogenase and 0.4 kcal/mole
to the NAD kinase, but an increased binding of over 2.4 kcal/mole for the
NADH pyrophosphatase. The further addition of a phosphate to form ADP
increases the binding approximately 0.7 kcal/mole for the NAD kinase,
1.1 kcal/mole for alcohol dehydrogenase, and 0.6 kcal/mole for the NAD :
NADP transhydrogenase, whereas the binding to NADH oxidase or NADH
pyrophosphatase is unchanged or slightly reduced. Addition of another phos-
phate to form ATP leads to increased binding only for the liver NADH
oxidase. Addition of a ribose to ADP to form ARPPR has no effect for
NAD kinase but increases the binding around 0.7 kcal/mole with NADH
pyrophosphatase. Final addition of nicotinamide to ARPPR to form NAD
increases the binding around 1.9 kcal/mole for NAD kinase and NADPH-
glutathione reductase, whereas a reduction of 1.9 kcal/mole in the binding
to NADH pyrophosphatase is observed. Addition of nicotinamide to 2'-P-
ARPPR to form NADP leads to a 2.3 kcal/mole increase in binding for the
NADPH-glutathione reductase and to very little change for the NADP-
cytochrome c reductase. The marked variation in behavior between en-
zymes and the uncertainty in the accuracy of the energy values make it im-
possible to draw definite conclusions or formulate rules for these inhibitions.
It appears that all the components of the NAD and NADP molecules can
participate in the binding, although not all of them need function for a
particular enzyme. The rather marked inhibition occasionally exerted by
NADH on NAD reactions, or by NAD(P) on NAD(P)H reactions, indicates
not only the specificity of these enzymes but points to a somewhat different
orientation of the oxidized and reduced forms on the enzymes.
A more interesting correlation emerges when one considers the variation
of inhibitory potency with the position of phosphate groups on the adenyl
ribose. In NAD the 5-position is phosphorylated and enzymes involving
NAD are more readily inhibited by 5'-AMP than by 2'- or 3'-AMP (NAD
ANALOGS OF NICOTINAMIDE
507
kinase and malate dehydrogenase). However, NADP is additionaUy phos-
phorylated in the 2-position and, as has been especially emphasized by
Neufeld et al. (1955), enzymes reacting with NADP are frequently inhibited
by 2'-AMP. In addition to the enzymes listed in Table 2-31 (NADPH dia-
phorase, NADP-cytochrome c reductase, glucose-6-P dehydrogenase, and
isocitrate dehydrogenase), they found phosphogluconate dehydrogenase and
NADP-activated oxalacetate decarboxylase to be inhibited by 2'-AMP more
than by the other AMP's. It was suggested that 2'-AMP may be a useful
inhibitor to distinguish between NAD and NADP enzymes. It must be
admitted that an insufficient number of NAD enzymes have been examined.
It is interesting that the NAD : NADP transhydrogenase is inhibited more
potently by 2'-AMP than by 3'- and 5'-AMP.
100
Fig. 2-17. Inhibitions of glutamate semialdehyde reductase by various nucleosides
and nucleotides. (From Smith and Greenberg, 1957.)
The importance of phosphate groups for the binding of inhibitors of this
type is seen strikingly in the study on glutamic semialdehyde reductase by
Smith and Greenberg (1957) (Fig. 2-17). The inhibitions by AMP, ADP,
and ATP are competitive, but NADP inhibits noncompetitively. Although
the addition of the first phosphate to form AMP has little effect (not more
than 0.1 kcal/mole), the addition of each of the next two phosphates to
508
2. ANALOGS OF ENZYME REACTION COMPONENTS
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ANALOGS OF NICOTINAMIDE 513
form ADP and ATP leads to about 1.2 kcal/mole increase in binding energy.
The addition of the 2'-phosphate to form NADP from NAD increases the
binding markedly and changes the nature of the inhibition. It is rather
strange that the addition of nicotinamide riboside to ADP lowers the bind-
ing energy about 1 kcal/mole.
Williams (1952) found that malate dehydrogenase is inhibited by adenine,
adenosine, and ATP. From this observation he concluded that such nor-
mally occurring substances may well affect dehydrogenases and other en-
zymes in the cell. His work stemmed from the report of Raska (1946) that
administration of 300-500 mg/day adenine to dogs on normal diets leads
to the development of multiple avitaminosis after 10-20 days; signs of ni-
cotinate deficiency, such as black tongue, were noted. The many more data
now available serve to strengthen Williams' conclusion, since even more
potent inhibitors have been reported. There has been much speculation
concerning the regulation of oxidative reactions by adenine nucleotides me-
diated through coupled phosphorylation. It is quite possible that other more
direct effects on dehydrogenases occur, both in the cell (particularly in the
compartmentalized mitochondria) and in experimental enzyme prepara-
tions where the concentrations of added nucleotides are often high enough
to inhibit appreciably. From Table 2-31 we see that five enzymes are inhi-
bited from 22% to 50% by ATP at concentrations from 1 mM to 3.4 mM;
ATP is commonly added to mitochondrial preparations at these or higher
concentrations. An experimental survey of dehydrogenase inhibitions by
nucleotides would be valuable. Chen and Plant (1963), on the basis of the
fairly potent inhibitions exerted by certain nucleotides on the NAD-link-
ed isocitrate dehydrogenase (Table 2-31), felt that some regidation of cycle
activity may be exerted, and if such does occur it would be a very impor-
tant factor in understanding not only the effects of nucleotide analogs but
also of many inhibitors which either primarily or secondarily alter the levels
of cellular or mitochondrial nucleotides. Another interesting point has been
brought out by Dalziel (1962) in connection with possible impurities in
preparations of the coenzymes. Although one might expect in many cases
an insignificant inhibitory effect of certain analogs because they have been
shown to bind less tightly than the normal coenzyme to the apoenzyme,
Dalziel correctly states that it is the relative values of K^ and K^^ which
are important, and K,,^ can be much higher than K^.. He calculated that
the presence of an analog of NADH as a 3% impurity can produce as much
as 70% inhibition of liver alcohol dehydrogenase if the analog and NADH
have the same affinity for the apoenzyme.
An NAD(P) analog which would bind to the NAD(P) site on dehydroge-
nases and then react chemically with some group at the site might well
be of some value in labeling these sites. Such an analog was investigated
by van Eys et al. (1962) on the basis that thiazole rings often open at al-
514 2. ANALOGS OF ENZYME KEACTION COMPONENTS
kaline pH to generate a free SH group. The NAD analog with 4-methyl-5-
(/?- hydroxy ethyl )thiazole replacing nicotinamide was found to behave in
this manner and to form a disulfide bond with SH groups at the site of
dehydrogenases, this binding being competitive with NAD. In the case of
horse liver alcohol dehydrogenase, 2 moles of this analog are bound tightly
to each mole of enzyme.
One of the most interesting studies of dehydrogenase inhibition by nu-
cleotides is that of the complex effects of GTP on glutamate dehydrogenase
(Frieden, 1962, 1963). The K^ is 0.0003-0.0005 mM and the kinetics being
uncompetitive point to different sites for NADP and GTP. Furthermore,
GTP not only inhibits directly but increases the ability of NADH to inhibit.
Since the NADH inhibition is due to the dissociation of the enzyme into
four subunits, it is likely that GTP enhances the process, and this was dem-
onstrated ultracentrifugaUy. The dissociation of the tetramer enzyme it-
self is not necessarily the basic cause of the loss of activity; it is possible
that structural changes brought about by NADH and GTP produce both
dissociation and reduced catalytic activity. The behavior can be explained
adequately on the basis of three binding sites: (1) a coenzyme site, (2) a
purine nucleotide site with which GTP and activating nucleotides react,
and (3) a NADH-binding site. The following complexes are thus possible
— EC, ECI, ECg, ECgl, and EI — where C represents the coenzyme. The
binding of GTP to the enzyme depends on the presence of NADP at a
vicinal site, the EI complex probably not being of much importance. The
importance of this situation for the regulation of cell metabolism is obvious,
particularly since this enzyme plays a central role in many pathways. Frie-
den pointed out the likely relationship between glutamate dehydrogenase
and the or-ketoglutarate step in the cycle; GDP is required for the conver-
sion of succinyl-CoA to succinate and GTP is formed, which can suppress
the activity of glutamate dehydrogenase, an enzyme which under certain
conditions controls the steady-state level of a-ketoglutarate in the cycle.
He also suggests that ammonia formation by the liver, protein synthesis,
and glyconeogenesis can all be regulated by this inhibition involving a feed-
back site.
ANALOGS OF THIAMINE
Thiamine functions in metabolism in the pyrophosphorylated form as the
coenzyme in various reactions where a bond adjacent to a carbonyl group is
broken (a-cleavage), the active complex in each case being an aldehyde-
thiamine-PP-enzyme structure wherein a C — C bond is formed at the 2-
position of the thiazole ring. These reactions would include (1) a-keto acid
decarboxylation (e.g. pyruvate decarboxylase), (2) a-keto acid oxidation
(e.g. pyruvate and a-ketoglutarate oxidases), (3) the phosphoroclastic reac-
tion of pyruvate, and (4) a-ketol formation (e.g. transketolase and phos-
ANALOGS OF THIAMINE
515
phoketolase). Thus three major metabolic sequences — the pentose-P path-
way, the tricarboxylate cycle, and photosynthetic carbon dioxide fixation —
are dependent on thiamine-PP, since a-cleavage occurs in all, and a variety
of other metabolic processes can be secondarily affected. Thiamine defi-
ciency, or interference with the formation or function of thiamine-PP, can
produce profound metabolic and physiological disturbances. Animals re-
quire preformed thiamine, most plants can synthesize the entire thiamine
molecule, and microorganisms vary widely from complete dependence on
exogenous supply to complete synthetic ability. The responses of organisms
to thiamine analogs wiU depend on these factors as well as the role of thia-
mine in metabolism. The pathways of thiamine biosynthesis are not com-
pletely understood and the accompanying scheme is to be taken as pro-
visional and not necessarily applicable to all organisms. Thiaminase is ap-
parently absent or relatively inactive in most tissues and thus the reactions
catalyzed by this enzyme are probably not common or important. The most
important reaction is the pyrophosphorylation of thiamine since certain
analogs can interfere here or be similarly phosphorylated. Thiamine-PPP
has been included because its formation from thiamine in yeast has been
demonstrated (Kiessling, 1956), although it is coenzymically inactive. It
may be noted that ATP is required for thiamine-PP synthesis and that
"Pyrimidine" "Thiazole"
+ PP
'pyrimidine -PP"
+ P
"thiazole- P'
"pyrimidine-B"
+ :
"thiazole"
"pyrimidine-B"
+
"thiazole-PP"
thiamine- P
+ B
Thiamine
+ H,0
thiamine-PP
+ H,0
"pyrimidine"
: +
"thiazole"
"pyrimidine"
: +
"thiazole-PP'
thiamine- PPP
enzyme-thiamine- PP
(The pyrimidine portion of thiamine is indicated in quotes and is 2-
methyl-4-amino-5-hydroxymethylpyrimidine; the thiazole portion is
designated likewise and is 4-methyl-5-{2-hydroxyethyl)thiazole. B is
any base that can replace the thiazole in the exchange reaction catalyzed
by thiaminase. )
516 2. ANALOGS OF ENZYME REACTION COMPONENTS
thiamine-PP is involved in metabolic reactions leading to ATP, so that
interference with thiamine-PP formation or function will tend to deplete
the cells of ATP and perhaps further depress thiamine-PP synthesis.
The possible sites of action for thiamine analogs can be broadly classified
as (1) inhibition of thiamine-PP synthesis, either on the formation of thia-
mine or its pyrophosphorylation, (2) interference with the formation of
complexes between thiamine-PP and enzymes, and (3) inhibition of thia-
minase. In any case the inhibition may be exerted by either the analog or
its phosphorylated derivatives. There is no evidence that any significant
effects of any of the analogs studied can be attributed to thiaminase inhi-
bition, so the first two mechanisms are undoubtedly the most important
in the induction of thiamine deficiency symptoms. There is some evidence,
which will be discussed later, that thiamine may have a function or func-
tions unassociated with coenzyme activity, particularly in the nervous sys-
tem, and, if this is true, one might consider the interference by analogs
in this function.
It would appear that most of the groups in the thiamine molecule parti-
cipate in either the binding or the catalysis inasmuch as the structure can
not be significantly altered without loss of activity, and the number of ef-
fective analogs is rather small. The first report of enzyme inhibition by a
thiamine analog was by Buchman et al. (1940), who found yeast pyruvate
decarboxylase activity to be depressed by 4-methyl-5-hydroxyethylthiazole
diphosphate (which they called "thiazole pyrophosphate"), the phosphory-
lated thiazole portion of thiamine. Neither the nonphosphorylated com-
pound nor the monophosphate is inhibitory. It requires about 10 times as
much analog as thiamine-PP to inhibit 50%, but if the analog is added
before the thiamine-PP, the inhibition is more pronounced. These results
point to the importance of the phosphate groups in the binding. They
state, "We conclude that there has been demonstrated here a not hitherto
recognized type of competitive inhibition of enzyme reactions, caused by
competition not between substrate and inhibitor but between coenzyme
and inhibitor." (See formulas on page 517).
Either the pyrimidine portion or the thiazole portion of the thiamine
molecule can be altered to form analogs. Replacement of the thiazole ring
with a similarly substituted pyridine ring gives pyrithiamine, which was
shown by Robbins (1941) to inhibit the growth of certain fungi, and by
Woolley and White (1943 b) to produce thiamine deficiency symptoms in
mice. Replacement of the pyrimidine amino group with a hydroxyl group
leads to oxy thiamine, found by Bergel and Todd (1937) to lack vitamin
activity, and by Soodak and Cerecedo (1944) to be quite toxic to mice.
These two analogs have been studied the most thoroughly of the thiamine-
like compounds and remain the most frequently used to produce experi-
ANALOGS OF THIAMINE
517
X
^^
1
X 1
o^
o
o^
a
a?
a?
u
o
4.
sf
K
1
X
^^■^^
^u
^O
OT
^u
QJ
c
.3
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7^
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+2:
"u
c
s
+z
^U
S
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^u^
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E
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s"
ci
K
u
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u
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U
ll^
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1
^^
1 1 ■
z^z
■z
r^
o
o
u
af
sT
1
X
1
s
o
• I
0-P-=0
I
o
. I
4.
as*
k^
+ Z
J3 .
a?
o
X
■^ /^
a?
o
1
u
aT
rn -^
af
8
o
1
o
L
w
o
X
a?
a.
1
a?
CO (
a
c
S
0)
c
^
X
X
0)
c
+ 2
af
X
.2
+z
aT
+z
4,
X
o
rt
u
"i^
p
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>.
u
§5
aT 1
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o
(1)
^
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a!
<
aT
z
Jx
4^-H
S o
¥^
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¥^
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¥Y
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Z"«>Z
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z^ ^z
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Z^ /;2
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T
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af
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af
518 2. ANALOGS OF ENZYME REACTION COMPONENTS
mental disturbances in thiamine function.* Two other types of analog per-
haps deserve more attention than they have received: the imidazole analog
(the thiazole ring replaced with a similarly substituted imidazole ring) exerts
an antivitamin effect on bacterial growth (Erlenmeyer et al., 1948), and the
2' -n-butylpyrimidine analog produces thiamine-deficiency states in rats (Em-
erson and South wick, 1945), both substances being roughly of the same
potency as pyrithiamine and oxythiamine. A large number of interesting
analogs, such as those synthesized by Livermore and Sealock (1947) and
by Mano and Tanaka (1960), have not yet been adequately examined.
Effects on Enzymes Dependent on Thiamine-PP
It is clear that neither oxythiamine nor pyrithiamine is inhibitory to
pyruvate decarboxylase, but that the diphosphate esters can interfere with
the binding of thiamine-PP to the apoenzyme. Thus the yeast decarboxylase
is inhibited by oxy thiamine-PP but not by oxythiamine (Eusebi and Ce-
recedo, 1950; Navazio et al., 1956), and wheat germ decarboxylase behaves
similarly (Eich and Cerecedo, 1954, 1955). Pyrithiamine is noninhibitory
whereas pyrithiamine-PP is as effective as oxythiamine-PP (Woolley, 1951;
Eich and Cerecedo, 1954). Oxythiamine-PPP is inhibitory (Velluz and Her-
bain, 1951; Navazio et al., 1956) but it is possible that some of the action
results from the splitting off of a phosphate to form oxythiamine-PP, just
as the cocarboxylase activity of thiamine-PPP has been found to depend
on hydrolysis to thiamine-PP (Kiessling, 1956). It is not certain if oxy-
thiamine-P is inhibitory and contradictory results have been obtained.
These data indicate the importance of the phosphate groups for the bind-
ing, and this is supported by the observation of Wiethoff et al. (1957) that
pyrophosphate inhibits wheat germ decarboxylase competitively with thia-
mine-PP (the pyrophosphate must be added before the thiamine-PP).
Thiamine-PP and the phosphorylated analogs are bound fairly tightly to
the apoenzyme {K„^ for thiamine-PP is usually near 0.001-0.003 mM) and
thus the order of addition of coenzyme and analog is important, especially
as the analogs are bound about 1.5-3.0 kcal/mole less tightly (Woolley,
1951; Stewart, 1957). If thiamine-PP is added 30 min before oxythiamine-
PP there is no inhibition, if they are added simultaneously there is slight
inhibition, but if oxythiamine-PP is added 30 min before the coenzyme
appreciable inhibition may be exerted (Eich and Cerecedo, 1954). Although
* It was found later that the material originally designated as pyrithiamine was a
mixture (Wilson and Harris, 1949). The pyridine analog was synthesized in pure
form and named "neopyri thiamine" but since the active compound in the early
preparation was this substance, it has been generally agreed that the original name be
restored. The general results of the early work are not invalidated, but the true po-
tency of pyrithiamine is greater than indicated there.
ANALOGS OF THIAMINE 519
some exchange between bound and free coenzyme and analog must occur,
it is too slow for equilibrium to be obtained easily. To determine the maximal
inhibiting power of an analog it is advisable to incubate the apoenzyme with
the analog previous to addition of the coenzjTne.
Pyruvate oxidase is inhibited similarly to the decarboxylase, as expected,
and in the case of pyrithiamine-PP it would appear to be competitive
(Woolley, 1951). Oxythiamine-PPP inhibits pyruvate oxidation in pigeon
breast muscle extracts but this may be mediated through the diphosphate
(Onrust et al., 1952). The formation of acetoin from pyruvate is also inhib-
ited by oxythiamine-PP (Eich and Cerecedo, 1954). However, Kuratomi
(1959) noted that oxythiamine, like thiamine, can form acetoin from py-
ruvate.
Transketolase from yeast is strongly inhibited by oxythiamine-PP (Datta
and Eacker, 1961) but oxythiamine itself has no effect (Dreyfus and Moniz,
1962). At 0.036 mM and 0.072 mM the inhibitions are 60% and 80%, re-
spectively, when the analog is added previous to thiamine-PP, but if the
oxythiamine-PP is added 2 min after the coenzyme, no inhibition is ob-
served. Addition of higher concentrations of thiamine-PP cannot reverse
the inhibition. Thus it is difficult for either the analog or the coenzyme to
displace each other from the apoenzyme. Oxythiamine-PP is bound more
tightly than thiamine-PP to the enzyme but it requires 2-3 hr to inhibit
50% when the enzyme initially contains thiamine-PP. The rate of dis-
placement for transketolase is even less than for decarboxylase, since
thiamine-PP was found to reverse oxythiamine-PP-inhibited enzyme 30%
in 20 min.
The inhibition of thiamine-PP-dependent enzymes by oxythiamine and
pyrithiamine will depend on whether these analogs can be phosphorylated
or not. Thus in the experiments of Kunz (1956), where oxythiamine and
pyrithiamine at 10 vaM inhibited pyruvate oxidation in rat liver mitochon-
dria 95% and 35%, respectively, one is not certain if there is direct inhibi-
tion or if the depression was due to the formation of small amounts of the
phosphorylated esters. Acetylthiamine inhibits to about the same degree
as pyrithiamine and this analog cannot be phosphorylated (unless it is first
deacetylated), so it would seem that these rather weak inhibitions may be
to some extent exerted directly. It is also interesting that oxythiamine
and pyrithiamine have no effect on pyruvate oxidation in brain mitochon-
dria, this being attributed by Kunz to a different structure or permeability
compared to liver mitochondria; different phosphorylative capacities might
also play a role. Phosphorylation of these analogs mediated through thia-
mine kinase seems to occur in most tissues and it is likely, as pointed out
by Woolley (1951), that the toxic reactions observed with oxythiamine
and pyrithiamine in animals are produced primarily by the phosphorylated
compounds.
520 2. ANALOGS OF ENZYME REACTION COMPONENTS
Accumulation of Intermediates and in Vivo Effects
Inasmuch as the oxidation of pyruvate requires thiamine-PP, one would
expect some accumulation of pyruvate in animals treated with thiamine
analogs if pyruvate oxidase is indeed inhibited in vivo. Such has been ob-
served in rats with oxythiamine (Frohman and Day, 1949; Gubler, 1961)
and pyrithiamine (de Caro et al., 1954). For example, rats injected intra-
peritoneally with 150 //g oxythiamine show an elevation in blood pyruvate
of 1.3 to 5.1 mg%; blood lactate also increases from 11.5 to 42.8 mg%. In
mice, pyrithiamine raises the blood pyruvate somewhat but oxythiamine
has no effect (de Caro et al., 1956), possibly indicating a species difference
since in rats oxythiamine is more effective than pyrithiamine (Gubler,
1961). The administration of oxythiamine to dogs at 6 mg/kg in three
doses leads to a marked rise in blood pyruvate (0.4 to 5.7 mg%) and thia-
mine is able to counteract this effectively (Wilson et al., 1962). Simultan-
eously there is a severe fall in liver glycogen (13 to 0.4 mg/g). Rats and
cats respond similarly but are less sensitive. Growth of Neurospora in the
presence of oxythiamine is accompanied by pyruvate accumulation and a
simultaneous reduction in pyruvate decarboxylase activity is demonstrable
(Sankar, 1958). Administration of increasing amounts of thiamine partially
or completely counteracts these effects on pyruvate levels, in all instances
where it has been tested.
We shall now turn to evidence of enzyme inhibition in the tissues of
analog-treated animals. It may be calculated from the data of Von Holt et al.
(1955) that feeding pjTithiamine to rats at 10 mg/kg for 7-12 days results
in some 63% reduction of pyruvate oxidation in liver homogenates. A
thorough investigation of the changing patterns of keto acid oxidation in
deficient and analog-treated rats has been made by Gubler (1958, 1961);
his results are summarized in Table 2-32. It is seen that the oxidation of
pyruvate is more sensitive than that of «-ketoglutarate to both dietary
deficiency and the analogs; this could relate to different displacing rates
in the two oxidases, or to different dependencies of enzyme activity on
thiamine-PP level. Oxidation of /5-keto acids, as expected, is not affected.
The effects of oxythiamine and pyrithiamine are roughly the same on all
tissues, with the exception of brain in which pyrithiamine is more effective.
The reason for this is not understood — it would seem unlikely that oxy-
thiamine is unable to penetrate the blood-brain barrier — but it may be
correlated with the fact that only pyrithiamine is able to produce poly-
neuritis in rats. Another difference between these two analogs lies in the
ability of thiamine-PP added in vitro to the mitochondrial suspensions to
counteract the depression of pyruvate oxidation. The loss of activity from
dietary deficiency of thiamine is readily reversed by adding thiamine-PP,
as anticipated; the loss due to pyrithiamine is surprisingly weU reversed
(to about 90% of the control values in brain and kidney); the loss due to
ANALOGS OF THIAMINE
521
Table 2-32
Effects of Thiamine Deficiency and Thiamine Analogs
ON THE Oxidative Decarboxylation of Keto Acids in Mitochondria "
Substrate
% Change
Tissue
Dietary
deficiency
Oxythiamine
Pyrithiamine
Liver
Pyruvate
-75
-56
-57
a-Ketoglutarate
-38
- 1
-11
a-Ketoisovalerate
+ 3
-22
-13
a-Keto-^-methylvalerate
-11
-11
-12
/S-Hydroxybutjrrate
+ 4
+ 2
+ 11
Brain
Pyruvate
-24
-18
-51
a-Ketoglutarate
+21
+ 6
-44
Kidney
Pyruvate
-56
-52
-61
a-Ketoglutarate
-55
+ 5
-37
Heart
PjTuvate
-32
-67
-58
a-Ketoglutarate
-24
-37
-17
° The analogs were administered to rats at oxythiamine/thiamine = 200, and
pyrithiamine/thiamine — 5, these doses producing polyneuritis in several days. The
figures give the changes observed in the oxidation of the substrates indicated in
mitochondrial suspensions. (From Gubler, 1961.)
oxythiamine is reversed poorly. The most obvious explanation of this is
that pyrithiamine blocks the synthesis of thiamine-PP, so that the tissues
are primarily deficient in cocarboxylase, whereas oxythiamine may exert
its inhibition mainly by binding to pyruvate oxidase in the form of its
diphosphate ester. These problems wiU be discussed after further effects
of these analogs have been presented.
Feeding pyrithiamine to pigeons leads to a 50% reduction in the p>Tuvate
decarboxylase activity in breast muscle, and this can be reversed by the
addition of thiamine-PP to the homogenates (Koedam et al., 1956). This is
accompanied by a marked reduction in the thiamine-PP content of muscle,
so that it was concluded that there is no essential difference between di-
etary thiamine deficiency and pjTithiamine feeding. Pyruvate dismutation
and acetoin formation in breast and heart muscle are likewise depressed
by pyrithiamine feeding (Koedam, 1958). A large single dose of pyrithia-
mine (2.5 mg) leads to a rapid inhibition of acetoin formation and even after
8 days the activity does not return to normal.
The respiratory quotient of rats treated with pyrithiamine (5 mg/day
522 2. ANALOGS or enzyme reaction components
intraperitoneally for 5-6 days) is lowered and, in contrast to normal or
thiamine-deficient animals, administration of glucose does not raise it (see
accompanying tabulation) (de Caro et al., 1954). There is thus an inhibition
R.Q.
Before glucose After glucose
Controls
Diet-deficient
Pyrithiamine-treated
0.81 0.93
0.78 0.92
0.72 0.77
of the total oxidation of carbohydrate, which can be directly attributed to
a block in pyruvate oxidation; however, it is difficult to explain the greater
effect in the diet-deficient animals. The only work showing an impairment
of transketolase function is that of Wolfe (1957). Rats deprived of thiamine
or given oxythiamine show a depression of the pentose-P pathway in the
erythrocytes, pentose accumulating, whereas pyrithiamine produces no
changes in transketolase activity even when the animals are paralyzed.
Some interference with amino acid metabolism by thiamine analogs, me-
diated through changes in the utilization of the a-keto acids, might be ex-
pected, but little study of this has been made. Pyrithiamine inhibits the
formation of aspartate and asparagine from glutamate in germinating Pha-
seolus seeds and there is an accumulation of ammonia, possibly due to the
lack of oxalacetate (Sivaramakrishnan and Sarma, 1954). These effects can
be attributed to a reduction in a-ketoglutarate oxidase. Oxythiamine and
pyrithiamine both inhibit the growth of Vibrio cholera and lead to the accu-
mulation of alanine, aspartate, and glutamate. The effects on the levels of
such amino acids will depend to a great extent on the pattern of metabolism,
i.e., on the over-all direction of transamination reactions.
Effects on Thiamine-PP Synthesis
Pyrithiamine has been found to inhibit thiamine kinase from chicken
blood (Woolley, 1950 a), rat liver (Eich and Cerecedo, 1954; Mano and Ta-
naka, 1960), rat intestine (Cerecedo et al., 1954), and pigeon liver (Koedam,
1958), and the inhibition appears to be competitive with thiamine. This in-
hibition is fairly potent: When the ratio pyrithiamine/thiamine is 1 the in-
hibition is around 50%, at a ratio of 5 it is 75%, and at a ratio of 10 it is
90% in rat liver and intestine (thiamine concentrations 0.01-0.1 raM). On
the other hand, oxythiamine inhibits thiamine kinase much less strongly
or not at all when present in similar ratios to thiamine (Eich and Cerecedo,
1954; Cerecedo et al, 1954; Koedam, 1958; Mano and Tanaka, 1960). The
ANALOGS OF THIAMINE 523
difference in inhibitory activity between these two analogs possibly indi-
cates the importance of the pyrimidyl 4 '-amino group. This was substan-
tiated in the demonstration by Cerecedo and Eich (1955) that oxypyri-
thiamine, in which the 4'-amino group is replaced by hydroxyl group, does
not inhibit rat liver thiamine kinase.
Mano and Tanaka (1960) studied a large series of thiamine analogs with
respect to their abilities to be phosphorylated by a rat liver thiamine kinase
system and their inhibitory potencies on thiamine phosphorylation (see ac-
companying tabulation). The analogs and thiamine were all at 0.1 milf.
Relative activity % Inhibition of
(thiamine — 100) thiamine kinase
Pyrithiamine
0
53
2'-Ethylthiamine
11
38
2'-/i-Butylthiamine
0
32
Oxythiamine
0
6
Diacetylthiamine
3
5
Thiothiamine
0
Stim 2
Dibenzoylthiamine
2
Stim 3
0-Acetylthiamine
7
Stim 6
Thiamine disulfide
0
Stim 12
The inhibitions by pyrithiamine and the 2'-alkylthiamines are competitive
and the following values of K^ were calculated: pyrithiamine 0.033 m.M,
2'-ethylthiamine 0.041 mM, and 2'-butylthiamine 0.043 mM. The inability
of the enzyme to catalyze the phosphorylation of oxythiamine and pyri-
thiamine is noteworthy in view of the theory that these analogs may exert
their effects in the diphosphate form. It may be recalled that Woolley (1951 )
failed to demonstrate the synthesis of pyrithiamine-PP in chicken blood.
There has been surprisingly little attention to this important problem of
analog phosphorylation in organisms.
Inhibition of Thiaminase
The importance of this enzyme in the mammalian metabolism of thiamine
is not known. The only evidence for its possible function is the appearance
in the urine of the pyrimidine and thiazole moieties of thiamine following
administration of thiamine. It would seem unlikely that inhibition of this
enzyme is an important factor in the toxicity of thiamine analogs, but it
could be of some significance in determining the effects on tissue levels of
thiamine-PP. It was stated by Soodak and Cerecedo (1944) that oxythia-
mine inhibits carp thiaminase but no data were given. Pyrithiamine inhibits
524 2. ANALOGS OF ENZYME REACTION COMPONENTS
this enzyme around 40% when analog and thiamine are both 0.5 mM (Sea-
lock and White, 1949). Pyrithiamine is split by the enzyme but at a slower
rate than is thiamine. Apparently it is bound more tightly than thiamine,
but reacts more slowly, since in mixtures of the two only the splitting of
thiamine is depressed. Thus too little is known of the effects of these analogs
on thiaminase to evaluate the importance of the inhibitions.
Sealock examined the effects of a series of substituted methylthiazolium
ions on fish thiaminase and found that 3-o-aminobenzyl-4-methylthiazole
(ABMT) is a particularly potent inhibitor (see tabulation) (Sealock and
Goodland, 1944). The inhibition is competitive, with K^^ = 0.0831 mM and
Ki = 0.00197 mM, possibly indicating that ABMT is bound 2.3 kcal/mole
Analog
3-o-Aminobenzyl-4-methylthiazole
3-^-Aminoethyl-4-methylthiazole
3-/3-Phthalimidoethyl-4-methylthiazole
3-o-Nitrobenzyl-4-methy]thiazole
3-Ethyl-4-methylthiazole
3-Phenyl-4-methylthiazole
3-Phenyl-2-methyl-4-methylthiazole
3-Ethyl-2-methyl-4-methylthiazole
Concentration
(mM)
Inhibition
Relative "X/'
0.5
100
< 0.005
0.5
48
0.54
0.5
6
7.8
0.5
2
24
10
9
101
10
5
190
5
0
>495
10
0
>990
more tightly than thiamine. The amino group is probably quite important,
since the replacement with a nitro group reduces the inhibition so markedly.
The aminobenzylthiazoles were later studied in greater detail (Sealock and
Livermore, 1949) and the position of the amino group was shown to be
critical, only the ortho compound being inhibitory (see accompanying ta-
bulation). On the other hand, the position of the thiazole methyl group is
not critical. Thiamine was 0.5 mM in these experiments. Kenten (1958) has
Amino position in Methyl position in Concentration „, ^ i -i • •
, , . XI,- 1 • / jiT\ /o Inhibition
benzyl ring thiazole ring (mM)
ortho
meta
para
ortho
meta
para
ortho
4
0.5
78.1
4
0.5
Stim
4
0.5
1.7
2
0.5
89.2
2
0.5
Stim
2
1
2.0
2,4
0.5
30.2
ANALOGS OF THIAMINE 525
found the thiaminase from bracken {Pteridium aquilinum) to be very sen-
sitive to ABMT, 15-20% inhibition being given by 0.002 mM and almost
complete inhibition by 0.05 mM. The inhibition is probably basically com-
petitive since it proceeds faster in the absence of thiamine. As far as I know,
this interesting compound has not been tested in whole animals to deter-
mine if thiaminase inhibition can be achieved and how this will alter thia-
mine metabolism.
Effects on Excretion and Tissue Levels of Thiamine
If these analogs displace thiamine or thiamine-PP from the tissues in
any way, or inhibit the transport or metabolism of thiamine, an increased
urinary excretion of thiamine would be expected, and this has been found
to occur in rats given 50 //g oxythiamine (Frohman and Day, 1949). One
might also predict that tissue levels of thiamine or its diphosphate would
be reduced, and this has been demonstrated for both pyrithiamine and oxy-
thiamine in mice, pigeons, and rats. The depression of tissue thiamine-PP
seems to be generally associated with a rise in blood pyruvate, so that at
least part of this depletion is related to enzymes involved in the oxidation
or decarboxylation of a-keto acids. Inasmuch as theories for the mechan-
isms by which these analogs act depend on the changes in tissue thiamine
levels, it will be necessary to examine the results with some care.
Pyrithiamine markedly depletes the tissues of thiamine-PP in pigeons.
Controls were fed 100 //g thiamine per day and another group was fed 623 //g
pyrithiamine each day in addition; after an average survival time of 19 days,
the thiamine-PP levels in the tissues were those shown in the following
tabulation (Koedam et al., 1956). The pyruvate decarboxylase activity in
muscle is reduced around 50% and adding thiamine-PP restores activity.
Thiamine-PP content (/<g/g)
Tissue
% Change
Controls
Pyrithiamine-fed
4.55
1.00
-78
2.61
0.78
-70
3.54
1.09
-69
4.42
1.80
-59
4.36
2.08
-52
Heart
Brain
Liver
Breast muscle
Kidney
The fall in thiamine-PP level is quite rapid; at 4 days it is mainly complete
in most tissues, and from the data on pjTuvate utilization it would appear
that a marked decrease occurs within 1 day (Koedam, 1958). Pigeons given
a single large dose of pyrithiamine (10 mg) and examined 64 days later show
526 2. ANALOGS OF ENZYME REACTION COMPONENTS
no permanent effect on the tissue thiamine-PP levels. An important obser-
vation was that pyrithiamine induces a more rapid depletion of tissue thia-
mine-PP than does elimination of exogenous thiamine. A comparison be-
tween dietary deficiency and pyrithiamine administration in rats was report-
ed by de Caro et at. (1954). The results in the accompanying tabulation were
Total thiamine content (//g/g)
Tissue
Controls Avitaminotic Pyrithiamine-fed
Liver 7.42 2.82 1.33
Muscle 1.60 0.80 0.71
Brain 3.38 2.89 0.57
obtained from rats injected with 5 mg pyrithiamine daily for 5-6 days and
rats subjected to a thiamine-free diet for a comparable time. In all cases
pyrithiamine produces a greater effect than simple elimination of thiamine
intake; the effect in brain is particularly striking and possibly correlated
with the polyneuritic symptoms produced by pyrithiamine.
Oxythiamine, on the other hand, does not seem to be so active in reducing
the tissue levels of thiamine-PP (Steyn-Parve, 1954). This analog at 1 mg/
day for 15 days to pigeons produces the changes summarized in the accom-
panying tabulation. No deficiency symptoms were noted and none would
Thiamine-PP content (fig/g)
Tissue
Controls Oxythiamine-fed % Change
Heart
Muscle
Cerebrum
Liver
6.70
3.40
-49
6.25
4.60
-26
4.15
3.75
-10
5.35
4.90
- 8
be expected at these tissue levels. The author believed that the change in
the liver is not significant; it is also possible that the large drop in the heart
thiamine-PP is too great, since in another experiment with twice the above
oxythiamine dosage the level decreases only 34%. The relative ineffective-
ness of oxythiamine was confirmed by de Caro et al. (1956) in mice, where
0.5-2 mg/day certainly produces little effect on the thiamine levels in muscle
and brain, although some decrease in liver is observed. There are likewise no
significant change in blood pyruvate. These results were confirmed and ex-
tended by Gurtner (1961), who administered pyrithiamine at 250 /yg/day
ANALOGS OF THIAMINE 527
and oxy thiamine at 10 mg/day to rats intraperitoneally for 29 days; a
thiamine-deficient group was also included (see accompanying tabulation).
Pyrithiamine
Oxythiamine
^i-deficient
Weight (% change)
-15
-43
-44
Cardiac rate (% change)
— 5
-18
-28
Paralysis (% occurrence)
93
0
46
Convulsions (% occurrence)
73
0
20
Blood pyruvate (% change)
+ 15
+288
+95
Tissue thiamine-PP (% change)
Liver
-35
+26
-97
Heart
-76
-20
-95
Brain
-86
- 6
-85
The differences in the actions of the two analogs is well illustrated here;
pyrithiamine produces marked neurological symptoms without much effect
on weight, cardiac rate, or blood pyruvate, although there is a very signi-
ficant fall in tissue thiamine-PP, whereas oxythiamine causes bradycardia
and weight loss without neurological effects, while the blood pyruvate is
elevated greatly without significant changes in tissue thiamine-PP.
Tissue Levels of Pyrithiamine
The pyrithiamine in rat tissues following the injection of 1 mg intraperi-
toneally was determined microfluorimetrically by Rindi and Perri (1961)
and Rindi et al. (1961) and the results are plotted in Fig. 2-18. These rats
had been maintained on a thiamine-deficient diet and within 1 day the pyri-
thiamine content of the tissues corresponded closely to the normal thiamine
content, indicating that the analog probably occupies the binding sites nor-
mally occupied by thiamine or its diphosphate. It was also shown that prac-
tically all of the pyrithiamine in the liver is phosphorylated. The concen-
tration in the brain increases progressively throughout the 12 days of the
experiment and it is likely that this reflects a transference of pyrithiamine
from the liver, the analog gradually replacing the thiamine in the brain.
Daily oral administration of 33 //g thiamine and 210 //g pyrithiamine leads
to a slow but very definite rise in tissue pyrithiamine over 20 days, the levels
eventually reached being higher than following the single intraperitoneal
injection. It is unfortunate that we have no data on oxythiamine distribu-
tion in the tissues, since this might well help to answer some of the problems
as to why its effects are often quite different from those of pyrithiamine.
528
2, ANALOGS OF ENZYME REACTION COMPONENTS
TIME (DAYS)
Fig. 2-18. Pyrithiaraine concentrations in rat tissues after intra-
peritoneal injection of 1 mg. (From Rindi and Perri, 1961.)
Effects on the Growth of Microorganisms
The degree of inhibition of various bacteria and fungi by pyrithiamine
has been related to the pattern of thiamine biosynthesis (Robbins, 1941;
Woolley and White, 1943 c). Sensitivity to pyrithiamine was correlated with
a requirement for intact thiamine, whereas those organisms able to syn-
thesize thiamine completely are poorly inhibited. If the organisms require
only part of the thiamine molecule, the growth depression by pyrithiamine
is intermediate. If pyrithiamine interferes with either the formation or
enzymic function of thiamine-PP, it would be difficult to understand how
the manner of obtaining thiamine could determine sensitivity to the analog.
However, another factor must be considered. It was shown that pyrithia-
mine-resistant organisms possess an enzyme capable of cleaving pyrithia-
mine, probably a thiaminase, while sensitive thiamine-requiring organisms
do not. At least part of the resistance might be attributed to the ability
of these organisms to inactivate the analog; the pyridine portion split from
pyrithiamine would not be inhibitory and the pyrimidine portion can ac-
ANALOGS OF THIAMINE 529
tually be utilized in thiamine synthesis. Supporting the importance of a
pyrithiaminase in resistance is the observation by Woolley (1944 a) that a
pyrithiamine-resistant strain of Endomyces vernalis, obtained by subcul-
turing in increasing concentrations of the analog and capable of withstand-
ing 25 times the concentration initially depressing growth 50%, contains
such an enzyme. Indeed, pyrithiamine is capable of stimulating growth in
the absence of thiamine since the pyrimidine portion (which is all that is
required by this organism) is provided by the splitting reaction. However,
destruction of pyrithiamine is not the only factor involved, since enough
of the analog remains unsplit to inhibit completely the parent strain. This
enzyme may be functional in the pathway biosynthesizing thiamine, which
would be the reason for the correlation with thiamine requirements. It might
also be well to consider another possible mechanism of inhibition, a block
of the transport of thiamine into the cells; only those organisms requiring
intact thiamine would be susceptible. A strain of S. aureus adapted to py-
rithiamine exhibits a variety of changes: the pigment color changes from
orange to lemon yellow, glucose utilization is severely depressed, and ace-
tate utilization is increased (Das and Chatterjee, 1962). A partial blocking
of the pentose-P pathway was also observed. These results indicate the
complex alterations occurring during the development of resistance.
Some of the effects on microorganisms will be briefly summarized, since
most of this work has no direct bearing on the mechanism of inhibition.
In most cases the growth depression by the analogs is counteracted by thia-
mine, as in the inhibition of Neurospora crassa by oxythiamine (Sankar,
1958), and, at least in some cases, the inhibition is formally cimpetit"'e
with thiamine (Quesnel, 1956). Growth depression can depend on various
factors. For example, Phycomyces blakesleeanus becomes more resistant to
pyrithiamine with culture age, the concentration required for 50% inhibi-
tion being 8 times greater at 13 days than at 4 days (Fluri, 1959). Is this
due to an alteration of thiamine metabolism with age, or to different me-
tabolic requirements for thiamine ? No change in sensitivity to oxythiamine
with age was noted. Furthermore, oxythiamine seemed to induce a thiamine
deficiency, determined by changes in carbohydrate content, whereas pyri-
thiamine did not. Euglena gracilis occurs in a normal green form and a
white form (chlorophyll-deficient from streptomycin treatment): The white
form is about 5 times more sensitive to pyrithiamine than is the green form
(Schopfer and Keller, 1951). Thiamine analogs have been considered as
possibly useful in certain infections. The growth of Microsporum audouini
is very strongly inhibited by 0.0012 mM pyrithiamine and the use of the
analog in tinea capitis was suggested (Ulrich and Fitzpatrick, 1951). The
infection of wheat with leaf rust (Puccinia) might be controlled with oxy-
thiamine inasmuch as this substances exerts a selective action on the fungus
when isolated infected leaves are tested (Samborski and Forsyth, 1960).
530 2. ANALOGS OF ENZYME REACTION COMPONENTS
A concentration of 0.75 n\M inhibits rust development completely and does
not exhibit phytotoxicity. Vibrio cholera is inhibited moderately by both
oxythiamine and pyrithiamine (Chatterjee and Haider, 1960), Lactobacillus
fermentum is inhibited by the imidazole analog of thiamine (Erlenmeyer et
al., 1948), and E. coli is inhibited 50% by the methylthio analog of thia-
mine at a ratio of 100 with respect to thiamine (Ulbricht and Gots, 1956).
Growth inhibition by thiamine analogs has been reviewed by Rogers (1962),
Toxic and Thiamine-Deficiency Effects in Animals
Both pyrithiamine and oxythiamine are toxic to animals and produce
states apparently related to thiamine deficiency. These analogs are of com-
parable potency; in most species pyrithiamine may be slightly more active
on a weight basis. The usual daily doses to induce the characteristic toxic
reactions and eventual death are usually between 0.01 and 0.1 mg, but this
depends on the thiamine intake, the effective ratios of analog/thiamine be-
ing around 5 to 50. The sequence of reactions following administration of
pyrithiamine to mice or rats may be summarized as: decreased food intake
(this may be noted within 24 hr), inactivity and a hunched position, ner-
vousness, tremors and occasional convulsions, spasticity followed by weak-
ness of the legs, incoordination, and paralysis. Death usually occurs within
24 hr after the development of polyneuritis. These are essentially the symp-
toms seen in thiamine deficiency but they occur more rapidly after the ana-
logs. Full polyneuritis and death may be produced within 5-12 days de-
pending on the dose. Pyrithiamine also produces typical thiamine-deficiency
polyneuritis in pigeons. The effects of oxythiamine in mice and rats are
somewhat different, although death may occur in approximately the same
time as from pyrithiamine. There is also anorexia and weight loss, and the
animals may become nervous, convulsive, and incoordinated during the
first 24 hr, but the later characteristic symptoms of polyneuritis do not
occur. Descriptions of the later reactions to oxythiamine have generally
been inadequate. In chicks apparently both analogs can induce polyneuritic
states. The above summary is derived mainly from the work of Woolley
and White (1943 b), Eusebi and Cerecedo (1949), Daniel and Norris (1949),
Frohman and Day (1949), Cerecedo et al. (1951), Naber et al. (1954), and
Wolfe (1957).
The differences between the effects of pyrithiamine and oxythiamine in
mice and rats have been emphasized by several workers, particularly the
absence of polyneuritis during treatment with oxythiamine, and have ini-
tiated speculations on the different mechanisms of action. It must be made
clear that the toxic effects of oxythiamine are not nonspecific and unre-
lated to thiamine function, since the reactions to both analogs may be
counteracted by administration of thiamine (Woolley and White, 1943 b;
Jones et al., 1948; Daniel and Norris, 1949; Cerecedo et al., 1951; and others).
ANALOGS OF THIAMINE 531
Oxypyrithiamine reduces the survival time of mice but does not produce
polyneuritis, so that it appears to behave like oxythiamine, indicating the
importance of the 4'-amino group on the pyrimidine ring for the effects on
the nervous system (Cerecedo and Eich, 1955). The 2'-w-butyl analog of
thiamine suppresses the growth of rats and leads to polyneuritis, these ef-
fects being antagonized by increased thiamine administration, so that this
analog superficially acts like pyrithiamine (Emerson and Southwick, 1945).
Another substance possibly interfering with thiamine metabolism is 2,4-
diamino-5-phenylthiazole (amiphenazole, Daptazole), a drug used as a res-
piratory stimulant. Rats on a thiamine-free diet given injections of ami-
phenazole and the pyrimidine portion of thiamine in low doses do not show
deficiency, indicating some ability to replace the normal thiazole compo-
nent, but at higher doses deficiency signs appear sooner (Shulman, 1956).
Amiphenazole alone even at high doses produces no effects. An abnormal
thiamine analog is apparently synthesized in the animals. The 2-trifluoro-
methyl analog of thiamine (trifluorothiamine) administered at 100 mg/kg/
day to mice on the thiamine-deficient diet leads to weight loss, paralysis,
convulsions, and an inhibition of the growth of transplanted carcinoma
(Barone et al., 1960). It inhibits the growth of Bacillus subtilis more po-
tently than does oxythiamine or pyrithiamine; this effect is antagonized
by thiamine, but is enhanced by either the pyrimidine or thiazole moieties
of thiamine. Further study of this interesting analog will be awaited with
anticipation.
A few miscellaneous observations relative to the pharmacological effects
of thiamine analogs on neuromuscular function will be summarized because
of the importance of such effects in developing theories of the mechanisms
of action. It has long been known that thiamine is involved in the forma-
tion of acetylcholine (providing the acetyl radical from pyruvate), and that
brain acetylcholine concentration falls during thiamine deficiency; it is pos-
sible that some of the effects of the analogs are mediated by a depression
of acetylcholine synthesis at synapses. Some have claimed that thiamine
is functional in axon conduction through its role in the synthesis of acetyl-
choline, and others have reported a release of thiamine during nerve stim-
ulation. It is also established that thiamine in rather high concentration
inhibits cholinesterase and can, under certain circumstances, augment the
action of acetylcholine. Pyrithiamine at 1-3 roM decreases the rates of de-
polarization and repolarization during the action potential in frog nerve,
whereas oxythiamine at these or higher concentrations produces no effect
(Kunz, 1956). This was interpreted as a blocking of the Na+ carrier me-
chanism and as evidence for the participation of thiamine in Na+ transport.
Depolarization is associated with Na+ entry, but repolarization in nerve is
not connected directly to Na+ flux. These concentrations are much higher
than occur following administration to animals. The tibialis twitch response
532 2. ANALOGS OF ENZYME REACTION COMPONENTS
in the cat is depressed by thiamine, pyrithiamine, and any of the pyrithia-
mine analogs having a hydroxyl group on the pyridinium ring; furthermore,
the neuromuscular blocks produced by tubocurarine and decamethonium
are antagonized by these compounds (Ngai et al., 1961). In the absence of
a hydroxyl group, there is a potentiation of twitch tension. Changes in
blood pressure parallel those in twitch tension. These compounds, of course,
may bear some relationship to acetylcholine because of the quaternary ni-
trogens and other functional groups an appropriate distance away, and it
is likely that these acute effects are unrelated to the metabolic aspects of
thiamine. It is interesting, however, that oxythiamine is without activity
on the neuromuscular junction and respiration, although it causes a fall in
blood pressure. No interference by the analogs in the actions of thiamine
was reported. Injections of thiamine, thiamine-PP, pyrithiamine, and oxy-
thiamine into frogs cause a miosis, which was interpreted as a direct effect
on the iris (Ber and Singer- Altbeker, 1961). It is possible that this is me-
diated through inhibition of cholinesterase, and it is not necessary to pos-
tulate special neural or muscular functions for thiamine. It would be more
valuable to study the possible changes in neuro-muscular activity during
administration of the analogs chronically and when there are evident motor
disturbances.
Mechanisms of Action and Comparison of Pyrithiamine and Oxythiamine
Pyrithiamine has been shown to do the following: (1) produce polyneur-
itis as in dietary thiamine deficiency, (2) deplete various tissues of thia-
mine-PP and increase its renal excretion, (3) inhibit a-keto acid metabolism
in vivo, which is reversed by adding thiamine-PP, (4) cause elevation of
blood pyruvate, (5) inhibit the phosphorylation of thiamine (thiamine ki-
nase), (6) apparently be phosphorylated in the tissues to pyrithiamine-PP,
(7) in the diphosphate form inhibit pyruvate decarboxylase, pyruvate oxi-
dation, and probably transketolase, and (8) be picked up by the tissues to
about the same extent as is thiamine normally. Most of these effects are
counteracted by the administration of sufficient thiamine. It is, therefore,
not difficult to establish possible sites of pyrithiamine inhibition; most of
the reactions to pyrithiamine can be explained on the basis of either a
block in thiamine phosphorylation or a direct inhibition of the enzymes
utilizing thiamine-PP through its diphosphate ester. If the site were only
on the kinase, the fundamental effect would be a depletion of thiamine-PP
such as occurs in dietary deficiency, and addition of thiamine-PP to tissue
preparations should restore the activity of pyruvate-metabolizing enzymes
completely. The reversal is, however, only partial (Gubler, 1961) and by
no means as great as in diet-deficient animals. It is thus likely that both
mechanisms play a role. The appearance of pyrithiamine mainly in the di-
phosphate form in tissues (Rindi and Perri, 1961) also points to the im-
ANALOGS OF THIAMINE 533
portance of direct a-keto acid enzyme inhibition. The rapidity with which
pyrithiamine exerts its toxicity (Eusebi and Cerecedo, 1949), relative to
dietary deficiency, would indicate an effect other than a block of thiamine-
PP synthesis. The exact role of thiamine kinase inhibition cannot be eval-
uated at this time.
When we turn to oxythiamine the problem becomes more complex. The
major differences from the actions of pyrithiamine may be summarized as
follows: (1) typical polyneuritis is not produced, (2) it is not as effective
in reducing tissue thiamine-PP levels, particularly in the brain, (3) it does
not produce a significant depression of pyruvate oxidation in the brain in
vivo, (4) inhibition of thiamine kinase is slight or absent, and (5) its toxic
effects are more readily overcome by thiamine. The most obvious explana-
tion would be that oxythiamine has generally the same actions as pyrithia-
mine in most tissues but for some reason does not interfere readily with
thiamine function in the nervous system. Failure to penetrate into nerve
tissue would adequately account for this but there is no direct evidence for
this, unless the failure of oxythiamine to affect the membrane potentials
of frog nerve, although pyrithiamine is effective, is interpreted in this way.
One aspect that has usually not been considered is the metabolism of these
analogs in the tissues, although Cerecedo et al. (1951) felt that oxythiamine
is more rapidly metabolized than pyrithiamine. Just as in resistant bacteria,
resistant tissues may contain a thiaminase-like enzyme capable of destroy-
ing the analog, and it is possible that a particular tissue can inactivate one
of these analogs more than the other.
If the major pathway of thiamine in animal tissues is simply (1 ) the trans-
port of thiamine into the cells, (2) its phosphorylation, and (3) the com-
bination of the thiamine-PP with the apoenzymes, the analogs must act on
one of these steps. Pyrithiamine inhibits steps (2) and (3), while oxythia-
mine affects only (3). No examination of interference with transport systems
has been reported but this should be done. The question of whether thiamine
has actions additional to its function in cc-cleavage, especially in nervous
tissue, must arise (Woolley and Merrifield, 1952; Gubler, 1961), but I doubt
if the evidence from the use of analogs is sufficient at the present time to
imply mechanisms other than on the established systems. One cannot judge
the state of a tissue function from changes in blood pyruvate (which re-
flects changes throughout the whole animal and perhaps particularly in the
liver); if central nervous system effects are to be evaluated, alterations in
the metabolism in the nerve cells must be determined. Also one must al-
ways consider the relationship between cell function and thiamine-PP level.
To what degree must thiamine-PP in the brain fall before symptoms occur?
This has often been judged by experiments in diet-deficient animals, but
in analog-treated animals it is not necessary that thiamine-PP levels be
reduced to the same degree to obtain the same functional disturbances.
534 2. ANALOGS OF ENZYME REACTION COMPONENTS
Finally, the arguments of Gubler (1961), that pyrithiamine must induce
disturbances other than in o;-keto acid metabolism, I feel are not entirely
valid. He states that since the a-keto acid oxidase activities are appreciably
lower in thiamine-deprived rat livers than in the livers of analog-treated
rats, some other disturbances in physiological function must contribute to
the deficiency symptoms and death. However, it is unlikely that the changes
in liver metabolism have much to do with either the symptoms or death,
and he actually found that the a-keto acid oxidase activities in the brain
are lower in pyrithiamine-treated rats than in thiamine-deprived rats. The
other argument, that pyrithiamine causes a polyneuritis that is difficult
or impossible to reverse by administration of thiamine whereas a-keto acid
oxidase activities can be readily restored in tissue extracts by adding thia-
mine-PP, may be significant, but these data could be just as easily explained
by an inhibition of thiamine kinase (preventing the synthesis of thiamine-
PP in the animal) or a very slow rate of exchange between thiamine and
enzyme-bound pyrithiamine-PP in the intact nervous tissue.
ANALOGS OF RIBOFLAVIN AND FAD
Riboflavin functions in metabolism as riboflavin-5'-phosphate (flavin mo-
nonucleotide, FMN) and flavin-adenine dinucleotide (FAD) in various oxi-
dizing enzymes and electron transport. The flavin coenzymes are usually
very tightly bound to their respective apoenzymes and are not dissociated
during extraction of the enzyme preparations. Indeed, in some cases, such
as succinate oxidase, the flavin component can be liberated only by pro-
teolytic digestion, with fragments of peptides attached, and the activity
cannot be restored by addition of any flavin compound. In most cases it
is thus difficult for analogs to replace or compete with the flavin coenzyme,
particularly in preparations from animal tissues, although in microorganisms
the flavoenzymes are generally more readily dissociable. The binding of
FAD seems to involve the isoalloxazine ring (perhaps the imino group at
position 3), possibly the ribityl portion, the phosphates, and the adenine
ring. Chelation to apoenzyme-bound metal ions, such as iron, is likely be-
cause most flavoenzymes contain such metal ions, but there is still some
doubt as to whether the metal ions function primarily in binding or in elec-
tron transfer.
Animals and a few bacteria depend on exogenous riboflavin but it is
synthesized in plants and most microorganisms. The pathway of riboflavin
biosynthesis is not well understood and has been studied mainly in a few
microorganisms used for the commercial production of riboflavin; the reac-
tions by which riboflavin is transformed into active coenzymes are better
documented. An abbreviated scheme of biosynthesis and breakdown is re-
presented here to facilitate discussion of the actions of analogs.
ANALOGS OF RIBOFRAVIN AND FAD
Diaminouracil
6,7-dimethyl-8-fo-l'-ribityl)-lumazine
535
Riboflavin
(3)
lumichrome + ribito'
riboflavin- 5 '-P + AMP
(1)
r
riboflavin- 5 '-P
(4)
^ riboflavin + P
(1) flavo kinase
(2) FAD pyrophosphorylase
(3) riboflavinase
(4) phosphomonoesterase
(5) nucleotide pyrophosphatase
(Reaction (6) represents a possible formation of FAD directly from riboflavin and
ATP, as postulated by Masuda (1955) in E. ashbyii; reactions (1) and (2) also require
ATP)
Many analogs, in which various parts of the riboflavin or FAD structure
have been modified, have been examined and very few are able to replace
riboflavin, possibly due both to inability to form the coenzyme analogs and
to relative inactivity of the coenzyme analogs when formed. Most of these
analogs are not significantly inhibitory to riboflavin function, indicating
the high degree of specificity in these reactions. Beinert (1960) has classified
the structural modifications producing most of the interesting analogs into
those where (1) the riboflavin is altered (as by replacement of ribitol by
other sugar alcohols or alkyl groups, replacement of the isoalloxazine ring
by other ring systems, or substitutions in the isoalloxazine ring), (2) ribo-
flavin esters other than the 5'-phosphate are synthesized (as the diphos-
phate, acetylphosphate, or sulfate), and (3) dinucleotides of riboflavin
have the AMP replaced by other nucleotides (such as GMP, IMP, CMP,
or UMP).
In this chapter we shall be concerned with the rather simple and obvious
analogs of riboflavin and FAD. There are several types of drug and me-
tabolic inhibitor that probably interfere in one way or another with ri-
boflavin metabolism or function — such as the promazines, the antibiotic
tetracyclines, the acridine antiseptics, certain antimalarials, and perhaps
some of the chelators of the phenanthroline and bipyridine types — and
these will be discussed for the most part in future chapters. Quinacrine
only will be treated in some detail at the end of this section because it is
the best studied riboflavin analog.
536
2. ANALOGS OF ENZYME REACTION COMPONENTS
HX
HX
Ribityl
H,C
Ribityl- 5^P04
Riboflavin
o
Riboflavin- 5 '-P
HX
HX
Dulcityl
1
Galactoflavin
H3C
HX
Lyxityl
NH
O
Lyxoflavin
HX
HX
HX
HX
CH2CH2OCOCH3
NH
Lumiflavin
U-2112
H3C
HX
CH,CHXCOCHXH,C(X)
^^^■>
U-6538
NH
HX
Ribityl
I
N.
NH
CH3 O
Isoriboflavin
ANALOGS OF KIBOFLAVIN AND FAD
537
Ribityl
H,CCH.
H3CCH;
6, 7 -Diethyl analog
of riboflavin
Ribityl
6-Chloro analog
of riboflavin
Ribityl
I
H,C
A^.
H NH,
2, 4-Diamino-7, 8-
dimethyl-10-
ribityl-5, lO-dihydrophenazine
Sorbityl
Flavotin
H,N
NH,
H,N
Acriflavine
(trypaflavine, euflavine)
NH,
Proflavine
CH,
N-^ ^
II
0
Toxoflavin
Effects on
Growth
A number of analogs have been found to inhibit the growth of Lacto-
bacillus casei, the standard test organism: isoriboflavin, the 6-chloro and 7-
chloro analogs of riboflavin (Lambooy, 1955), the 2,4-diaminophenazine
analog (Woolley, 1944 b), and riboflavin-5'-sulfate (Egami et al., 1956).
Other bacteria are occasionally inhibited, for example S. aureus and Str.
plantarum by dichlororiboflavin (Lambooy, 1955). The acridines, proflavine
and 5-aminoacridine, suppress the growth of L. casei but this is not coun-
teracted by increasing the riboflavin concentration, so the mechanism is
538 2. ANALOGS OF ENZYME REACTION COMPONENTS
not clear (Madinaveitia, 1946). Toxoflavin is a poisonous substance from
Pseudomonas cocovenenans and responsible for some fatal food poisonings in
Java. It quite potently inhibits the growth of E. coli, S. aureus, B. subtilis,
and Shigella, as well as being toxic to experimental animals (Latuasan and
Berends, 1961). The mechanism is unknown but it was postulated that tox-
oflavin is an effective electron-acceptor and removes electrons from the
transport chains to form hydrogen peroxide, possibly competing with the
natural flavin coenzymes (or accepting electrons from them). Relatively
little has been done on the induction of riboflavin deficiency in animals.
Isoriboflavin almost stops the growth of young rats at 2 mg/day and ribo-
flavin is able to reverse this (Emerson and Tishler, 1944), and galactoflavin
appears to produce very similar effects (Emerson et al., 1945). The 6-chloro
and 7-chloro analogs of riboflavin (Lambooy, 1955) and the dinitrophena-
zine analog (Woolley, 1944 b) produce mild deficiency states in rats and
mice. Effects have usually been determined by growth rates, and typical
symptoms of riboflavin deficiency have seldom been noted, mainly because
it requires a fairly long time to deplete the tissues of enzyme-bound co-
enzymes.
Some interesting and suggestive reports on the carcinostatic activity of
certain analogs have appeared. The 6,7-dichloro-9-(r-D-sorbityl)isoalloxa-
zine analog causes regression of mouse lymphosarcoma, the ribityl compound
has slight activity, and the other sugar alcohol derivatives are inactive
(Holly et al., 1950). Replacement of the ribityl group with nonsugar residues
gives riboflavin antagonists which inhibit L. casei and produce deficiency
states in rats. The 9-hydroxyethyl analog of riboflavin (U-2113) weakly
suppresses mouse adenocarcinoma (Shapiro et al., 1956), and the 9-aceto-
xyethyl analog (11-2112) behaves similarly (Lane et al., 1958). However,
U-2112 given to patients with various types of cancer (0.25-6 g/day for
5-84 days, with total doses 1.25-226 g) exhibits no beneficial action and
no evidence of riboflavin deficiency is seen; this was attributed to the rapid
hydrolysis of this substance in man. The 9-hemisuccinoxyethyl analog (U-
6538) does not inhibit L. casei but depresses growth in rats at 10 mg/kg/
day, which is reversible with riboflavin (Lane et al., 1959). It is quite ef-
fective against lymphosarcoma in rats and at 5 mg/kg/day leads to a 66%
inhibition of tumor growth without depressing the over-all growth rate.
Two of four patients receiving the compound showed changes suggestive
of riboflavin deficiency but no evidence of carcinostasis was observed, per-
haps due to the terminal nature of the disease and the metabolism of the
analog. Galactoflavin is known to cause tumor regression in rodents. It is
tolerated by patients at a dose of 1 g every 8 hr for 2-5 months, and de-
ficiency symptoms do not occur unless the diet is low in riboflavin (Lane
and Brindley, 1964). Presumably reports on its carcinostatic activity will
be published.
ANALOGS OF RIBOFLAVIN AND FAD 539
Metabolism of Riboflavin Analogs and Effects on Riboflavin Metabolism
Certain analogs, such as the 6,7-diethyl derivative, are able to replace
riboflavin to some extent at low concentration but are inhibitory at higher
concentration. This analog supports the growth of L. casei, and Lambooy
(1950) believed that it must be phosphorylated. This was demonstrated in
the rat where 6,7-diethylriboflavin-5'-P was found in the liver although no
FAD analog was demonstrable (Aposhian and Lambooy, 1955). L. lactis is
able to incorporate lyxoflavin into lyxoflavin-5'-P and the corresponding
dinucleotide (Huennekens et al., 1957 b). Scala and Lambooy (1958) were
able to modify L. casei by prolonged riboflavin deficiency and high con-
centrations of the 6-chloro analog so that the organism could use either ribo-
flavin or the analog. They believe that the analog inhibits the phosphor-
ylation of riboflavin. It is interesting that the adapted organism cannot
use the 7-chloro analog.
Flavokinase catalyzes the phosphorylation of riboflavin and shows a high
degree of specificity toward substrates. The yeast enzyme phosphorylates
dichlororiboflavin as well as riboflavin, arabitylflavin poorly, and all other
analogs tested not at all (including isoriboflavin, galactoflavin, dulcitylfla-
vin, and sorbitylflavin) (Kearney, 1952). The only analog that inhibits the
enzyme is lumiflavin (35% at 0.18 mM with riboflavin 0.051 mM) and this
occurs only when the analog is in excess of the riboflavin. McCormick (1962)
has extended this work to partially purified rat liver flavokinase and found
similar behavior, only riboflavin, dichlororiboflavin, and arabitylflavin be-
ing phosphorylated (all with KJs between 0.012 and 0.017 mM). Four
analogs were found to be inhibitory: lumichrome {K^ = 0.048 mM), lumi-
flavin {K^ = 0.031 mM), the 9-formylmethyl analog {K^ = 0.0097 mM),
and the 9-(2'-hydroxyethyl) analog {K^ = 0.0068 mM). The following are
not phosphorylated and do not inhibit: isoriboflavin, galactoflavin, sorbityl-
flavin, dichloroarabitylflavin, 7-methylmannitylflavin, and 7-methyldulci-
tyLflavin. The fact that most analogs are not attacked by flavokinase is per-
haps the primary reason for the failure of these compounds to replace ribo-
flavin. It is also clear that the data are insufficient to draw conclusions
relative to the possibility of some of the most commonly used analogs inhi-
biting the phosphorylation of riboflavin, but what evidence we have would
indicate that such inhibition is unlikely to be important. The synthesis of
riboflavin from 6,7-dimethyl-8-(r-D-ribityl)lumazine by an enzyme system
from Ashbya gossypii is potently inhibited by a variety of analogs of this
precursor, of which the 6,7-dihydroxy derivative is the most active {K^ =
0.000009 mM) (Winestock et al, 1963). It is interesting that 5'-deoxyribo-
flavin is fairly inhibitory {K, = 0.019 mM) and, indeed, it was concluded
that the sugar moiety is necessary for inhibition.
Very little information on the effects of analogs on the tissue levels of
riboflavin compounds is available. Rats given galactoflavin for 10-28 days
540 2. ANALOGS OF ENZYME REACTION COMPONENTS
show a 70-75% depression of liver mitochondrial flavin, and dietary ribo-
flavin restriction also reduced the level (Beyer et al., 1961). The evidence
from depression of enzyme activity will be discussed later. The thorough
analysis of the effects of riboflavin deficiency on rats by Burch et al. (1956)
has shown that various tissues differ markedly in ability to retain FMN
and FAD (see accompanying tabulation). Deficiency of 23-day duration
% Change in deficient rats
Tissue
FMN
FAD
NADH oxidase
D-Amino acid
oxidase
Xanthine
oxidase
Brain
-24
-19
0
+ 8
Liver
-86
-61
+ 10
-66
-22
Kidney
-41
-19
-11
-17
-19
Heart
0
-32
2.3
1 (•)
0
has little effect on brain flavins while liver levels drop rapidly. No necessary
correlation between total FMN or FAD and enzyme activity is evident,
indicating that some enzymes will lose their FMN or FAD much more
readily than others and that a fraction of the cellular flavin may be nonen-
zymically bound. On the basis of these results, one might expect riboflavin
analogs, when active, to exert differential effects on the various flavoen-
zymes, and it is likely that analyses for total flavins will not provide me-
tabolically significant figures.
Effects on Flavoenzymes
The enzymes most commonly used to test the inhibitory activity of
riboflavin analogs are those with dissociable flavin coenzymes, such as the
old yellow enzyme and D-amino acid oxidase, and in such cases an inhibi-
tion of a competitive nature is not unexpected. However, there are many
instances of the inhibition of enzymes which have very tightly bound co-
enzymes and these are more difficult to interpret. Some inhibitions of both
types are shown in Table 2-33. Inhibitions by riboflavin, FMN, and FAD
are also included because these show that it is not alwaj^s necessary to
consider inhibition as resulting from structural analogs.
Most of these inhibitions appear to be noncompetitive. Thus the inhibi-
tions of L-galactono-y-lactone dehydrogenase by riboflavin, L-amino acid
oxidase by riboflavin and its analogs, and D-amino acid oxidase by ribo-
flavin are not reduced by increasing concentrations of FMN or FAD. How-
ever, the inhibitions of kidney D-amino acid oxidase by FMN and ribo-
flavin-5' -sulfate are competitive with respect to FAD, and the inhibition
ANALOGS OF RIBOFLAVIN AND FAD
541
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2. ANALOGS OF ENZYME REACTION COMPONENTS
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544 2. ANALOGS OF ENZYME REACTION COMPONENTS
of glutamate racemase by riboflavin is reduced by FAD. It is clear that
there is not much information on the inhibition of enzymes by riboflavin
analogs, especially by their phosphates, or FAD analogs. The mechanism
for the inhibition of enzymes in which the flavin coenzyme is tightly bound
is not known. However, it is possible to suggest three mechanisms. It is
now known that various flavins and their nucleotides form molecular com-
plexes with one another, and the formation of such complexes with bound
FMN or FAD may occur, preventing the normal interactions of the coen-
zymes in oxidation. In some instances the inhibitors may interfere with
the experimental electron acceptor, particularly when this is a dye. Lastly,
one must consider the possibility of nonspecific binding of these polyhetero-
cyclic compounds to the enzymes; one might predict that a number of en-
zymes not involving flavins would be inhibited by such analogs, but few
have been examined. None of the riboflavin analogs in Table 2-33 is a po-
tent inhibitor and it is unlikely that these inhibitions are responsible for
any of the in vivo effects observed.
The coenzyme of the old yellow enzyme is FMN, and riboflavin-5'-sulfate
does not interfere with its binding to the apoenzyme, which Theorell et al.
(1957) explain by the less negative charge on the sulfate group. Riboflavin-
5'-sulfate, however, inhibits D-amino acid oxidase (Egami and Yagi, 1956),
so that the structural requirements for binding must be different in these
two enzymes. Yagi and Nagatsu (1960) have studied the effects of ribofla-
vin-5'-sulfate on rat liver mitochondrial oxidations of a-ketoglutarate, suc-
cinate, malate, and D-alanine, and found that no inhibition is exerted at
0.1 mM, which they interpret as due to the tight binding of the FAD in
the mitochondria. Aged mitochondria are stimulated by FAD and here in-
hibition by riboflavin-5' -sulfate can be demonstrated. The FAD analogs of
the various flavins have not been studied often but Huennekens et al.
(1957 b) found lyxoflavin-5'-phosphate to be active in the NADPH-cyto-
chrome c reductase (although less than FMN) and lyxoflavin dinucleotide
to be active in the D-amino acid oxidase (but less than FAD). We have
seen that riboflavin deficiency leads to reduction in the activities of certain
enzymes in the tissues. Administration of galactoflavin to rats for 15-28
days leads to an approximately 40% reduction in glutamate and /5-hy-
droxybutyrate oxidation in liver mitochondria, but no change in succinate
oxidation or in the P : 0 ratios (Beyer et al, 1961). U-2113, the 9-hydrox-
yethyl analog of riboflavin, causes a slight (15%) decrease in tumor xan-
thine oxidase in mice. It is not known if this is due to FAD depletion or a
more direct inhibition. 5-Hydroxytryptamine (serotonin) is metabolized by
monoamine oxidase to 5-hydroxyindoleacetate; both riboflavin deficiency
and galactoflavin increase the urinary excretion of this product, indicating
that one of the other metabolic pathways for serotonin is depressed by
interference with flavin function (Wiseman and Sourkes, 1961). These mis-
ANALOGS OF RIBOFLAVIN AND FAD 545
cellaneous observations do not provide a satisfactory basis for understand-
ing the metabolic effects of riboflavin analogs.
We have been discussing analogs of the riboflavin portion of FAD and
some mention of the adenine nucleotides as inhibitors should be made. The
D-amino acid oxidase of sheep kidney is inhibited competitively by various
purines and nucleotides (see accompanying tabulation) (Burton, 1951 a).
Inhibitor
{mM)
5'-AMP
1.05
S'-AMP
No inhibition
ADP
13
ATP
11
Adenosine
45
Adenine
22
Hypoxanthine
24
Caffeine
11
It was shown that complexes between riboflavin and purines are formed
and have the following dissociation constants: caffeine 10 n\M, adenosine
30 mM, AMP 40 mM, ADP 37 mM, and ATP 39 mM. The formation of
such complexes might account for the enzyme inhibition in the case of
adenosine and caffeine, but cannot account for the more potent effects of
AMP and ADP, these latter substances competing with FAD for the apo-
enzyme site. The D-amino acid oxidase from pig kidney is likewise inhibited:
50% inhibition is given by 0.4-0.6 mM AMP, ADP, ATP, and IMP; by
6 mM adenine, adenosine, and hypoxanthine; and by 15-20 mM uracil, cyto-
sine, and ribose-5'-P (FAD 0.00014 mM in all cases) (Walaas and Walaas,
1956). The K, for the competitive 5'-AMP is 0.64 mM. Crandall (1959)
determined K^ for AMP as 0.1 mM for this enzyme. Flavokinase is inhi-
bited strongly by 5'- AMP {K^ = 0.025 mM), and it is possible that the
adenine portion of the nucleotide competes with the alloxazine ring of ribo-
flavin for the active center (Kearney, 1955).
Quinacrine (Mepacrine, Atabrine, Atebrin)
Quinacrine is an acridine derivative introduced by the Germans in 1932
for malaria therapy as a suppressive agent and is more effective than qui-
nine on the asexual forms of the plasmodia. Since the observation by Wright
and Sabine (1944) that FAD counteracts the inhibition of tissue respiration
by quinacrine, it has been commonly assumed that quinacrine exerts its
546 2. ANALOGS OF ENZYME REACTION COMPONENTS
primary effects by interference with flavoenzymes, and has come to be the
most widely used substance to detect the participation of a flavin com-
ponent in an enzyme or metabolic system. Quinacrine not only inhibits the
CH,CH,CH3-N^^'^'
CH,0
Quinacrine
malarial organism but, in common with other acridines, suppresses the
growth of various bacteria. For example, Lactobacillus casei is inhibited
and this seems to be related to flavin metabolism since the maximal con-
centration at which growth occurs is 0.12 niM when riboflavin is 0.00066 mM
and 0.49 mM when riboflavin is 10 times the previous concentration (Ma-
dinaveitia, 1946). Spore germination of B. subtilis (Falcone et at., 1959) and
B. coagvlans (Amaha and Nakahara, 1959) induced by L-alanine is inhibited
58% by 0.1 mM quinacrine and nearly completely by 1 mM.
The toxic efl^ects observed in experimental animals and man — for ex-
ample, gastrointestinal (abdominal pain, diarrhea, nausea), dermatological
(eczematoid dermatitis, lichen planus), central nervous system (psychoses),
and others — do not appear to be related to riboflavin deficiency, and the
typical syndrome of deficiency has never been produced by quinacrine.
Thus one must assume that other actions are probably of more importance
in animals. One characteristic of quinacrine is its remarkable ability to be
accumulated in the tissues during chronic administration, and eventually
the tissue levels are hundreds or thousands of time higher than in the serum
(see Table 1-8-1). This is evident from the yellow coloration of the tissues.
These levels, of course, do not represent free quinacrine and the accumula-
tion is due to the high affinity of various tissue components for quinacrine.
It is bound in the cytoplasm, the mitochondria, and the nucleus; equilibra-
tion of isolated nuclei with quinacrine results in a 200-fold concentration
differential (Reiner and Gellhorn, 1956). Proteins, nucleic acids, and nu-
cleoproteins bind quinacrine strongly, and some of the inhibitory, muta-
genic, and carcinostatic effects have been attributed to such binding. It
is thus clear that quinacrine can be bound at many loci in the cell.
Effects of Quinacrine on Enzymes
Many enzymes are inhibited by quinacrine (Table 2-34). In some cases
the inhibition is competitive (or at least reduced by increasing the concen-
tration of FMN or FAD) and in others it is not. Quinacrine has come to
ANALOGS OF RIBOFLAVIN AND FAD 547
be the most commonly used detector for the participation of flavins in
enzyme systems, but before this is subjected to analysis we shall discuss
some of the results which have been obtained. The following summary pre-
sents the data reported on antagonism but does not in any case imply a
truly competitive mechanism.
Enzymes in which inhibition is reduced by FMN or FAD
Adenosinetriphosphatase (mitochondrial NAD-activated): FMN and FAD
(Low, 1959)
Aldehyde oxidase (rat and monkey liver): FAD (Lakshmanan et al., 1964;
Mahadevan et al, 1962)
Aliesterase (liver): FMN (Hemker and Hiilsman, 1960)
D- Amino acid oxidase (lamb and sheep kidney): FAD (Hellerman et al.,
1946; Burton. 1951 a)
Catechol oxidase (spinach): FAD (Nair and Vining, 1964)
Choline dehydrogenase (rat liver): FAD (Bargoni, 1963)
Cytochrome reductase: FMN (Haas, 1944)
Hydroxylamine: cytochrome c oxidoreductase {Nitrosomonas): FMN and
FAD (Aleem and Lees, 1963)
Lactate dehydrogenase {Lactobacillus and yeast): FMN and FAD (Snos-
well, 1959; Iwatsubo and Labeyrie, 1962)
NADH: nitrite oxidoreductase (Neurospora): FAD (Nicholas et al., 1960)
NADH oxidase {Azotobacter, Clostridium, and Lactobacillus): FAD (Re-
paske and Josten, 1958; Dolin, 1959; C. F. Strittmatter, 1959)
Nitrate reductase (Pseudomonas): FAD (Fewson and Nicholas, 1961)
Succinate oxidase {Tetrahyrnena and Xanthomonas): FMN and FAD (Ei-
chel, 1956 b; Madsen, 1960)
Enzymes in which inhibition is not reduced by FMA or FAD
Adenosinetriphosphatase (mitochondrial sonicate) (Beyer, 1960)
Allohydroxy-D-proline oxidase {Pseudomonis) (Yoneya and Adams, 1961)
L-Amino acid oxidase (moccasin venom) (Singer and Kearney, 1950)
Ethanolamine oxidase (Arthrobacter) (Narrod and Jakoby, 1964)
L-Galactono-5^-lactone dehydrogenase (cauliflower) (Mapson and Bres-
low, 1958)
Lactate dehydrogenase (Propionibacteritim) (Molinari and Lara. 1960)
Nitrate reductase {E. coli) (Heredia and Medina, 1960)
Old yellow enzyme (yeast) (Kistner, 1960)
548
2. ANALOGS OF ENZYME REACTION COMPONENTS
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556 2. ANALOGS OF ENZYME REACTION COMPONENTS
Some of the enzymes inhibited by quinacrine do not involve flavin co-
enzymes, as emphasized by Hellerman et al. (1946) and Hemker and Hiils-
man (1960), so that no direct antagonism with FMN or FAD would be
expected. Nevertheless, reversal of the inhibition is sometimes seen and
may be due to the formation of a complex between the quinacrine and the
added coenzyme, as was shown to occur between quinacrine and FMN in
the inhibition of aliesterase, this presumably removing some of the quina-
crine from the enzyme. In any case, the degree of reversal or even whether
reversal occurs with FMN or FAD will depend on the way in which the ex-
periment is run and the relative concentrations. If interaction of the en-
zyme and quinacrine is allowed to occur, the chance of reducing the inhi-
bition by adding coenzyme is less than if both are added together, since it
may be difficult to reach equilibrium due to the tight binding of quinacrine.
The effects of quinacrine on ATPases and oxidative phosphorylation are
interesting and perhaps important in explaining some of the metabolic
changes. Myofibrillar ATPase does not depend on any flavin component
and yet is inhibited rather well by quinacrine (Kaldor, 1960). Furthermore,
increasing the ATP concentration from 1 raM (where 1 mM quinacrine in-
hibits about 50%) to 7 vaM almost abolishes the inhibition. The inhibition
is potentiated by Mg++ and counteracted by Ca++, so that a quinacrine-Mg
complex was postulated as the possible active inhibitor. Interactions of
Mg++ and ATP with quinacrine were shown by fluorescence changes. Irvin
and Irvin (1954) had found that quinacrine forms complexes with AMP
and ATP at physiological pH's. The dissociation constant for the ATP com-
plex is 1.38 X 10"^, so that 1 raM quinacrine would lower the ATP con-
centration from 1 vaM to 0.67 raM, whereas it would have a negligible ef-
fect on ATP around 7 raM. Increasing the ATP might also reduce the Mg++
available for a quinacrine complex, and if this is the active inhibitor, the
inhibition would be lessened. The ATPase activity in mitochondrial pre-
parations may be quite different from the myofibrillar enzyme and could
involve a flavin component. Quinacrine inhibits the ATPase of beef heart
mitochondria and simultaneously uncouples oxidative phosphorylation even
more potently (Penefsky et al., 1960), while the DNP-stimulated ATPase
of rat liver mitochondria is stimulated by quinacrine at lower concentra-
tions (0.75 mM) and inhibited by higher (3 raM) (Low, 1959 a), the P,-
ATP exchange reaction and oxidative phosphorylation being depressed.
Low felt that this provides evidence for the participation of flavin in such
ATPase activity, especially, as FMN and FAD can reverse the inhibition,
but other interpretations are possible (e.g., complexes between quinacrine
and ATP or the flavins). There is no doubt, however, that DNP alters the
response of mitochondrial ATPase to quinacrine, but no DNP effect is ob-
served in muscle ATPase (Pennington, 1961). Uncoupling by quinacrine
was first reported by Loomis and Lipmann (1948) and recent work seems
ANALOGS OF RIBOFLAVIN AND FAD 557
to establish a true uncoupling action, although by no means so specific as
with DNP since Og uptake is usually reduced simultaneously with the
P : 0 ratio.
Baltscheffsky (1960 b) found that light-induced phosphorylation in spin-
ach chloroplasts is strongly inhibited by quinacrine, 0.04 mM producing
almost complete block, and in cell-free extracts of Rhodospirillum rubrum
less potently (Baltscheffsky and Baltscheffsky, 1958; Baltscheffsky, 1960 a).
The inhibition is reversed by FMN and FAD in the the bacterial extracts,
but not at all in the chloroplasts; indeed, in the latter FMN and FAD are
quite potent inhibitors. It was suggested that an endogenous flavin is a
necessary component of the system. Photophosphorylation has recently been
found to be very sensitive to quinacrine. In Rhodospirillum chromatophores
quinacrine begins to depress the photophosphorylation at 0.0001 mM, in-
hibits 65% at 0.028 mM, and blocks completely at 0.1 mM, the K, being
0.003 mM (Horio and Kamen, 1962 a). The characteristic response to ribo-
flavin is, however, not prevented and it was postulated that quinacrine
binds at some locus in the respiratory chain. Quinacrine at concentrations
around 0.05 mM uncouples all types of photophosphorylation in Swiss chard
chloroplasts and simultaneously stimulates the photoreduction of dichloro-
phenolindophenol (Gromet-Elhanen and Avron, 1963). Similar effects were
observed in spinach chloroplasts, quinacrine at 0.02 mM inhibiting photo-
phosphorylation 61% and at 0.05 mM inhibiting completely, at the same
time stimulating the photoreduction of trimethyl-l,4-benzoquinone (Dilley
and Vernon, 1964). Changes in light absorption and scattering indicate a
relationship between photosphosphorylation and structural alterations in
the chloroplasts, but it is not known if quinacrine modifies directly these
structural changes.
The determination of the inhibitor constant, K„ for quinacrine and sim-
ilar substances is somewhat more complex than with most inhibitors, due
to the fact that equilibrium is difficult to achieve and mutual depletion
kinetics must be applied, the free concentrations of both quinacrine and
FAD being much lower than the total concentration. HeUerman et al. (1946)
considered these problems relative to the inhibition of D-amino acid oxidase
and described a very useful technique with the appropriate equations for
the calculation of K^, The ^fad is 0.00057 mM, and K^ for quinine is
0.67 mM (means for two enzyme preparations). The quinine inhibition is
quite competitive but quinacrine behaves atypically and its K^ varies with
the experimental conditions (it is somewhat smaller than the K^ for qui-
nine).
An interesting example of the effects of pH on inhibition was reported
by Molinari and Lara (1960) for the lactate dehydrogenase of Propionihac-
terium pentosaceum (Fig. 2-19). Increase of pH augments the inhibition by
quinacrine whereas the opposite effect is seen on Dicumarol inhibition,
558
2. ANALOGS OF ENZYME REACTION COMPONENTS
but these relationships are reasonable if one assumes that the negatively
charged Dicumarol reacts with positively charged enzyme groups, and the
positively charged basic quinacrine reacts with enzyme anionic groups.
We must finally evaluate the reliability of quinacrine as an indicator of
flavin participation in enzyme reactions. Certainly the mere inhibition of
an enzyme by quinacrine does not imply involvement of a flavin coenzyme.
100
80
60
40
n 20
%
INH
Fig. 2-19. Effects of pH on the inhibitions of lactate dehydrogenase
from Propionibacterium pentosaceum by quinacrine and Dicumarol
at 0.1 mM. (From Molinari and Lara, 1960.)
The observations of Hemker and Hiilsman (1960) support the opinion of
Hellerman et al. (1946) that quinacrine is a relatively nonspecific inhibitor,
due to its affinity for proteins in general. If a reversal of the inhibition by
FMN or FAD is demonstrated, there is more likelihood for the participation
of a flavin, but even here one must consider the possible complexing of the
quinacrine by the reversing flavin. Also it seems that flavin nucleotides
which are not coenzymically active are often as good reversors as the co-
enzyme itself, indicating some other mechanism than competition for the
active center. It would seem to me that quinacrine would be one of the least
likely specific antagonists of FMN or FAD, since structurally it is not as
close as many other analogs. An ideal indicator for flavins would be phos-
ANALOGS OF RIBOFLAVIN AND FAD 559
phorylated or in the form of a dinucleotide analog, and such has not yet
been found.
Many flavin-dependent enzymes bind FAD or other flavin coenzymes very
tightly and it is difficult to understand how quinacrine could displace these,
or how exogenous FMN or FAD could antagonize the action of quinacrine
if this is the case. If an enzyme after extraction is flavin-dependent and is
catalytically active, it must have very tightly bound coenzyme; perhaps
quinacrine can react with the bound coenzyme but there is no evidence for
this. Certainly the failure of quinacrine to inhibit should not be taken as
evidence for the absence of a flavin component. Actually it must be said
that many of the experiments with quinacrine have not been done properly.
In some instances one concentration of quinacrine has been used and, if
inhibition of any degree is noted it is stated that this is evidence for a
flavoenzyme, even though no antagonism has been demonstrated; it should
be obvious that no conclusions can be drawn from these results. In other
cases two experiments have been run, one with quinacrine alone and one
with both quinacrine and either FMN or FAD; if the inhibition is less in
the presence of the FMN or FAD it is concluded that quinacrine is inter-
fering with the normal function of a flavin coenzyme. A control with FMN
or FAD alone must also be run, since in many cases these substances will
stimulate activity. The results of Bargoni (1963) are difficult to interpret
in that FAD would prevent the inhibition by quinacrine only if the FAD
were incubated with the enzyme for 30 min before the inhibitor is added.
The binding of quinacrine to some enzymes is readily reversible, but yeast
lactate dehydrogenase is irreversibly inactivated; FAD will slow this inac-
tivation but will not reactivate (Iwatsubo and Labeyrie, 1962). Several
suggestions as to the design of such antagonism tests may be made: (1) use
a flavin derivative which is the most likely coenzyme involved, (2) use co-
enzyme-dissociated and reconstituted enzymes whenever possible, (3) al-
ways have a control with the reversor alone, and (4) attempt to establish
competitive relations between the quinacrine and the reversor.
Effects of Quinacrine on Metabolism
Quinacrine and other antimalarials were found by Fulton and Christo-
phers (1938) to depress the respiration of trypanosomes. Concentrations as
low at 0.004 mM exert inhibitory effects on multiplication but it requires
0.3 mM to inhibit the respiration 11.5%, at which concentration the count
is reduced 26.3%. These observations and several others afterward dem-
onstrate that concentrations presumably higher than are present in vivo
must be used to depress the respiration of these organisms. Wright and
Sabine (1944) studied the effects of quinacrine on the respiration of rat
tissue slices. In most instances there is an initial stimulation followed by a
slowly developing inhibition, often not complete after 100-200 min. Liver
560 2. ANALOGS OF ENZYME REACTION COMPONENTS
slice respiration is depressed only slightly by 0.5 n\M, even after 2 hr,
while 1-2 mM produces a maximal depression of around 65% after 1 hr.
Brain respiration is more sensitive and is reduced around 75% by 0.5 mM.
Following inhibition by quinacrine, addition of pyruvate, lactate, citrate,
fumarate, or malate does not restore oxygen uptake, but addition of succi-
nate brings about a rapid rise in respiration, indicating only that succinate
oxidase is not blocked significantly at these concentrations. Because of the
reversal of D-amino acid oxidase inhibition by coenzyme, they suggested
that the block may be around the flavoenzyme locus in the respiratory
chain, but actually there is no evidence for this. The respiration of Plasmo-
dium lophurae with different substrates is depressed 15-24% by 0.1 mM
quinacrine and somewhat above 50% by 1 mM (Bovarnick et al., 1946).
Actually it is very difficult to compare in vitro and in vivo effects and con-
centrations because of the progressive binding and accumulation of quina-
crine; in other words, the free plasma concentration of quinacrine means
very little, nor does total tissue concentration necessarily relate to any
enzyme effects.
A glycolytic inhibition by quinacrine was suggested by the early work
of Marshall (1948), who found a depression of glucose utilization, a decrease
in glucose- 1-P, an increase in glucose-6-P, some decrease in triose-P's, and
decreases in pyruvate and lactate in washed chick erythrocytes parasitized
by Plasmodium gallinaceum. It is difficult to separate the metabolic effects
on parasite and erythrocyte, but it is probable that the major fraction of
the glucose utilization was due to the parasites. The most marked effect of
quinacrine is an accumulation of ATP, which Marshall attributed to an in-
hibition of hexokinase.
The only analysis of the effects of quinacrine on metabolism was made
by Bowman et al. (1961). The glucose utilization of P. bergkei free parasites,
parasitized reticulocytes, and reticulocytes was determined, and low con-
centrations of quinacrine (claimed to be near those found therapeutically)
exhibit a selective action on the parasites. The glucose utilization over 1 hr
is reduced 34% by 0.0125 mM and 94% by 0.035 mM quinacrine. There
is no effect on the pattern of glucose-1-C^* and glucose-6-C^^ distribution.
The amount of lactate formed from glucose is reduced and there is an accu-
mulation of hexose-6-P, so it was concluded that quinacrine inhibits some
enzyme which is involved in the utilization of hexose-6-P and is normally
rate-limiting in the free parasites; this enzyme may be phosphofructokinase.
Depression of respiration could thus, in part, be attributed to a glycolytic
inhibition and, if so, is probably not related to a flavoenzyme.
There is probably need for more investigation of the effects of quinacrine
on tissues and parasites in animals given the drug for varying times, be-
cause of the difficulty in estimating the proper in vitro concentrations to
use. It may well be that some enzyme system not previously examined is
ANALOGS OF PYRIDOXAL 561
most potently inhibited. With regard to the inhibition of growth, it might
be well to consider more carefully the changes resulting from complexes
formed between quinacrine and nucleotides or nucleic acids.
ANALOGS OF PYRIDOXAL
A group of substances, including pyridoxol, pjTidoxal, pyridoxamine, and
their phosphates, possess vitamin Bg activity and these will be designated
as pyridoxine in accordance with Braunstein (1960) and the Commission
on Chemical Terminology of the International Union of Pure and Applied
Chemistry (the substance previously called pyridoxine now being pyridoxol).
These substances are converted to pyridoxal which is metabolically func-
tional in the form of its phosphate. Pyridoxal-P is the coenzyme for a large
number of enzymes involved in the decarboxylation, transamination, oxi-
dative deamination, racemization, a,/?-cleavage, and /?- and y-substituent
replacement in amino acid metabolism, and, in addition, may be active in
amino acid transport. Disturbances in pyridoxal metabolism or functions
will thus bring about primarily alterations in the biosynthesis and degra-
dation of amino acids, and indirectly will affect protein synthesis and a
variety of other metabolic pathways. Possibly the most important biochem-
ical defect will be the reduction in transaminations involving glutamate,
inasmuch as these reactions are central in amino acid metabolism. More
recently it has been found that phosphorylase a contains pyridoxal-P, per-
haps bound in an aldamine linkage, and, although initially it was believed
that it is enzymically nonfuctional, the demonstration by Illingworth et al.
(1958) that pyridoxal and 5-deoxy pyridoxal will prevent the binding of
pyridoxal-P and enzyme activity points to some role of the pyridoxal-P
in the catalysis.
Animals generally require pyridoxine whereas plants and most micro-
organisms can synthesize pyridoxal. Little is known about the biosynthesis
of pyridoxal, but the rather complex interrelationships between the pyri-
doxines and their phosphates are now fairly clear. The pathways and the
enzymes involved are summarized in the accompanying diagram (Braun-
stein, 1960; Wada and Snell, 1961). It is believed that the major pathway
for the formation of pyridoxal-P is
Pyridoxol -^ pyridoxol-P -> pyridoxal-P
The primary excretory metabolite of pyridoxal is 4-pyridoxate and its lac-
tone. Most of the pyridoxine in tissues is present as pyridoxal-P bound quite
tightly to enzymes and other proteins. Certain analogs can inhibit the for-
mation of pyridoxal-P and may act partly in this way, while other analogs
may be phosphorylated and compete with pyridoxal-P.
562
2. ANALOGS OF ENZYME REACTION COMPONENTS
(2) ^
Pyridoxol
4-pyridoxate
(6)
(1)
r
pyridoxal
(5) (4)
(3)
(2)
pyridoxol- P
(5)
pyridoxamine ^^
(3)
(2) ,
pyridoxal -P
(5)^
(4)
pyridoxamine- P
(1) pyridoxol oxidase
(2) pyridoxal kinase
(3) phosphatase
(3)
(4) transaminase
(5) pyridoxol- P oxidase
(6) aldehyde oxidase
Two general types of analog are theoretically possible — those resulting
from alteration of the pyridine ring and those in which the substituent
groups are modified, replaced, or eliminated — but practically it has been
found thus far that only analogs of the second type are effective. Actually,
not many really effective analogs have been found. The most commonly
used analog has been deoxypyridoxol* and we shall limit our discussion
mainly to this substance. Unless otherwise noted, the name deoxypyridoxol
will refer only to the 4-derivative.
Deoxypyridoxol was found to have no vitamin Bg activity by Unna (1940)
and to be an antagonist of pyridoxine in the chick by Ott (1946). Chicks
on a low pyridoxine diet can be killed by as little as 16 //g deoxypyridoxol,
whereas normal chicks on an adequate pyridoxine diet can withstand as
much as 600 //g. By varying the relative doses of both vitamin and analog,
Ott showed that approximately 2 molecules of analog can counteract 1
molecule of pyridoxine. Deoxypyridoxol has since been found to inhibit cer-
tain microorganisms and to produce symptoms of vitamin Bg deficiency in
animals, including man. 4-Methoxymethylpyridoxol (usually called methox-
ypyridoxine) was found by Unna to have slight vitamin activity in the
rat, but Ott (1947) demonstrated a potent inhibitory effect in the chick.
The ability of rats to use this analog is related to its transformation to
pyridoxal in these animals (Porter et al., 1947). In the chick it is about 25
times as effective as deoxypyridoxol. These are apparently the only analogs
so far tested that can produce rather typical pyridoxine-deficiency symp-
toms in animals, although several others can inhibit microbial growth by
disturbing pyridoxal function. Toxopyrimidine is undoubtedly toxic to ani-
mals and can be antagonized by pyridoxine, but it is debatable whether
* This is 4-deoxypyridoxol and has previously been called desoxypjTidoxine or
deoxypyridoxine. However, if we are to conform to the modern nomenclature, the
specific compound must be deoxypyridoxol. Deoxypyridoxine might be used to refer
to the entire group of deoxy substances exhibiting vitamin Bg activity antagonism.
ANALOGS OF PYRIDOXAL
563
CH20H
CHO
CHO
H0^J:;;;^^CH20H
HO.^^4^^^
CH2OH
HO ^^^Av^ CHjOPd;
H3C'^N+
^ H+
Pyridoxol
(pyridoxine)
Pyridoxal
Pyridoxal-P
CH2NH3"
HO^ >\ ^CH,OH
H,C N.
CHjOH
CH2OH
H3C N
H
CH3
HO^^X/CH^OH
XJ
H3C N'
Pyridoxamine
3-Deoxypyridoxol
4-Deoxypyridoxol
(deoxypyridoxine)
CH2OH
HO^J^CH3
CH2OCH3
H0^A^.^CH30H
•XJ
H^
CH2OH
HO, X^ ^CH,OH
H,C— CKT ^NI
5 - Deoxypy r idoxo 1
4 - Methoxy methyl -
pyridoxol
w-Methylpyridoxol
CH2OCH2CH3 CH=NOH
^^^..^^'Cv^CHaNHa HO^ JX /CH,OH
--L y^
H,C— CH, Nl
3 -2 JJ+
"XT'
H^
nh:
H3C "N
H
CH2OH
2-Ethyl-3-amino-
4-ethoxyniethyl-5-
aminomethylpyridine
Pyridoxal oxime
CHO
Toxopyrimidine
CH3
HOv. A^ /OH
HaC^ Te
NO,
4-Nitrosalicylaldehyde , 2, 4-Dimethyl- cvcjo-
•' ^ telluropentane-3, 5-dione
564 2. ANALOGS OF ENZYME REACTION COMPONENTS
it can be termed a true pyridoxine analog; this substance will be discussed
separately at the end of this section.
Effects on Pyridoxine Metabolisnn and Tissue Levels of Pyridoxine
The only analog found in the early work to inhibit pyridoxal kinase strong-
ly is 2-ethyl-3-amino-4-ethoxymethyl-5-aminomethylpyridine (Hurwitz,
1952). This substance cannot be phosphorylated because of the lack of a
hydroxyl group at position 5, but has a rather high aJBinity for the yeast
enzyme {K^ = 0.073 mM) and competitively inhibits the phosphorylation
of pyridoxal. Certain analogs can be phosphorylated by this enzyme (e.g.,
deoxypyridoxol, 3-deoxypyridoxol, and the 3-amino analog of pyridoxol)
(Umbreit and Waddell, 1949; Hurwitz, 1955 b), and presumably could re-
duce the phosphorylation of pyridoxal through substrate competition. Phos-
phorylation was shown to require a hydroxymethyl group at the 5-position
and the absence of a substituent at position 6. The more recent work of
McCormick and Snell (1961 ) demonstrated other potent inhibitors and made
it clear that the kinases from different sources vary markedly with respect
to affinity for the analogs (Table 2-35). Furthermore, the relative affinities
are fairly well correlated with the abilities to inhibit growth of the various
bacteria. The inhibitions are not always competitive and, in some instances,
increase with pyridoxal concentration. The 3-hydroxy group can be replaced
by an amino group or omitted without affecting affinity adversely, but
substitution in the 6-position reduces the affinity without necessarily abol-
ishing it. It is interesting that A^-methylpyridoxal is completely inactive as
an inhibitor. The most potent inhibitors of pyridoxal kinase are derivatives
obtained by the reaction of pyridoxal with various carbonyl agents. The
beef brain kinase is inhibited 50% by 0.00005 mM pyridoxal semicarbazone
and by 0.000065 mM pyridoxal azine (the product of the reaction of 2
pyridoxals with hydrazine), but the discussion of such inhibitions is more
pertinent to the subject of the carbonyl agents.
Various enzymes oxidizing pyridoxol, pyridoxol-P, or pyridoxamine-P
have recently been found in liver, and are occasionally inhibited by analogs.
The oxidation of pyridoxol is competitively inhibited by deoxypyridoxol
(67% when pyridoxol is 10 mM and the analog is 12.5 mM) (Morino et al.,
1960), while the oxidation of pyridoxol-P is competitively inhibited by
deoxypyridoxol-P {K„, = 0.02 mM, and K, = 0.35 mM) (Morisue et al,
1960). These enzymes thus follow the general rule that nonphosphorylated
analogs inhibit the reactions of nonphosphorylated substrates, and phos-
phorylated analogs inhibit the reactions of phosphorylated substrates. The
oxidative deamination of pyridoxamine-P is inhibited by pyridoxamine (the
latter is also deaminated at a slower rate) and rather weakly by pyridoxol
(Pogell, 1958). Wada and Snell (1961) examined the competitive inhibitions
of pyridoxol-P oxidase by a variety of substances (see accompanying tab-
ANALOGS OF PYRIDOXAL
565
PQ
^ o
d d
d d
2Q
o o
d d
in
O lO
d -«
^
-r -A >>
o
o
o
n
';-i
^
>•.
>i
p
fO
Ph
ft
^^
^
>■-
>.
t>>
>■.
X
X
^
f^
o
O
IB
1
-*
lO
3
<N
-a
P-i
o
«
o
^
r2
'E
^
>>
o
ft
^^
t3
,
C
"«
eS
^.^
CO
c3
X
-c
O
c
'2
cS
'E
^
ft
•p
t-i
c
o
(->
Ch
a
in
0)
_2
a
>
2
«
^
&^
H
566
2. ANALOGS OF ENZYME REACTION COMPONENTS
% Inhibition of oxidation of:
Inhibitor
Concentration
(mM)
Pyridoxol-P
Pyridoxamine-P
(0.3 mM)
(0.3 mM)
Pyridoxal
1
6
7
Pyridoxol
1
<3
<3
Pyridoxamine
1
<3
<3
4-Pyridoxate
1
3
5
4-PyTidoxate-P
0.2
23
52
1
33
62
Deoxypyridoxol
1
0
0
Deoxypyridoxol-P
0.2
34
54
1
42
70
Pyridoxal oxime
0.001
17
31
0.01
57
67
ulation) and noted that only the phosphorylated derivatives are signifi-
cantly inhibitory. Whether inhibition of these oxidases by analogs plays a
role in the depressant or toxic effects produced is at present unknown, but
it must be admitted that for deoxypyridoxol and its phosphate none of
the inhibitions is probably potent enough to be important in vivo.
The effects of analogs on the tissue levels of the vitamin Bg group are
particularly important in certain arguments relative to the mechanisms by
which these analogs are toxic. Umbreit (1955 a) believes that deoxypyri-
doxol exerts actions other than the antagonism of vitamin Bg function.
The basis for this is principally that deoxypyridoxol accelerates the ap-
pearance of deficiency symptoms when animals are on a diet lacking pyri-
doxine and yet does not reduce the tissue levels of pyridoxal coenzymes.
He has also pointed out that in some cases there is also no fall in transamin-
ase or decarboxylase activity during the "acute" deficiency produced by
deoxypyridoxol. Nevertheless, it is admitted that the toxic effects of de-
oxypyridoxol can be readily counteracted by pyridoxine administration.
Actually there is very little published on tissue levels of vitamin Bg as af-
fected by deoxypyridoxol. Stoerk (1950) reported that dietary deficiency
lowers liver pyridoxine content but that deoxypyridoxol produces no fur-
ther lowering despite a more rapidly appearing deficiency syndrome. Similar
results were obtained by Beaton and McHenry (1953) in rats exhibiting
acrodynia following deoxypyridoxol feeding (see accompanying tabulation).
Umbreit also cites unpublished data supporting these results. Effects of
deoxypyridoxol on enzyme activity in vivo will be taken up in the following
ANALOGS OF PYRIDOXAL 567
section, but there is now sufficient evidence that the activities of certain
pyridoxal-P-dependent enzymes are reduced.
„ .J . -p. J 1 Liver vitamin Br
ryndoxine JJeoxypyridoxol
(/'g/g)
- - 6.3
+ - 11.0
- + 6.3
+ + 9.6
Interpretation of this apparent discrepancy between toxic reactions and
insignificant changes in liver vitamin Bg during deoxypyridoxol treatment
can be made along several lines. In the first place, it is generally believed
that much of the tissue pyridoxal is bound to nonenzyme protein, possibly
in part through the aldehyde group, so that analyses of total tissue levels
do not necessarily reflect changes in coenzyme concentration. The fact that
enzyme activity is often depressed without significant changes in total vi-
tamin Bg suggests that this can be an explanation. That bound to nonen-
zyme protein may not be displaced by deoxypyridoxol since this analog
contains no aldehyde group. In the second place, analyses have been made
only in the liver and it is unlikely that changes in liver pyridoxal function
are responsible for any of the common toxic symptoms of deficiency. It is
even possible that during treatment with deoxypyridoxol there is a transfer
of vitamin Bg substances from one tissue to another. It is conceivable that
deoxypyridoxol has actions other than interference with pyridoxal function,
but the fact that its toxicity can be reduced by pyridoxine administration
points to a close relationship between its effects and pyridoxal. I think
that more emphasis must be placed on the changes in enzyme activity
rather than on the tissue levels of vitamin Bg for the reasons given above.
The most complete investigation of the effects of deoxypyridoxol on the
concentrations of tissue Bg vitamers is that of Bain and Williams (1960),
who utilized chromatographic separation. The results are summarized in
Table 2-36. The extreme fall in brain pyridoxal-P they believe must in some
way be related to the convulsions. The rapidity with which the analog can
deplete the brain coenzyme is surprising. Is the pyridoxal-P replaced on
the apoenzymes by deoxypyridoxol-P ? It seems unlikely that interference
with transport or metabolism of pyridoxol could produce such marked ef-
fects so soon. Also the displaced pyridoxal-P must be metabolized or leave
the tissue. It is not known why pyridoxamine-P does not fall comparably.
Dietary vitamin Bg deficiency for 37-51 days does not cause such marked
losses of pyridoxal-P or total Bg vitamers from the brain as the single dose
of deoxypyridoxol. The severe drop in pyridoxal-P in brain following deoxy-
568
2. ANALOGS OF ENZYME REACTION COMPONENTS
O)
CO
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to
o
^
K!
w
a
o
pq
H
<
<i1
H
OS
H
^
H
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;z;
o
o
cq
Q
Ph
>> O
Ph 'TS
COOM OOi-O-^ OfM>0 C^iO
1— IGO-* OifC-* 5<lTt<C<l lOi— I
00^1 locoi r~>0| (Mfo
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„H
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00 t^ ■*
lO o o
fS
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d Q
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o
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r-(
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T3 t3
ANALOGS OF PYRIDOXAL 569
pyridoxol must be reflected in depression of the activity of enzymes requir-
ing this coenzyme, but not necessarily in all equally. The failure of pyridox-
amine-P to fall as much as pyridoxal-P might indicate that transaminases
are not depleted as readily as other pyridoxal-P enzymes,-^ but interpreta-
tion is made difficult by the fact that we do not know what fractions of
these substances are bound to apoenzymes and to nonenzyme protein. The
effects on the tumors are similar but of less magnitude, and the changes in
pyridoxamine-P are again variable, even increasing in ascites tumor. The
rise in pyridoxal-P and total vitamers in ascites fluid may reflect the loss
of these substances from the cells.
Effects of Deoxypyridoxol on Pyridoxal-P-Dependent Enzymes in Vivo
The results reported on transaminase activity during administration of
deoxypyridoxol are variable. Transamination in hamster hearts is reduced
30-40% in animals with a dietary deficiency, but injecting deoxypyridoxol
at 50 //g per animal 3 times a week does not lower the activity further
(Shwartzman and Hift, 1951). However, there is some growth inhibition
beyond that shown in the deficient animals, although no specific symptoms
were noted. Deoxypyridoxol at 100 //g/day in rats does not alter the aspar-
tate-glutamate transaminase and actually seems to increase the alanine-
glutamate transaminase activity in liver compared to animals on a deficient
diet (Caldwell and McHenry, 1953). Since the animals receiving deoxypyri-
doxol had severe dermatitis, it was justifiably concluded that the production
of dermatitis is unrelated to liver transaminase. On the other hand, Dietrich
and Shapiro (1953 a) found a greater fall in liver transaminase when mice
were injected with deoxypyridoxol at 150 mg/kg/day than in simple dietary
deficiency ( — 49% and —37%, respectively). Indeed, transaminase levels
in several tissues fall very markedly in mice on 175 mg/kg/day of deoxy-
pyridoxol (Shapiro et al., 1953). There is not much difference in the rates
of decrease in the various tissues (Fig. 2-20). It is difficult to reconcile all
of these results unless it is a matter of species variation, which is unlikely.
Transaminases are not directly inhibited very potently by deoxypyridoxol;
the K^ is 0.12 m.M for the alanine-pyruvate transaminase of Pseudomonas,
for example (Dempsey and Snell, 1963). The very high inhibitory activity
of pyridoxyl-L-alanine {K, = 0.00018 mM) is surprising.
The results on decarboxylases are similar. Dietary pyridoxine deficiency
causes a 50% fall in rat brain glutamate decarboxylase, but administering
deoxypyridoxol in addition produces no further depletion (Roberts et al.,
1951). More recent studies, summarized in Table 2-37, clearly indicate a
lack of correlation between the brain decarboxylase levels and the occur-
rence of convulsions; e.g., 3-deoxy pyridoxol convulses without a significant
change in enzyme activity, whereas c/j-methylpyridoxol lowers the enzyme
level without producing convulsions. Liver dopa decarboxylase is decreased
570
2. ANALOGS OF ENZYME REACTION COMPONENTS
moderately in mice from both deficiency and injections of deoxypyridoxol
(Dietrich and Shapiro, 1953 a). Despite the lowered levels of transaminase
and decarboxylase, cysteine desulfhydrase (also dependent on pyridoxal-P)
is not affected by deoxypyridoxol, due to the fact that the apodesulfhydrase
has little affinity for deoxypyridoxol-P, no inhibition of the binding of py-
ridoxal-P being observed (Dietrich and Borries, 1956).
600
500
400
300
200
^ 100
LIVER
KIDNEY
TIME (DAYS)
Fig. 2-20. Effects of deoxypyridoxol given intraperitoneally at
150 mg/kg/day on the aspartate: a-ketoglutarate transaminase of
mouse tissues. (From Shapiro et al., 1953.)
Serine biosynthesis from formate and glycine, involving serine transhy-
droxymethylase, is dependent on pyridoxal-P. The incorporation of for-
mate-C^* into serine in chick liver extracts is much reduced when these
are obtained from deoxypyridoxol-treated animals, the depression being
around 50% and reversible with pyridoxal-P in vitro (Sakami, 1955). Renal
ANALOGS OF PYRIDOXAL
571
Table 2-37
Effects of Pyridoxine Analogs on Rat Brain Gltjtamate Decarboxylase °
Glutamate decarboxylase
Treatment
Dose
(mg/kg)
Convulsions
(//moles/g/hr)
Endogenous
+Pyridoxal-P
Pjrridoxine-deficient
Control
36
218
Toxopyrimidine
15
4-
40 ( + 11%)
202 (- 7%)
Control
54
235
Toxopyrimidine
25
+
56 (+ 4%)
233 (- 1%)
Normal diet
Control
88
274
Toxopyrimidine
700
+
50 (-43%)
266 (- 3%)
3-Deoxypyridoxol
150
-
56 (-36%)
245 (-11%)
cy-Methylpyridoxol
200
-
31 (-65%)
261 (- 5%)
Pyridoxine-supplemented *
(100 //g%)
Control
70
276
3-DeoxypyridoxoI
100
+
72 (+ 4%)
252 (- 9%)
4-Deoxypyridoxol
100x7
-
71 (+ 2%)
296 (+ 7%)
5-Deoxypyridoxol
100x7
-
82 ( + 18%)
283 (+ 3%)
co-Methylpyridoxol
100x7
—
30 (-57%)
233 (-16%)
° From Rosen et al. (1960).
'' The analogs were injected intraperitoneally for 7 days, except for 3-deoxyp>Ti-
doxol, which convulsed the animals after a single dose.
glutaminase is also reduced by deoxypyridoxol at 21 days, at which time
dietary deficiency produces no change (Beaton and Goodwin, 1955).
It is impossible to evaluate the importance of these changes in enzyme
levels for the toxic effects of deoxypyridoxol. The toxic effects are mani-
fested principally in the central nervous system, skin, and hematopoietic
system, and no enzyme determinations during deoxypyridoxol treatment
have been reported in any of these tissues, except for brain glutamate de-
carboxylase. There is no reason to attribute the toxic effects to reduction
of transaminase rather than to the possible reduction of many other en-
zymes dependent on pyridoxal-P, most of which have never been examined
in this connection. It is more likely that the growth inhibition observed
572 2. ANALOGS OF ENZYME REACTION COMPONENTS
with both deficiency and deoxypyridoxol is related to lowered transami-
nase activity and protein synthesis. Analyses of total brain do not, of
course, necessarily reflect local changes in amino acid metabolism, and the
site of origin of the convulsions has yet to be determined. It must be re-
membered that transaminase, glutamate decarboxylase, and y-aminobu-
tyrate levels vary in different regions of the central nervous system and,
furthermore, the functional dependence of various regions on pyridoxal-
P-dependent metabolism must also vary. The displacement of pyridoxal-P
from apoenzymes by deoxypyridoxol-P in vivo depends on several factors:
(1) rate of penetration of the analog into the cells, (2) ability of the tissue
to phosphorylate the analog, and (3) relative affinities of the apoenzyme
for the coenzyme and the phosphorylated analog. Thus one might for sev-
eral reasons expect the pattern of enzyme depression from deoxypyridoxol
to be different from that produced by simple dietary deficiency.
Effects on Metabolism
Kynurenate is a normal metabolite of tryptophan in the rat, but in pyri-
doxine-deficient animals one finds kynurenine and xanthurenate also. The
metabolic pathway involved here may be summarized as follows:
kynurenate
y
Tryptophan -> formylkynurenine -> kynurenine -> 3-OH-kynurenine -> xanthurenate
\ "
anthranilate
This is an interesting situation since the formation of all three of these
products involves pyridoxal-P enzymes, namely, kynurenine transaminase
for the formation of kynurenate and xanthurenate, and kynureninase for
the formation of anthranilate. The administration of deoxypyridoxal to
otherwise normal rats produces no particular effect, but if tryptophan is
given to deoxypyridoxol-treated animals there is an increase in the appear-
ance of kynurenine and xanthurenate, just as in dietary deficiency (Porter
et al., 1947). The rise in kynurenine would be expected because two of its
degradative pathways are depressed (including the normally most impor-
tant one), and the increase in xanthurenate excretion must be due to a
diversion of the metabolic flow through the remaining pathway. However,
it is difficult to understand why xanthurenate excretion should increase
relative to kynurenate, since both are presumably formed with the same
enzyme, unless a rise in kynurenine concentration increases relatively more
the rate of the xanthurenate pathway. It would be interesting to know
what happens to the level of 3-hydroxykynurenine during deoxypyridoxol
administration.
Pyridoxine-deficient rats have higher blood urea than normal animals
and this has been attributed to an impaired utilization of amino acids,
ANALOGS OF PYRIDOXAL 573
since it is not of renal origin and hence due to an increased urea formation.
Administration of 100 //g/rat/day of deoxypyridoxol for 28 days apparently
increases urea formation in liver slices in both deficient and pyridoxine-fed
animals (see accompanying tabulation), although the authors stated that
Pyridoxine
Deoxypyridoxol
Vurea
4.32
+
—
3.38
—
+
4.76
+
+
4.00
deoxypyridoxol lessens the deficiency abnormalities rather than accentuat-
ing them (Beaton et al., 1954). The cycle of urea formation does not require
pyridoxal directly, but the aspartate to condense with citrulline must be
formed by transamination reactions, so that one might expect impaired
pyridoxal function to depress urea formation by this mechanism, and per-
haps it counteracts to some extent the efl"ect of an increased supply of amino
acids for catabolism.
Convulsive seizures in mice are produced by the injection of 4-methoxy-
methylpyridoxol, and are completely prevented by pyridoxol in a dose 3
times that of the analog (Gammon et al., 1960). The convulsions can also
be prevented by prior administration of y-aminobutyrate but not by any
of the other related amino acids or products of glutamate metabolism test-
ed. Since glutamate decarboxylase requires pyridoxal-P, the most obvious
explanation would be that the analog reduces brain y-aminobutyrate, there-
by initiating convulsions, and that the administration of y-aminobutyrate
simply restores the normal level or prevents depletion. Others have suggest-
ed that certain carbonyl reagents, such as thiosemicarbazide, produce sei-
zures by reducing brain y-aminobutyrate, and many now believe that ceU'
tral motor activity is controlled bj^ the levels of such amino acids and their
corresponding amines. Analyses of mouse brains obtained during convul-'
sions from 4-methoxymethylpyridoxol and other analeptics were thus made,
and it was found that y-aminobutyrate drops 50-70% when the analog is
used but shows no significant change when the convulsions are due to Me-
trazol, picrotoxin, or electroshock (Gammon et al., 1960; Kamrin and Kam-
rin, 1961). Despite the coherence of these observations in supporting the
role of y-aminobutyrate in antipyridoxine convulsions, one additional fact
is difficult to fit in: the administration of pyridoxol, which blocks the sei-
zures, does not alter the fall in brain y-aminobutyrate. It has also been
shown that administration of y-aminobutyrate to animals with seizures pro-
duced by the analog does not reduce the seizures although the y-amino-
574 2. ANALOGS OF ENZYME REACTION COMPONENTS
butyrate content of the brain increases (Purpura et al., 1960). These dis-
crepancies might be removed if one could determine local changes in y-ami-
nobutyrate in the central nervous system. On the other hand, it is possible
that other disturbances are more pertinent to the convulsive state, and, as
Rosen et al. (1960) have pointed out, perhaps more thought should be given
to the interference with the transport of amino acids into the brain cells.
Variations in the metabolism of amino acids other than glutamate and the
levels of physiologically active amines have not been studied. In this con-
nection it is interesting that Schrodt et al. (1960) administered deoxypyri-
doxol to two patients with malignant carcinoid syndrome at doses of 100-
200 mg/day, and found in one patient a fall in the urinary excretion of 5-
hydroxyindoleacetate, which is the primary product of serotonin metab-
olism, along with symptoms of vitamin Bg deficiency.
The role of pyridoxal in lipid metabolism is not yet clear but there is
some evidence in animals for a requirement in fatty acid synthesis. When
deoxypyridoxol was administered to seven subjects at 300 mg/day, six de-
veloped symptoms of pyridoxine deficiency (Mueller et al., 1959). There was
a general decrease in the polyunsaturated fatty acids of the blood, but no
change is phospholipids or cholesterol. This was believed to be evidence
for the involvement of pyridoxine in maintaining blood fatty acids through
participation in the synthetic reactions, but the data do not indicate a role
in the interconversion of the unsaturated fatty acids.
Effects on Active Transport
Although relatively Httle has been done with respect to the actions of
pyridoxine analogs, there is accumulating evidence that amino acid trans-
port is often related to pyridoxal-P function, and it is likely that some of
the toxic effects of the analogs will be explained on this basis. Part of the
transport of amino acids across the intestinal wall is active and is inhibited
by deoxypyridoxol (Fridhandler and Quastel, 1955). A 41% inhibition of
L-alanine transport was observed with 10 mM deoxypyridoxol, which is
certainly a very high concentration; however, pyridoxol at the same con-
centration has no effect. The inhibition is not antagonized by pyridoxol,
pyridoxal, or pyridoxal-P, which may indicate that phosphorylation is not
rapid enough intracellularly and that the coenzyme itself cannot penetrate,
or that the inhibition is not an antagonism of pyridoxine. A rather disturb-
ing fact is that glucose and fructose transport is also inhibited by deoxy-
pyridoxol to about the same extent as alanine absorption, so that this is
not a specific effect on amino acid transport.
The transport of d- and L-methionine across rat intestine is depressed
by deoxypyridoxol injected at 200-400 //g/day (Jacobs, 1958; Jacobs and
Hillman, 1958). The effect appears within an hour after intraperitoneal
injection and can be abolished by injection of pyridoxol (see accompanying
ANALOGS OF PYRIDOXAL 575
tabulation) (Jacobs et al., 1960). These results definitely implicate pyridoxal
in amino acid transport but do not prove that it functions directly in the
transport mechanism, since the effect could be an indirect one.
Pyridoxol Deoxypyridoxol % Change in transport
- - —56
+ — +25
+ -35
+ + +6
The uptake of glycine by ascites carcinoma cells is inhibited by 5-25 mM
deoxypyridoxol (Christensen et al., 1954). Since pyridoxine deficiency re-
duces the accumulating ability and this is restored by pyridoxal in vitro,
it would appear that pyridoxal functions here in some manner, although
again not necessarily in the membrane transport system.
Effects on Growth
Deoxypyridoxol suppresses the growth of a variety of microorganisms.
Rabinowitz and Snell (1953 a) emphasized that sensitive organisms are
those requiring an exogenous source of pyridoxine, and in these the inhibi-
tion can be counteracted by pyridoxine; those synthesizing their own p\Ti-
doxal can effectively resist the analog. The situation is quite complex, how-
ever, and when different analogs are examined a marked variability in sus-
ceptibility is found (Rabinowitz and Snell, 1953 b). For example, w-me-
thylpyridoxol is inhibitory to yeast but not at all to Streptococcus faecalis or
Lactobacillus helveticus, the 5-deoxypyridoxol derivatives being the most ef-
fective in these latter organisms; in L. helveticus, only 5-deoxypyridoxol is
inhibitory, 4-deoxypyridoxol, 5-deoxypyridoxol, and 5-deoxypyridoxamine
being without action. A factor that is very important in determining the
susceptibility of bacteria to these analogs is the nature of the exogenous
amino acids supplied. Streptococcus faecalis grows well if all amino acids are
provided even though pyridoxine is absent, but a requirement for pyrido-
xine and a sensitivity to analogs are created by restriction of the amino
acids in the medium (Olivard and Snell, 1955). Under certain circumstances
the growth can be limited by conversion of l- to D-alanine by alanine race-
mase, which involves pyridoxal-P and is quite sensitive to 5-deoxypyridoxol
{K, = 0.089 mM) and w-methylpyridoxol {K^ = 0.53 mM). The inhibition
of growth by these analogs can be explained on the basis of the inhibition
of this enzyme under these conditions. On the other hand, in most circum-
stances the inhibition must be on amino acid metabolism, as in Vibrio
576 2. ANALOGS OF ENZYME REACTION COMPONENTS
cholera where alanine and aspartate accumulate when growth is suppressed
55% by 0.53 vciM deoxypyridoxol (Chatterjee and Haider, 1960). Perhaps
the first pyridoxine analogs to be recognized as inhibitors of bacterial growth
were the tellurium compounds studied by Morgan, Cooper, and their col-
leagues between 1923 and 1926. Gulland and Farrar (1944) postulated that
2,4-dimethyl-c^c^o-telluropentane-3,5-dione is toxic because of its structural
similarity to pyridoxine, but no further work or attempts to counteract the
inhibition with pyridoxine have come to my attention. An in vivo effect on
bacteria has been demonstrated in at least one case for deoxypyridoxol,
which prolonged the survival time of mice infected with Toxoplasma gondii
from 5 days to 12.1 days when it was incorported into the diet at 0.1%,
although some toxic symptoms were noted (Summers, 1957). Pyridoxine,
can counteract both the beneficial and toxic effects.
Chick embryogenesis is disturbed by deoxypyridoxol and other analogs.
When 1 mg of deoxypyridoxol is injected into eggs, there is 100% mortality
of the embryos and this can be prevented by injecting pyridoxine (Cravens
and Snell, 1949). However, after 4 or more days the embryos become less
sensitive, and although higher doses are toxic they are not counteracted
by pyridoxine. Similar effects are noted for 4-methoxymethylpyridoxol but
it is at least 25 times more toxic than deoxypyridoxol to the early chick
embryo (Karnofsky et al., 1950). Mammalian fetal development is also dis-
turbed by deoxypyridoxol — fetuses resorbed, still-births, and abnormal
young — but administration of estrone and progesterone together prevents
these effects and pregnancy is maintained in the majority of the animals,
indicating that the action of the analog is primarily on the maternal tissues
rather than the embryo (Nelson, 1955).
Some of the observations relative to the inhibition of tumor growth by
deoxypyridoxol will be summarized since these effects are interesting in
light of the fairly rapid amino acid metabolism in tumors and the generally
low levels of the Bg vitamers in solid tumors. Regression of mouse lympho-
sarcoma implants with deoxypyridoxol was achieved by Stoerk (1947, 1950)
when the animals were on a low-pyridoxine diet, but there was also loss
of body weight, suggesting an insufficiently specific inhibition. The fre-
quency of successful fibrosarcoma implants in rats is increased by pyri-
doxol and decreased by deoxypyridoxol, even though in the latter case no
severe deficiency symptoms are observed (Loefer, 1951). On the basis of
these findings, Gellhorn and Jones (1949) gave deoxypyridoxol to patients
with disseminated lymphosarcoma and acute leukemia in combination with
a pyridoxine-deficient diet. Although there was some weight loss and weak-
ness, there were no specific signs of deficiency, no changes in tryptophan
metabolism, no hematopoietic depression, and no retardation in the growth
of lymphoid tissue, the results being clinically insignificant. Deoxypyridoxol
appears to be reasonably effective in suppressing mammary carcinoma in
ANALOGS OF PYEIDOXAL 577
mice, with a specific regression of the tumor and no weight loss (Shapiro
and Gellhorn, 1951). Human carcinoma cells (Eagle's KB strain) are quite
sensitive to deoxypyridoxol in tissue culture, 0.08 mM inhibiting the growth
50% (Smith etal., 1959). It appears doubtful that deoxypyridoxol is suffi-
ciently specific as a carcinostatic agent but its use in conjunction with other
inhibitors remains a possibility, especially as Doctor (1959) has shown that
deoxypyridoxol at 20 mg/kg/day in the rat has no effect on the leucocyte
count, but combined with a moderately effective dose of aminopterin exerts
a very marked suppression of the leucocytes, and also potentiates the action
of oxythiamine.
Toxic Effects in Whole Animals
Some of the evidence that deoxypyridoxol can produce rather typical
vitamin Bg-deficiency states will be summarized to emphasize that, what-
ever the basic biochemical disturbances, the effects are primarily related to
an interference with formation or function of pyridoxal-P. It is first of all
quite clear that the doses of deoxypyridoxol to induce toxic reactions must
be much higher when the animals are adequately supplied with the Bg vi-
tamers than when the animals are subjected to a dietary deficiency, and
that administration of pjTidoxol can overcome the toxic reactions produced
by the analog. In general the responses to deoxypyridoxol are the same
as in pyridoxine deficiency, except that they appear earlier, producing an
acute deficiency syndrome. Thus in rats and mice there is a dermatitis
characterized by acanthosis, parakeratosis, and hyperkeratosis, sometimes
with a superimposed infection, which is similar to deficiency dermatitis
(Stoerk, 1950). There is atrophy and degeneration of the hematopoietic
organs, evidenced by decreases in thymus and spleen weights, and these are
reflected in the peripheral blood picture (Mushett et al., 1947). The nervous
system hyperirritability leading eventually to convulsions has been men-
tioned in connection with the metabolic changes in the brain.
A state resembling pyridoxine deficiency has been produced in man by
Mueller and Vilter (1950). Eight individuals on a pyridoxine-poor diet were
injected intramuscularly with 60-150 mg/day. Within 2-3 weeks a sebor-
rheic dermatitis appeared around the eyes, nose, and mouth, with simulta-
neous glossitis and stomatitis. These symptoms disappeared in 2-3 days
upon administration of pyridoxol. The total white count did not fall, nor
was there evidence of anemia, but the lymphocytes dropped to around half
the initial level. Schrodt et al. (1960) in their two carcinoid patients also
observed seborrheic dermatitis and glossitis. The general pharmacology of
analogs and inhibitors of pyridoxal function has been reviewed by Holtz
and Palm (1964).
Although several workers have stated that reactions may be seen in acute
deoxypyridoxol-treated animals which are not seen in simple dietary de-
578 2. ANALOGS OF ENZYME REACTION COMPONENTS
ficiency, there appears to be no really good evidence for any of these reac-
tions being unassociated with pyridoxal function. We have discussed (p. 567)
various possible reasons for different effects of deoxypyridoxol and dietary
deficiency on tissue Bg vitamer levels and, hence, on metabolic disturbances
in the tissues.
Toxopyrimidine
This substance (4-amino-5-hydroxymethyl-2-methylpyrimidine), which is
essentially the pyrimidine portion of thiamine, has been known for many
years to produce abnormal motor behavior and convulsions, and a search
for antidotes led to the discovery that the pyridoxine group is specific in
preventing the toxic reactions. It was then realized that toxopyrimidine
bears a structural resemblance to pyridoxal. Makino and Koike (1954 a,b)
believed that toxopyrimidine acts in the phosphorylated form since tyrosine
decarboxylase is not inhibited by toxopyrimidine up to 1 mM, whereas to-
xopyrimidine-P exerted some inhibition at 0.001 mM and almost complete
inhibition at 0.1 mM. The inhibition is competitive with respect to pyridox-
al-P. Haughton and King (1958) confirmed this inhibition but stated that
it required an (I)/(C) ratio of 1000 to get 50% inhibition, whereas Makino
and Koike found around 50% inhibition with a ratio near 3. No inhibition
of tryptophanase, transaminase, glutamate decarboxylase, or arginine de-
carboxylase was observed, and they concluded that toxopyrimidine is not
of much value in the study of pyridoxal-P enzymes. The failure to inhibit
significantly the tryptophanase of E. coli was also reported by Wada et
al. (1958). McCormick and Snell (1961) found no inhibition of pyridoxal
kinase at concentrations of 0.01-0.1 mM toxopyrimidine. Rindi and Fer-
rari (1959) found that 90-120 min after the intraperitoneal injection of
125 mg/kg of toxopyrimidine in pyridoxine-deficient rats, convulsions hav-
ing been produced, the y-aminobutyrate levels in the brain have fallen
some 23%, although glutamate is unchanged. Administration of pyridoxa-
mine stops the convulsions and increases brain y-aminobutyrate. This dose
of toxopyrimidine reduces brain glutamate decarboxylase 20% but does
not significantly alter transaminase activity (Rindi et al., 1959). Again pyri-
doxamine restores activity. We have already noted (Table 2-37) that toxo-
pyrimidine can reduce brain glutamate decarboxylase at high doses, but at
lower convulsive doses in pyridoxine-deficient animals it does not. It would
seem that if toxopyrimidine causes convulsions by interfering with pyri-
doxal function, it is not mediated through a general fall in y-aminobutyrate
or transaminase activity, and the status of the mechanism is much the
same as for deoxypyridoxol, namely, uncertain.
ANALOGS OF PTEROYLGLUTAMATE (FOLATE)
579
ANALOGS OF PTEROYLGLUTAMATE (FOLATE)
Tetrahydrofolate functions metabolically in the transfer of C^ units at
the oxidation level of formaldehyde or formate, and thus is important in
the biosynthesis of purines, pyrimidines (thymine), certain amino acids (ser-
ine, histidine, methionine), choline, and other biologically important sub-
stances. Interference with its function leads secondarily to a depression of
nucleic acid and protein synthesis and, because of this, to a general sup-
pression of cellular growth and multiplication. The possible sites of block
for folate analogs may be summarized as: (1) pathway for the synthesis
of folate, (2) reduction of folate to tetrahydrofolate, (3) reactions of C^ unit
transfer, and (4) degradative reactions of folate and its derivatives. The
transport of folate into cells should probably also be considered as a pos-
sible site of inhibition but little about this process is known at the present
Folate (F)
(1)
dihydrofolate (FH,)
(3)
(2)
S-formyl-FH^
(5)
(8) 5-formimino-FHj
(9) (10)
5, 10-methenyl-FH,
(1) folate reductase
(2) dihydrofolate reductase
(3) glutamate-FH4 transformylase
(4) serine hydroxymethylase
(5) formimino transferase
(6) 10-formyl-FH4 deacylase
— tetrahydrofolate (FHJ —
(4)
-•^ lO-hydroxymethyl- FH^
(6y\(7)
10-formyl-FH4
(11)
5, 10- methylene- FH4
(7) tetrahydrofolate formylase
(8) (cyclohydrolase)
(9) formimino- FH4 cyclodeaminase
(10) cyclohydrolase
(11) (cyclohydrolase)
(12) 5, 10-methylene-FH4 dehydrogenase
time. A number of substances can inhibit the synthesis of folate in micro-
organisms — e.g., the sulfonamides and certain pteridines — but they will
not be treated in this chapter. There is no evidence that the important
actions of folate analogs are related to inhibition of degradative reactions.
We shall therefore limit the subject in this section to inhibitions of folate
reduction and transformylation reactions, namely, those pathways shown
in the accompanying diagram.
580
2. ANALOGS OF ENZYME REACTION COMPONENTS
H,N
-NH
v\ //
COO
I
CH,
I "
CHo
I ""
CH— COO"
I
CONH
Folate
H,N
OH CHO
5-Formyltetrahydrofolate
(folinate, citrovorum factor)
H,N^ /N. /N^ ^CH,
OH
7-Methylfolate
H,Nv. M^ M
H,N
NH,
Amethopterin
(methotrexate)
Pyrimethamine
(Daraprim)
2, 4, 7 Triamino-6-
o-methylphenylpteridine
ANALOGS OF PTEROYLGLUTAMATE (fOLATE) 581
The most thoroughly studied folate analogs are aminopterin (4-hydroxy
group replaced by amino group) and amethopterin (10-methylaminopterin)
because of their importance in the chemotherapy of cancer. These substances
produce states of folate deficiency in all types of organism and, in most
instances, the symptoms of this deficiency can be prevented by providing
tetrahydrofolate or 5-formyltetrahydrofolate, but not with folate, indicating
that the site of block lies somewhere in the pathway of the transformation
of folate to its coenzymically active forms. Although this block produces a
general disturbance in tetrahydrofolate function and, it appears, nucleic
acid and protein synthesis, there are two reasons for some degree of speci-
ficity. In the first place, the syntheses of the various substances requiring
Ci units from folate coenzymes are not necessarily all depressed equally,
since it is a general rule that a lowering of the concentration of some sub-
stance from which several pathways lead will produce varying effects on
these pathways, depending mainly on the nature of the enzymes involved
and the supply of reactants for each pathway (in this case C^ unit accep-
tors). In the second place, those cells or tissues with the highest rates of
synthesis and dependent on these synthetic reactions for growth or multi-
plication wiU be most adversely affected by the folate analogs. Here it is
not a matter of degree of functional activity but of proliferation; the heart
is not readily affected by these analogs whereas the hematopoietic system is.
We shall not discuss the more biological aspects of the actions of these
analogs, since this is covered adequately in a number of books and reviews
(e.g. Holland, 1961; Delmonte and Jukes, 1962), but confine attention to
the basic enzyme and metabolic effects.
Inhibition of the Reduction of Folate to Tetrahydrofolate
This reduction occurs in two steps and in most instances it appears that
each step is catalyzed by a specific enzyme, but possibly in other cases a
single enzyme is responsible. Most assays of folate reduction for analog
inhibition have involved determination of the formation of either tetrahy-
drofolate or folinate, or the disappearance of folate, and it is difficult to
differentiate between the two steps with regard to inhibition. In some re-
ports the term "folate reductase" is applied to the over-all reaction. One
thing is certain: Dihydrofolate reductase, which has been better purified
and more thorougly studied than folate reductase, is very potently inhibited
by aminopterin and amethopterin. The enzyme from chicken liver is inhib-
ited 74% by 0.000053 mM aminopterin (Futterman, 1957) and from hu-
man leukemic leucocytes 64% by 0.00001 mM (Bertino et al, 1960), in
both cases the substrate being 10,000-fold or more in excess of the analog.
Osborn et al. (1958) calculated the K's for aminopterin and amethopterin
to be 0.000001 mM and 0.0000023 mM, respectively, using the chicken liver
enzyme, Blakley and McDougall (1961) reported a value of 0.0000024 mM
582 2. ANALOGS OF ENZYME REACTION COMPONENTS
for aminopterin and the enzyme from Streptococcus faecalis, and Nath and
Greenberg (1962) gave K, as 0.0000023 mM for amethopterin and the calf
thymus enzyme. The inhibitions of the full reduction of folate to tetrahy-
drofolate by aminopterin and amethopterin are very similar (Futterman,
1957; Futterman and Silverman, 1957; Zakrzewski and Nichol, 1958; Silber
et al., 1962), and point to the dihydrofolate reductase as being the more
sensitive enzyme. Furthermore, the formation of folinate (citrovorum fac-
tor) from folate is very potently inhibited in rat liver slices (Nichol and
Welch, 1950), Lactobacillus casei and Streptococcus faecalis (Hendlin et al.,
1953), mouse leukemic cells (Nichol, 1954), and chicken liver extracts
(Doctor, 1958). The most potent pteridine analog was found by Doctor
(1958) to be 2,4,7-triamino-6-o-methylphenylpteridine, although it is not
as potent as aminopterin, and he showed that the site of inhibition is pre-
vious to tetrahydrofolate. There is thus much evidence that folate reduc-
tion is blocked by low concentrations of these analogs, and it is generally
agreed that this must be the primary mechanism by which folate defi-
ciency and growth depression are produced.
The inhibitions of folate reduction by aminopterin and amethopterin have
been reported to be noncompetitive by several investigators, but it is very
likely that the inhibitions are truly competitive, this being obscured by the
much greater affinity of the enzyme for the analog than for dihydrofolate.
In no case has the rate of inhibition in the presence of varying concentra-
tions of substrate been determined, but one might predict that the competi-
tive nature of the inhibition would be demonstrated in this way. Once the
enzyme is inhibited, it is very difficult to recover the activity because the
rate of dissociation of the analog from the enzyme is extremely slow. In
other words, this is an example of pseudoirreversible inhibition obeying mu-
tual depletion kinetics. This was shown by Peters and Greenberg (1959) on a
sheep liver folate reductase, the inhibition at constant analog concentration
being dependent on the enzyme concentration. It is thus possible to titrate
this enzyme in tissues or extracts. This has been well discussed by Werk-
heiser (1961), who also showed that the amount of analog bound by rat
liver supernates is equivalent to the amount required to inhibit folate re-
duction; in other words, the tightly bound analog seems to be combined
only with dihydrofolate reductase. If rats are injected with amethopterin,
the supernatant fraction of the liver contains most of the analog and only
10-15% of this is lost during dialysis for 6 days (Werkheiser, 1959). The
amount of amethopterin or aminopterin to inhibit completely the reductase
in liver extracts in 0.56 //g/g of liver, and the supernates of livers from
analog-treated rats contain 0.52 //g/g of tissue. The reductase is the only
protein binding these analogs significantly in chicken liver homogenates
(Schrecker and Huennekens, 1964). Fountain et al. (1953) had found that
there is a remarkable retention of amethopterin in the tissues of mice, the
ANALOGS OF PTEROYLGLUTAMATE (fOLATE) 583
concentration in the liver remaining approximately constant for at least 3
weeks after a single intravenous dose. It is noteworthy that some tissues,
such as the lung and spleen, do not pick up much of the analog, and that
the kidney loses the analog relatively more rapidly than the liver. Werk-
heiser (1960) likewise found that the liver retains aminopterin for long pe-
riods, while the intestine does not. In mice given a lethal dose of aminop-
terin 1 hr after a protective injection of folate, the liver folate reductase
activity is depressed 96% after 24 hr and remains at this level for 7 days,
following which there is a slow recovery (Werkheiser, 1962). The intestinal
enzyme is similarly inhibited but recovers faster. The loss of aminopterin
from the intestine is characterized by a half-life of 60 hr, but the liver shows
two components with half-lives of 60 hr and 90 days, respectively. It was
suggested that rapidly proliferating cells are dependent on folate reductase
activity and that the disappearance of the inhibition is faster in such tissue
because of the more rapid turnover of cells; in other words, the 60 hr com-
ponent would arise from proliferating tissue while the 90 day component
would relate to nonproliferating tissue. If this is the case, the binding of
aminopterin to the enzyme in vivo must be essentially irreversible. These
results all point to a very high degree of specificity in the binding and inhi-
bition, and confirm the major site of attack as being on folate reduction.
Further evidence comes from the reduced urinary folinate levels in rats on
25 //g/day of aminopterin (Nichol and Welch, 1950). Less direct evidence
is provided by Nichol (1954) and Broquist et al. (1953), who showed that
resistant streptococci or leukemic cells have a much greater ability to pro-
duce folinate from folate than do normal cells. It is possible that this in-
creased activity allows enough active folinate to be formed to enable the
cells to grow and multiply in the presence of the analogs, but it is probable
that this is not the only mechanism of resistance to these agents.
The nature of the binding of these folate analogs to the reductase has
not been fully elucidated, but Zakrzewski (1963) has determined the ther-
modynamic characteristics for the dissociation of the EI complexes (see ac-
companying tabulation). The inhibitions by the substituted pteridines are
competitive. The K^ values for aminopterin were taken from Werkheiser
Changes for
EI dissociation
Inhibitor
Ki (mM) AH
AFo
AS
2,6-Diaminopurine
0.0018 +6.0
+ 7.9
-6.4
2,4-Diamino-6-methylpteridine
0.0018 +5.0
+ 7.9
-9.5
2,4-Diamino-6-formylpteridine
0.0081 +4.1
+6.9
-9.5
2,4-Diamino-6-hydroxypteridine
0.37 +1.8
+4.7
-9.6
Aminopterin
10 '-10-8 +11.6
+ 13.7-15.1 -
-7.0-11.7
584 2. ANALOGS OF ENZYME REACTION COMPONENTS
(1961) and are uncertain because it is not known whether competitive or
noncompetitive inhibition occurs; it may be noted that the values are one-
hundredth to one-tenth those given by previous workers. The pyrimidine
amino groups are essential for tight binding and the pyrazine ring presum-
ably does not participate in the binding. A binding mechanism was proposed
in which emphasis is placed on the tautomeric state of the analogs and the
formation of hydrogen bonds between the ring nitrogens and the amino
groups with the enzyme, the replacement of the 4-OH group of folate with
an amino group favoring greater hydrogen bonding. It may also be ob-
served that the benzoylglutamate portion of the molecule must contribute
around 6-7 kcal/mole binding energy.
Another analog with less obvious structural similarity to folate is pyri-
methamine (Daraprim), an antimalarial drug that in chronic dosage pro-
duces folate deficiency in bacteria and animals (Wood and Hitchings, 1959 a;
Hitchings, 1960). Pyrimethamine, like the analogs previously discussed,
inhibits the reduction of folate, and does this at a concentration equivalent
to that required for growth inhibition. A 35% inhibition of folinate forma-
tion is caused in extracts of S. faecalis by 0.000012 mM pyrimethamine,
so that its potency is comparable to that of aminopterin. There are no ef-
fects on the biosynthesis or assimilation of folate. It is believed that the
antimalarial action is due to the tighter binding of the drug to the plasmo-
dial folate reductase than to the host enzyme. However, the uptake of
pyrimethamine by bacterial cells is unique inasmuch as it is inhibited
strongly by glucose, whereas the uptake of aminopterin is augmented by
glucose (see accompanying tabulation) (Wood and Hitchings, 1959 b). Fur-
Analog taken
up (cpm)
Analog
No glucose
Glucose
Pyrimethamine
Aminopterin
51.6
55
0.7
523
thermore, aminopterin uptake is increased by a rise in temperature, whereas
less pyrimethamine appears in the cells at higher temperatures. Despite
the apparent similarity of site of action of these two analogs, there is some
basic difference in the movement or disposition of the materials in the cells.
Effects on Synthetic Processes Mediated by Tetrahydrofolate
A block in the reduction of folate would be expected to depress the C^
unit transfers and the synthesis of nucleic acids and proteins as long as
there is no supply of tetrahydrofolate or folinate. The analogs in addition
ANALOGS OF PTEROYLGLUTAMATE (FOLATE) 585
might inhibit directly the reactions in which tetrahydrofolate functions.
There is essentially no information on this second possibility. Cyclohydro-
lase is inhibited weakly by amethopterin (43% at 0.5 mM) and aminopterin
(69% at 0.5 mM) (Tabor and Wyngarden, 1959), and formyltetrahydro-
folate synthetase is even more weakly inhibited by several analogs ( Jaenicke
and Erode, 1961; Whiteley et al., 1959). The many other reactions involved
have never been examined for inhibition by analogs. All one can say at the
present time is that the known inhibitions on folate reduction are sufficient
to explain most or all of the effects of these analogs.
Some examples of the inhibition of syntheses will be mentioned to illus-
trate the nature of the metabolic actions of these analogs. Aminopterin and
amethopterin invariably depress the incorporation of formate-C^* into pu-
rines and nucleic acids; this has been shown in rabbit bone marrow (Totter
and Best, 1955), leukemic spleen extracts (Balis and Dancis, 1955), and the
whole animal (Skipper et al., 1950). It appears that thymine synthesis is
more sensitive than purine synthesis to inhibition by these analogs. In bone
marrow 0.0021 mM aminopterin inhibits incorporation into thymine 72%
but into adenine or guanine only 22%. It would be interesting to know
what the effect on adenine nucleotides is, but little is known. Aminopterin
elevates ATP in liver, has no effect on spleen or muscle ATP, and reduces
tumor ATP (Zahl and Albaum, 1955). Indeed, the total adenine nucleotides
in the sarcoma decrease. One might expect the effects to depend on the
relative rates of adenine synthesis and turnover in the tissues. Aminopterin
lowers liver NAD levels — 39% fall at 60 //g/day and 56% fall at 100 //g/day
— and if this occurs throughout the body it could be an important conse-
quence of interference with folate metabolism (Strength et al., 1954). The
in vitro depression of respiration by aminopterin is not completely reversed
by added NAD, and it is very possible that other coenzymes (e.g. NADP,
coenzyme A. or FAD) are decreased.
The interconversion of glycine and serine is inhibited by aminopterin
when only folate is supplied, but the activity is restored with folinate (Blak-
ley, 1954). Tetrahydroaminopterin does not inhibit serine synthesis when
folinate is provided, but 2-deaminofolate inhibits some 55% at 0.75 mM in
rabbit liver extracts (Blakley, 1957). The incorporation of formate-C^^ into
lymphoma proteins is inhibited 72% by 0.0073 mM amethopterin, but little
effect is observed in normal liver (Williams et al., 1955). In general these
analogs block nucleic acid synthesis more than protein synthesis, but this
may vary a good deal from one tissue to another, or one organism to an-
other. Most of the inhibitions in nucleic acid and protein synthesis have been
attributed solely to defects in the formation of the constituent units, and
little consideration has been given to other possible contributing factors,
such as lowered levels of various coenzjTnes with impairment of oxidative
and phosphorylative reactions.
586 2. ANALOGS OF ENZYME REACTION COMPONENTS
Effects on Certain Enzymes Unrelated to Q Transfer
Brief mention should be made of the early demonstrations that 7-methyI-
folate is inhibitory to dopa decarboxylase and that this is reversible with
folate at 10-100 times the analog concentration (Martin and Beiler, 1947).
Pteroylaspartate and 7-methylpteroate are less potent inhibitors, but the
difference is not great, so that the glutamate portion is not very important
for the binding (Martin and Beiler, 1948). Tyrosine decarboxylase is not
inhibited by 0.67 mM 7-methylfolate whereas 0.067 mM inhibits dopa de-
carboxylase 25%. The mechanism of this inhibition, the role of folate in
decarboxylase activity, and the effects of the more commonly used folate
analogs are all unknown. The decarboxylase inhibition prompted a study
of the effects of 7-methylfolate on blood pressure (Martin et al., 1947), and
it was found that 5 mg/kg in the dog depresses the blood pressure quite
significantly and for an extended period of time, but whether this is related
to decarboxylase inhibition is problematical.
The inhibition of acetyl transfer by amethopterin (K, = 0.032 mM) in
pigeon liver extracts is interesting because it represents another possible
site of action for this analog, even though it is obviously much less potent
here than on folate reduction (Jacobson, 1960). The inhibition is not coun-
teracted by folate, tetrahydrofolate, or folinate, and is competitive with
the acetyl donor (p-nitroacetanilide) but noncompetitive with the acetyl
acceptor (aniline or sulfanilamide). These results do not implicate a folate
compound in the acetylation reaction — indeed, folate and folinate are
weak inhibitors, and the mechanism is more likely a simple binding to the
substrate site. The 10-methyl group is important since aminopterin is only
one-tenth, or less, as inhibitory as amethopterin. Evidence for inhibition
of acetylation in vivo is the higher sulfanilamide level and the lower acetyl-
sulfanilamide level in rabbit plasma of animals treated with amethopterin
(Johnson et al., 1958).
ANALOGS OF OTHER VITAMINS, COENZYMES,
AND THEIR COMPONENTS
There has been a great deal of work on the growth inhibitions produced
by numerous analogs of pantothenate, biotin, cobalamin, and other meta-
bolically necessary cofactors, but relatively few reports on enzyme inhibi-
tions are available. However, some of this isolated work on enzyme systems
is interesting in itself and perhaps some reference to it will stimulate further
study.
Pantothenate is required by many microorganisms and animals because
it is a component of coenzyme A, the biosynthetic pathway being:
Pantothenate -> pantothenylcysteine ->• pantetheine ->• CoA
ANALOGS OF OTHER VITAMINS 587
Mcllwain (1945) found that pantothenate analogs, such as pantoyltaurine,
do not displace bound pantothenate from bacterial cells, and concluded
that these analogs are bacteriostatic because they block the formation of
the metabolically active form of pantothenate (which was not known at
that time). Furthermore, as Martin et al. (1950) showed, the analogs in gen-
eral do not interfere in the reactions involving CoA. Pantoyltaurine and
other analogs do not inhibit brain choline acetylase, even at concentrations
around 5 vaM. One analog, salicyloyl-/?-alanide, does inhibit this enzyme
(20% at 0.47 mM and 100% at 4.7 mM), but since neither pantothenate
nor CoA reverses this inhibition it is doubtful if it is specific. Most effective
analogs thus seem to block the pathway of pantothenate -^ CoA, and no
known direct antagonists of CoA are known. Pantoylaminoethanethiol in-
hibits the synthesis of CoA from pantetheine (50% at a ratio of analog to
pantetheine of 13) and thus inhibits sulfonamide acetylation in liver ex-
tracts provided with pantetheine (Boxer et al., 1955). One cannot help but
wonder in some of these instances if abnormal CoA analogs are formed,
CH,
I
HOCH,— C— CHOH— CONH— CH,CH,— COO'
"■ I
CH3
Pantothenate
CH3
I
HOCH2— C— CHOH— CONH - CHoCH,— SO:
I - 2 3
CH,
Pantoyltaurine
CH3
I
HOCH2— C-CHOH-CONH-CH2CHi-CONH-CH2CH2SH
CH3
Pantetheine
CH,
I
OH
HOCH2—C — CHOH— CONH- CH2CH2SH / A— CONH-CH2CH2— COO"
CH3 \ /
Pantoylaminoethanethiol Salicyloyl-/3-alanide
588
2. ANALOGS OF ENZYME REACTION COMPONENTS
rather than a simple inhibition of the biosynthetic pathway. It is possible
to go back farther and inhibit the synthesis of pantothenate by analogs of
pantoate or /5-alanine. For example, 2,3-dichloroisobutyrate blocks the cou-
pling of these two components of pantothenate, competitive with pantoate
and uncompetitive with /5-alanine (K^ = 1.4-6.4 mM) (Hilton, 1958).
Biotin functions in several metabolic pathways (synthesis of aspartate
and higher fatty acids, and COg fixation) and in certain organisms can be
formed from desthiobiotin. 2-Oxo-4-imidazolidinecaproate (desmethyldes-
thiobiotin) inhibits the growth of E. coli by competing with desthiobiotin
for an enzyme involved in biotin synthesis, and this inhibition is competi-
tive (Rogers and Shive, 1947). Biotin by an unknown mechanism stimulates
fermentation in biotin-deficient yeast and this is inhibited by homooxy-
biotin, oxybiotinsulfonate (COO- group replaced by SOg" group in oxybio-
tin), and 7-(3,4-ureylenecyclohexyl)butyrate when the analogs are added
n O
HN"
■NH
HN'
■^NH
Biotin
■(CH2)^COO"
H3C CH.— (CH2)4— COO
Desthiobiotin
O
II
c
CH.r-(CH2) — COO'
-(CH2)5— COO'
Desmethyldes-
thiobiotin
Homobiotin
HN"^ NH
HN^ NH
(CH,) — COO'
/ V(CH2)3— COO'
Oxybiotin
y-(3, 4-Ureylenecyclo-
hexyl)butyrate
ANALOGS OF OTHER VITAMINS 589
before the biotin, but not afterward (Axelrod et al., 1948). This could mean
that these analogs cannot displace biotin once it is bound or that they in-
hibit in some way the formation of an active form of biotin. Biotin is inac-
tivated by kidney slices, possibly by removal of fragments from the side
chain, by biotin oxidase, and this enzyme is inhibited by several analogs
competitively (see accompanying tabulation) (Baxter and Quastel, 1953).
There is thus the possibility that some analogs can conserve biotin in the
tissues as well as inhibit its svnthesis or function.
4 1 Concentration , . , , , , . .
Analog (Analog)/ (biotin) % Inhibition
{raM )
DL-Homobiotin
1.65
10
96
DL-Desthiobiotin
1.63
10
96
L-Biotin
0.4
10
82
DL-Biotindiaminecarboxylate
0.4
10
82
D-Biotinoldiamine
0.4
10
45
D-Biotinol
0.41
10
24
2.1
50
61
D-Biotinsulfone
0.41
10
16
Analogs of cyanocobalamin (vitamin B^g) have not been extensively stud-
ied because of the complexity of the structure. 5,6-Dimethylbenzimidazole
is a component of cyanocobalamin and l,2-dimethyl-4,5-diaminobenzene is
a precursor in the synthesis. Analogs of these substances are often inhibi-
tory to bacterial growth and the biosynthesis of vitamin B^g. 1,2-Dichloro-
4,5-diaminobenzene inhibits the synthesis of vitamin B^g in bacteria and the
HaC^^^^^^X^ ^ Clv^/^^^NHa
,iU,
H,C^ ^^^ ^N^ Cr ^^ ^NHa
5, 6-Dimethylbenzimi- 1, 2-Dichloro-4, 5-
dazole diaminobenzene
growth of those bacteria requiring vitamin B^g (WooUey and Pringle, 1951).
Vitamin B^g is unable to counteract these inhibitions. Although 5,6-dimeth-
ylbenzimidazole can be used by the rat to form vitamin B^g, this sub-
stance is inhibitory to Lactobacillus lactis, and like a number of analogs, is
able to inhibit the synthesis of vitamin B^g in these bacteria (Hendlin and
Soars, 1951). These inhibitions do not appear to be competitive with vi-
590 2. ANALOGS OF ENZYME REACTION COMPONENTS
tamin B^g, but the inhibition by l,2-diamino-4,5-dimethylbenzene is com-
petitive. The only instance of the inhibition of vitamin B^g function is in
the synthesis of methionine from serine in E. coli where the methylamide,
ethylamide, and aniHde analogs of cyanocobalamin inhibit competitively
with respect to vitamin B^j (Guest, 1960). These analogs also inhibit the
growth of organisms that require methionine or vitamin B^g, but do not
when methionine is supplied. The inhibitions are reasonably potent, 61%
depression being given by 0.029 mM of the anilide derivative when cyano-
cobalamin is 0.000032 mM, but the analogs are obviously bound less tightly
than the cyanocobalamin to the enzyme involved. Hydroxocobalamin and
cyanocobalamin, which are analogs of cobalamin coenzyme, inhibit potent-
ly and competitively the diol dehydrase from Aerobacter, but once inhibition
occurs it cannot be reversed by either dialysis or the coenzyme (Lee and
Abeles, 1963).
Lipoate functions in acyl transfer during the oxidation of keto acids and
this is inhibited by 6-ethyl-8-mercaptooctanoate, an analog of 6-acetyl-6,8-
dimercaptooctanoate (a functional form of lipoate) (Albrecht, 1957). This
analog does not inhibit the anaerobic decarboxylation of pyruvate in ex-
tracts from E. coli but inhibits pyruvate oxidation. The phosphotransacetyl-
ase reaction and the formation of acetyllipoate are inhibited.
MrSCELLANEOUS ANALOG INHIBfTiONS
There are a number of reports of inhibitions by analogs that do not readily
fall into any general classification. Some of these have been put into Table
2-38 in order to illustrate further the various types of analog, although in
some cases it is not quite certain whether the inhibitor should be considered
as an analog or not. In most instances the inhibitions are competitive, but
in others it may be noncompetitive or mixed, indicating that the inhibitions
do not all involve a simple competition between the analog and the sub-
strate for an enzyme site. Unfortunately, most of these inhibitions have not
been adequately studied with respect to mechanism. Some more important
examples that could not be easily summarized in the table will be discussed
briefly.
Inhibition of Morphine A/-Demethylase
The antagonistic actions of nalorphine (A'^-allylnormorphine) to the phar-
macological responses to morphine have been extended to the enzyme sys-
tem for morphine inactivation, and it appears that the configurations of
the tissue receptor groups and the enzyme active site are very similar
(Axelrod and Cochin, 1957). Several normorphine analogs were tested on
the A-demethylation of morphine by rat liver enzyme (see accompanying
tabulation) and the role of the alkyl substituent in the binding is evident.
MISCELLANEOUS ANALOG INHIBITIONS
591
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MISCELLANEOUS ANALOG INHIBITIONS
603
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604 2. ANALOGS OF ENZYME REACTION COMPONENTS
Analog (0.2 mM)"
% Inhibition
Relative —AF of binding
(kcal/mole)
iV^-Methallylnormorphine
74
4.90
iV-Isobutylnormorphine
69
4.75
iV^-AUylnormorphine
64
4.61
i\7^-Hexylnormorphine
44
4.13
iV-Butylnormorphine
35
3.88
2\r-Propylnormorphine
20
3.40
iV-Ethylnormorphine
13
3.09
iV-Isopropylnormorphine
11
2.97
Normorphine
0
<2.45
" Morphine = 1 mM.
Some generalizations may be made: (1) increase in chain length increases
the binding energy, (2) unsaturation increases the binding by 1.0-1.2 kcal/
mole, (3) each additional methylene group augments binding by approxi-
mately 0.3 kcal/mole, and (4) the inhibition by nalorphine is noncompetitive
(it was stated that this may be a slow pseudoirreversible inhibition but in-
cubation with morphine and inhibitors was for 2 hr.
Dehydroshikimate Reductase
The inhibition of this enzyme by various phenolic compounds points to
the manner in which the substrate is bound and the configuration of the
active site. Relative binding energies are given in Table 2-39. These inhibi-
tions are all strictly competitive. It is seen that all effective inhibitors have
a p-OH group, and Balinsky and Davies (1961 b) postulated from the pos-
sible ring configurations of shikimate that this group must lie approximately
in the equatorial plane. Additional OH groups increase the binding slightly
or not at all, so that m-OH groups seem to participate little in the binding.
One might expect the carboxylate group to be bound to an enzyme cationic
group, but this does not appear likely; e.g., the addition of a C00~ group
to catechol increases the binding very little, and the benzoates without a
p-OH are bound very poorly. The stronger reaction of the aldehyde group
in vanillin also indicates that the forces here are not merely electrostatic.
Substitution of benzoate in the o-position is detrimental to the binding and
this may be due to steric hindrance, as shown in the diagram of the active
site presented by Balinsky and Davies (Fig. 2-21). The energy of binding
of the p-OH group is greater than 2.3 kcal/mole and thus hydrogen bond-
ing may be involved. It is worth noting that the experiments were run at
MISCELLANEOUS ANALOG INHIBITIONS
605
Table 2-39
Analog Inhibition of Dehydroshikimate Reductase"
Analog
Structure
Relative - AF
of binding
(kcal/mole)
Vanillin
CH3O
HO— (\ V-CHO
5.72
(Shikimate)
Gallate
HO
HO— ( />— COO'
HO
HO
HO
COO
HO
(5.29)'
4.85
/»-Hydroxybenzoate
^0—{\ ,)— COO"
4.13
Protocatechuate
HO
HO— <x /)— COO
4.12
Catechol
HO
HO
3.86
Guaiacol
CH3O
2.78
606 2, ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-39 (continued)
Analog
Structure
Relative - AF
of binding
(kcal/mole)
Phenol
HO
1.84'
2, 4-Dihydroxybenzoate
HO
COO
OH
1.84'
Benzoate
COO
< 1.84'
w -Hydroxybenzoate
HO
<Q>-coo-
< 1.84^^
HO
3, 5-Dihydroxybenzoate
COO'
< 1.84'
Salicylate
<Q^coo-
OH
< 1.84'
«From Balinsky and Davies (1961b).
f> The value for shikimate was obtained from K,,, assuming that this is a true substrate
constant.
'"Values calculated on the basis of inhibitions reported and are very approximate.
pH 9 where the phenolic groups are partially ionized, and this may account
for some of the differences in inhibitory activity between the compounds.
Variation of the inhibition with pH might provide some interesting infor-
mation.
MISCELLANEOUS ANALOG INHIBITIONS 607
Kynurenine Transaminase
The transamination between kynurenine and a-ketoglutarate catalyzed
by an enzyme from rat kidney is inhibited by a variety of mono- and di-
carboxylates (Mason, 1959). The inhibitions are competitive with respect
to kynurenine and are reversible. The results are interpreted in terms of
NAOP
Fig. 2-21. Topographical scheme of
the active site of dehydroshikimate
reductase. (From BaHnsky and Da-
vies, 1961 b.)
two types of interaction: (1) electrostatic binding of carboxylate groups to
cationic groups on the enzyme, and (2) van der Waals' forces between the
hydrocarbon portions of the inhibitors and the enzyme surface. The varia-
tion of inhibition in the dicarboxylate series (Table 2-40) indicates that two
cationic groups interact maximally with adipate. The distance between
these groups was given by Mason as 11 A on the basis of an extended adipate
molecule; the intercarboxylate distance for adipate is given as 6.87 A in
Table 1-1. However, the cationic groups need not be the same distance
apart as the carboxylate groups, and not only is the distance important
but the allowed configuration of the methylene chain to interact maximally
with the enzyme. As Mason points out, the data from the phthalates do
not support this distance entirely, since terephthalate inhibits least of the
three isomers and its intercarboxylate distance is the closest to that of
adipate. The intercarboxylate distance in isophthalate is 5.84 A (Table 1-1)
and this might indicate that the cationic groups are closer than might be
expected from the data on the flexible dicarboxylates, but again there is
the problem of the orientation of the benzene ring. The importance of van
der Waals' interactions is shown by the increasing inhibition given by the
higher fatty acids, and the greater inhibition by the alkyl-substituted glu-
608
2. ANALOGS OF ENZYME REACTION COMPONENTS
Table 2-40
Inhibition of Kynurenine Transaminase by Mono- and Dicarboxylates "
Relative —AF
Inhibitor
Concentration
(mM)
% Inhibition
of binding
(kcal/mole)
Alkyl monocarboxylates
Formate
6
0
<1.43
Acetate
6
0
<1.43
Propionate
6
0
<1.43
Butyrate
6
0
<1.43
Valerate
6
0
<1.43
Caproate
6
1
1.43
Heptanoate
6
8
2.75
Caprylate
6
15
3.19
Nonanoate
6
24
3.55
Caprate
6
41
4.04
Straight-chain dicarboxylates
Oxalate
6
0
<1.43
Malonate
6
0
<1.43
Succinate
6
0
<1.43
Glutarate
6
8
2.75
Adipate
6
65
4.64
Pimelate
6
30
3.74
Suberate
6
2
1.86
Azelaeate
6
35
3.88
Sebacate
6
65
4.64
1,10-Decanedicarboxylate
3
74
5.33
1,11 -Undecanedicarboxylate
3
78
5.47
1 , 14-Tetradecanedicarboxylate
3
87
5.86
Cyclic monocarboxylates
Benzoate
6
0
<1.43
y-Phenylbutyrate
6
9
2.91
Cyclohexanecarboxylate
6
0
<1.43
y-Cyclohexanebutyrate
6
64
4.62
Cyclic dicarboxylates
o-Phthalate
12
14
2.72
Isophthalate
12
35
3.45
Terephthalate
12
9
2.48
Cyclohexane- 1 ,2-dicarboxyiate
6
9
2.91
Glutarate derivatives
2-Methylglutarate
6
15
3.19
2,2-Diniethylglutarate
6
35
3.88
2,4-Dimethylglutarate
6
33
3.82
3-Methylglutarate
6
47
4.18
3,3-Dimethylglutarate
6
56
4.41
3-Methyl-3-ethylglutarate
6
39
3.98
3,3-Diethylglutarate
6
20
3.40
/J-Ketoglutarate
6
0
<1.43
2,2-Dimethylsuccinate
fi
2
1.86
" Kynurenine was 3.7 mM and a-ketoglutarate was 6 mM; pH
Mason, 1959.)
6.3. (From
MISCELLANEOUS ANALOG INHIBITIONS
609
tarates compared to glutarate. One of the cationic groups on the enzyme
seems to have a pK,, around 6.7, since the mhibition by the dicarboxylates
decreases from pH 5.5 to 8.5 and approaches that of the monocarboxylates
(Fig. 2-22). The increase in inhibition of the dicarboxylates with longer
chain lengths than suberate is explained by the ability of the flexible hy-
drocarbon portions to orient for effective interaction with the enzyme sur-
face between or around the cationic groups. The contribution of a methyl-
ene group to the binding is around 0.2-0.4 kcal/mole. It is likely that a
100
VALERATE
5.5
6.0
6.5
7.0
7.5
8.0
8.5
pH
Fig. 2-22. Effects of pH on the inhibitions of kynurenine transaminase by
various fatty acid anions at 6 mM. (From Mason, 1959.)
hydrophobic region of the enzyme lies at some distance from the cationic
groups since the fatty acids do not begin to inhibit until the chain length
reaches 5 or 6 carbon atoms, and the marked differences between the bind-
ing energies of benzoate and y-phenylbutyrate, and cyclohexanecarboxylate
and y-cyclohexanebutyrate, indicate that the ring interaction is effective
when the ring is separated from the carboxylate group by several angstroms.
However, these differences may be due more to steric factors, a ring close
to the carboxylate group interfering with its electrostatic interaction. In-
deed, there seems to be a region for van der Waals' interactions between the
610 2. ANALOGS OF ENZYME REACTION COMPONENTS
two cationic groups on the enzyme, since the 3-substituted gkitarates are
bound more tightly than the 2-substituted isomers.
The administration of nicotinylalanine to rats leads to a 4-fold increase
in the urinary level of A"-methylnicotinamide (Decker et al., 1963). Nicotin-
ylalanine is possibly formed from tryptophan through 3-hydroxykynure-
nine but studies with tryptophan-C^^ indicate it not to be a metabolite but
actually a strong inhibitor of kynureninase and kynurenine hydroxylase.
This inhibition presumably occurs in vivo, resulting in a sequential block
in the major route of kynurenine degradation. The effect of this analog on
the transaminase is not known.
Inhibition of Urease by Methylurea and Thiourea
There is a good deal of disagreement on the analog inhibitions of urease
and the kinetics are certainly not simple. Takeuchi (1933) originally found
very little inhibition (perhaps 5%) by 83 mM methylurea (although mark-
ed inhibition by oxyurea — which may be HgN — CO — NHgOH — occurs,
this is probably not competitive). Sophianopoulos and Corley (1959) ob-
tained competitive inhibition by methylurea at lower concentrations (un-
specified), but higher concentrations (300-1000 mM) increase the substrate
inhibition produced by urea; these latter effects may be related to enzyme
denaturation. The inhibition was found to depend on the pH by Shaw and
Raval (1961), it being noncompetitive between pH 7 and 8.9, and competi-
tive below pH 7. Furthermore, the kinetics correspond to the reaction of 2
molecules of methylurea with the active center, which may relate this type
of inhibition to substrate inhibition.
Thiourea was claimed to stimulate urease at 5 mM (Sizer and Tytell,
1941), to have no effect below 50 mM, and to inhibit 35% at 500 mM
(Kistiakowsky and Shaw, 1953). This inhibition is completely reversible
and occurs rapidly. At pH 6 the inhibition is competitive but as the pH is
raised, deviation occurs. The kinetics again point to 2 molecules of thiourea
reacting with the enzyme. The reaction
E -f 2 I -> EI2
is not affected by change of pH, whereas the reaction
ES + 2 I -> ESl2
is sensitive to pH, which serves to explain the change in inhibition type
with the pH. Lister (1956) reported that of the 17 urea analogs tested,
only thiourea is inhibitory — 35% at 200 mM, 70% at 400 mM, and 85%
at 1000 mM, when urea is 500 mM. The inhibition is prevented by cysteine,
which brings up the possibility of disulfide bond formation by thiourea at
high concentrations.
MISCELLANEOUS ANALOG INHIBITIONS 611
Inhibition of Catechol-0-methyitransferase by Pyrogallol
This inhibition is of interest because of the bearing it has on the metab-
olism of epinephrine and norepinephrine. Bacq (1936) observed that pyro-
gallol increases the responses of tissues to sympathetic nerve stimulation and
to epinephrine. However, he then attributed this action to the antioxidant
properties of pyrogallol. Lembeck and Resch (1960) and Vanov (1962) have
recently confirmed this by showing that the pressor response to epineph-
rine is prolonged by pyrogallol. The inhibition of the catechol-0-methyl-
transferase was reported by Bacq et al. (1959), who believed that this could
explain the sensitization of smooth muscles to the catecholamines by pyro-
gallol and other phenolic compounds. Axelrod and Laroche (1959) also
found a potent inhibition of this enzyme (50% when pyrogallol = epineph-
rine = 0.01 niM), which decreases with increasing substrate concentration,
indicating a competitive action. Furthermore, about 70% of intravenously
injected epinephrine-H^ is metabolized in 10 min in mice, but pretreatment
with 100 mg/kg pyrogallol reduces the amount metabolized to 22%. The
half-life of norepinephrine in mice is increased from 22 to 42 min by 10 mg
pyrogallol, while at the same time 0-methylation is inhibited 99%, indi-
cating other pathways for norepinephrine metabolism (Udenfriend et al.,
1959). Probably the monoamine oxidase pathway is also important. The
administration of pyrogallol to rats does not by itself increase brain nor-
epinephrine levels, but in conjunction with iproniazid (which inhibits mo-
noamine oxidase) it does, in this case the two major degradative pathways
being blocked (Jaattela and Paasonen, 1961). This is a good example of
the action of two inhibitors on a divergent multienzyme system and, in
addition, has interesting possibilities for clinical application.
Repeated administration of pyrogallol causes a rise in the blood pressure
but this is soon followed by a loss of response or tachyphylaxis (Wylie et
al., 1960). The rate of urinary excretion of 0-methylated derivatives of the
catecholamines is briefly decreased by pyrogallol, but if the administration
is continued the rate returns to normal (Nukada et al., 1962). Long-term
treatment with pyrogallol leads to an increase in 0-methyltransferase and
monoamine oxidase in the liver of rats, so it may well be that these enzymes
are adaptively altered. The urinary excretion changes of the catecholamines
and their 0-methylated products are shown in Fig. 2-23.
The kinetics of the in vivo inhibition have been studied by Crout (1961).
Inhibition of 0-methyltransferase occurs very rapidly in liver, heart, and
brain even when the pyrogallol is injected intraperitoneally, and by 30 min
has developed appreciably. The inhibition of the enzyme obtained from rat
tissues, however, is only partly competitive (actually the curves appear to
indicate pure noncompetitive inhibition) despite the fact that pyrogallol
is a substrate for the enzyme. The K, of 0.008 vs\M for pyrogallol indicates
the high potency of the inhibition (^,„ for norepinephrine is 0.3 vaM). Fur-
612
2. ANALOGS OF ENZYME REACTION COMPONENTS
ther work on the exact^mechanism of this inhibition might provide interest-
ing information. Wylie et al. (1960) found pyrogallol to be the most potent
inhibitor of a series of polyphenols and epinephrine analogs, inhibiting 50%
at 0.03 mM when epinephrine is 0.3 mM. However, gallate, adrenalone, and
arterenone are almost as active. A new inhibitor of 0-methyltransferase was
studied briefly by D'lorio and Mavrides (19^3). This is 3.5-diiodo-4-hy-
droxybenzoate and it inhibits the rat liver enzyme competitively with
Ki = 0.013 mM.
Inhibition of the Oxidation of Aromatic Compounds by Bacteria
The metabolism of o-nitro- and p-nitrobenzoate by Nocardia is quite
strongly inhibited by m-nitrobenzoate, o-nitrophenol, and p-nitrophenol
(Cain, 1958). The competitive nature of this interference was demonstrated
by reciprocal plotting. The enzymes involved here are not well characterized
and, indeed, the action could be on a transport mechanism at the mem-
+ 100
+ 50
%
CHANGE
-50
-100
METANEPHRINE
AND
NORMETANEPHRINE
30
DAYS
Fig. 2-23. Effects of pyrogallol injected subcutaneously
at a dose of 20 mg/kg on the urinary excretion of free
and methylated catecholamines by rabbits. (From
Nukada et al, 1962.)
MISCELLANEOUS ANALOG INHIBITIONS 613
brane. Durham and Hubbard (1959, 1960) favor competition for a transport
system in the inhibition by p- aminosalicylate of the oxidative assimilation
of p-aminobenzoate in Flavobacterium. In the presence of ^-aminosalicylate
there is more p-aminobenzoate remaining in the medium and almost com-
plete inhibition of uptake is seen at a p-AS/p-AB ratio of 10. p-Aminoben-
zoate may be not only a necessary metabolite for folate synthesis, but also
the principal source of energy for growth. It is very difficult in such cases
to determine whether the inhibition is on a surface transport or an intra-
cellular enzyme until the enzymes responsible for the metabolism have been
isolated and examined.
Inhibition of Acetate and Fat Metabolism by Propionate
The original work in this field was done by Jowett and Quastel (1935 a,b).
The transformation of butyrate into acetoacetate in guinea pig liver slices
is inhibited strongly by benzoate, /9-phenylpropionate, and cinnamate. Pro-
pionate also inhibits but more weakly (59% at 10 mM). Much later Felts
et al. (1956) reported that 4 mM propionate almost completely depresses
the incorporation of acetate- 1-C^* into fatty acids in rat liver slices. The
formation of C^Og is also suppressed. Propionate is known to be inhibitory
to the growth of many bacteria and fungi, so the question of the mechanism
of its action is of some importance. It has often been attributed to a com-
bination with and depletion of coenzyme A. This inhibition has been studied
most thoroughly by Pennington (1956, 1957), who found that the reaction
acetate- 1-C^* -^ C^^Og in rat liver can be inhibited readily and almost com-
pletely, while simultaneously the total amount of acetate disappearing is
reduced. This also occurs in kidney, heart, and diaphragm, but to a lesser
extent. Even concentrations as low as 0.5 mM are 40% inhibitory in the
liver. It was felt that propionate blocks both the uptake of acetate and the
formation of acetyl-CoA. The oxidation of pyruvate is inhibited much less
and that of butyrate not at all. However, most of the action must be on
the intracellular metabolism inasmuch as marked inhibition is seen in liver
homogenates (Pennington and Appleton, 1958). Addition of coenzyme A
in the presence of propionate increases the amount of COg formed from
acetate slightly but does not reverse the inhibition, indicating that a simple
depletion of coenzyme A is not the mechanism. It was postulated that
perhaps propionate inhibits after being metabolically altered, possibly to
propionyl-CoA, or directly inhibits acetyl-CoA synthetase. This is an in-
teresting and metabolically important inhibition so that one looks forward
to studies on the enzymes involved in acetate metabolism.
A few instances of the inhibition of acyl-CoA metabolism by analogs have
been reported. Tetrolyl-CoA and propiolyl-CoA, the acetylenic analogs of
butyryl-CoA and propionyl-CoA, respectively, are potent noncompetitive
inhibitors of fatty acid synthesis in brain and liver extracts (Brady, 1963;
614 2. ANALOGS OF ENZYME REACTION COMPONENTS
Robinson et al., 1963). The mechanism of inhibition here, however, appears
to be by reaction with enzyme SH groups. The coenzyme A is necessary
since free tetrolate does not inhibit, and is split off during the reaction.
Palmityl-CoA noncompetitively inhibits the condensing enzyme with respect
to acetyl-CoA, and perhaps competitively with respect to oxalacetate (Wiel-
and and Weiss, 1963). The inhibition develops slowly and the authors sug-
gest configurational changes in the enzyme, although there is no direct
evidence for this. Such an inhibition might be regulatory with regard to
the operation of the cycle and the formation of acetoacetate in the liver by
controlling the rate of oxidation of acetyl-CoA through the cycle. Higher
acyl-CoA's inhibit rat liver acetyl-CoA carboxylase very strongly, oleyl-
CoA and stearyl-CoA being competitive with K/s of 0.0013 milf and 0.00071
mM, respectively, this possibly playing an important role in the homeo-
static control of fatty acid synthesis (Bortz and Lynen, 1963).
Inhibition of cholesterol biosynthesis will be discussed in a subsequent
chapter, but it is worthwhile mentioning at this point that a-phenyl-n-bu-
tyrate not only lowers serum cholesterol but also inhibits the incorporation
of acetate into fatty acids, whereas the oxidation of acetate is only weakly
inhibited (Steinberg and Fredrickson, 1955). The evidence points to an ac-
tion early in acetate metabolism, possibly the acetylation of coenzyme A
or transacetylations from acetyl-CoA.
Antagonism between Tungstate and Molybdate
Molybdate is a necessary cof actor in the growth of many organisms and
has been found to participate in certain enzyme reactions, such as those
catalyzed by xanthine oxidase and nitrate reductase. If chicks are fed on
a low-Mo diet containing 4.5-9.4% mg sodium tungstate, the growth rates
are depressed and signs of molybdenum deficiency appear (Higgins et al.,
1956 a). The levels of molybdenum in the tissues fall to less than 10% of
the normal and xanthine oxidase activity is severely depressed, leading to
an alteration in the excretory pattern of purines. These changes are re-
versed by adding 2-6 mg% sodium molybdate to the diet. Similar falls in
xanthine oxidase were observed in rats.
Aspergillus niger requires molybdate especially when nitrate is the sole
source of nitrogen, since the enzymic reduction of nitrate by nitrate reduc-
tase involves molybdate as a prosthetic group (Higgins et al., 1956 b). Tung-
state is able to compete with molybdate and inhibition of growth occurs
when the (tungstate)/ (molybdate) ratio is 20. Azotobacter vinelandii is like-
wise inhibited by tungstate when nitrogen or nitrate is the source of amino
acids and proteins, but not when ammonia is provided. The uptake of Mo^^
by the cells is also depressed by tungstate. The ability of tungstate to inhibit
growth is dependent on the level of molybdate in the medium and it requires
rather high ratios of (tungstate)/ (molybdate) to inhibit well (Bulen, 1961).
MISCELLANEOUS ANALOG INHIBITIONS 615
The question of the site, or sites, of tungstate inhibition is not yet settled.
Bulen believes that the primary effect is a depression of molybdate uptake
and has provided evidence that that there is no antagonism of the enzymi-
caUy functioning molybdate. The growth of the crown-gall organism Agro-
bacterium tumefaciens is also inhibited in nitrate medium (about 50% at
0.05 milf), but when ammonia is added there is only slight inhibition at
1 ToM (Kurup and Vaidyanathan, 1963). Molybdate antagonizes the growth
depression. A decrease in nitrate reductase activity during the inhibition
was observed. Before the problem of the site of inhibition can be finally
settled, more work must be done on isolated molybdenum-dependent en-
zymes. The NADH-dependent nitrate reductase from wheat is not inhibited
by 1 mM tungstate (Spencer, 1959). In any event, this represents a unique
type of competitive inhibition which is basically due to the similar structures
and properties of tungstate and molybdate.
Inhibition of Penicillinases
Some of the data are summarized in Table 2-38, and it is evident that the
penicillinases from different bacteria exhibit various patterns of sensitivity.
In particular, the gram-negative and gram-positive organisms possess dif-
ferent types of enzyme, the former usually being more sensitive to penicillin
analogs. The inhibitions are generally competitive. The exact mechanism of
the inhibition, however, is not clear, since there is some evidence that the
analogs alter the configuration of the enzyme (Garber and Citri, 1962; Citri
and Garber, 1963). In the first place, the analogs accelerate the tempera-
ture inactivation of penicillinase and, in the second place, the inhibition by
6-(2,6-dimethoxybenzamido)penicillanate is accompanied by the appear-
ance of groups sensitive to iodine. These penicillinase inhibitors are of use
in determining the mechanism of penicillin resistance in bacteria; if the
resistance is due to the increased synthesis of penicillinase, the inhibitor
wiU abolish the resistance. (Hamilton-Miller et al., 1964).
Inhibition of Cycle Enzymes by y-Hydroxy-a-ketoglutarate
Glyoxylate and oxalacetate condense under physiological conditions to
form a product which inhibits certain steps in the tricarboxylate cycle.
The reaction is quite rapid at 20° and pH 7.4, and since both glyoxylate
and oxalacetate are produced normally in cells, it was of some interest to
study the properties of this condensation product, which was considered to
be «-hydroxy-/?-oxalosuccinate by Ruffo et al. (1962 a). It was found to be
competitive with respect to citrate and cis-aconitate, and to inhibit 50%
at concentrations around 0.12 mM. When glyoxylate is added to mito-
chondria there is respiratory inhibition and some accumulation of citrate
if oxalacetate is present, which is due to the inhibition of aconitase and
616 2. ANALOGS OF ENZYME REACTION COMPONENTS
isocitrate dehydrogenase (Ruffo et al., 1962 b; Ruffo and Adinolfi, 1963).
If no oxalacetate is present, glyoxylate directly inhibits the oxidations of
a-ketoglutarate and succinate, so that two sites of cycle inhibition can occur,
one by glyoxylate alone and one by its condensation product with oxalace-
tate. Payes and Laties (1963) have claimed that the of-hydroxy-/?-oxalosuc-
cinate (oxalomalate) initially formed in the condensation reaction is rapid-
ly decarboxylated to y-hydroxy-or-ketoglutarate, which is the actual in-
hibitor. y-Hydroxy-a-ketoglutarate competitively inhibits yeast aconitase
[K^ = 0.14 mM), potato a-ketoglutarate dehydrogenase {K^ = 0.7 mM),
and isocitrate dehydrogenase. It is also moderately inhibitory to the res-
piration of potato slices, 35% depression being observed with 2 mM and
71% with 5 mM. It is rather surprising that it does not serve as a sub-
strate for a-ketoglutarate dehydrogenase and enter into the sequence of
reactions involving coenzyme A, as does y-methyl- y-hydroxy-or-ketogluta-
rate (parapyruvate).
CHAPTER 3
DEHYDROACETATE
Since this substance inhibits succinate dehydrogenase and may exert some
of its effects because of a structure analogous to certain cellular metabolites,
it is appropriate to discuss it at this time, although little of its basic meta-
bolic effects is understood. In a study of the formation of ethyl acetoacetate,
Geuther (1866) found that distillation of this substance yielded a crystalline
material, to which he gave the name dehydroacetic acid. It was soon shown
to contain a pyran ring structure (Feist, 1890) and has been used extensively
in organic preparative procedures. The effects on biological systems were not
studied until Brodersen and Kjaer (1946) in Copenhagen investigated a se-
ries of unsaturated lactones for antibacterial activity. They reasoned that
several such compounds are antimicrobial — anemonin from species of Ra-
nunculaceae, parasorbic acid from the mountain ash (Sorbus), Dicumarol
from sweet clover, penicillic acid from certain species of Penicillium, kojic
acid from Aspergillus, patulin (clavacin) from various fungi, and others —
and that a correlation between activity and the — 0 — CO — C=C — struc-
ture might exist:
Q 0
^O o
OH
H,C
.CH3
^CH,
OCH,
Anemonin
Parasorbic acid
Penicillic acid
O^ .OH
O^ ^CH,OH
Patulin
Kojic acid
617
618 3. DEHYDROACETATE
Dehydroacetic acid was found to be rather inactive against most of the 25
types of bacteria studied and no further mention was made of it. McGowan
et al. (1948) investigated 80 compounds with ethylenic linkages for the pur-
pose of correlating fungistatic activity with the abihty of the substituents
to withdraw electrons from these double bonds, and found dehydroacetate
to exert very little effect. An investigation stimulated by the antibacterial
effects of usnic acid from lichens led Ukita et al. (1949) to examine dehydro-
acetate, which they found to inhibit staphylococci and mycobacteria slight-
ly; however, other substances were more potent and of greater interest.
The Dow Chemical Company meanwhile had been studying the antimicro-
bial action of dehydroacetate and on June 28, 1949 issued three patents for
its use in food preservation.* These were based on work started in 1946 in
cooperation with the Department of Pharmacology of the University of
Michigan Medical School, the results of which were published in a series of
papers in 1950. Almost all of our present basic knowledge of dehydroacetate
stems from this work and essentially no fundamental biochemical reports
have been made since, although a great many papers on its practical use
appear annually.
CHEMICAL PROPERTIES
The structure of dehydroacetic acid was debated for many years until
Rassweiler and Adams (1924) proved that the formula suggested by Feist
(1890) is basically correct. The dipole moment of 2.83 was claimed by Le-
Dehydroacetic acid
Fevre and LeFevre (1937) to be consistent with this structure if restrictions
were imposed on the rotation of the acetyl group. The formula is commonly
written in the keto form, but it is likely that a tautomeric equilibrium with
the following enolic forms occurs:
H,c. o ^o H,C
(a) (b) (c)
* Dehydroacetate is actually a rather weak inhibitor of microbial growth but can
be used to ^reserve food because it is so little toxic to humans.
CHEMICAL PROPERTIES 619
Hydrogen bonding of the type suggested by Forsen and Nilsson (1961) is
represented; this undoubtedly occurs in the pure form but may not in
aqueous solution. Proton magnetic resonance and infrared spectra suggest
structure (b), but the other forms are not excluded. The restriction of the
acetyl group rotation may derive from such hydrogen bonding.
Ionization
It is important with regard to the action of dehydroacetate on enzymes
and its penetration into cells to determine the predominant forms in aqueous
solution at physiological pH. It is usually stated that dehydroacetic acid is
a very weak acid, and Wolf and Westveer (1950) remarked that it would
exist primarily in the ionized state at pH 9. The acidic property is the result
of enolization and each of the enolic forms shown above could lose a proton
to form the corresponding anion. However, once this occurs, resonance be-
tween the three forms is possible, stabilizing the anion and increasing the
acidity and, furthermore, producing a more diffuse negative charge. In order
to determine the state of the inhibitor under physiological conditions, titra-
tions were done, starting at pH 3.64 (the pH of a saturated solution). It
was found that two equivalents of base are taken up between pH 4.2 and
6.4, with a mean pK^ of 5.20, so that around pH 7 dianions of the a-pyran
and y-pyrone type must be present.
H,C ^O^ ^O H,C
O
-CH,
General Properties
Dehydroacetic acid has an absorption peak around 313 m// (Calvin et al.,
1941), sublimes at 109°, is fairly soluble in organic solvents but poorly sol-
uble in water (around 0.25% at 37°). The sodium salt, however, is quite
soluble in water (33%) (Wolf, 1950) and sufficiently stable in solution for
most purposes. It may be catalytically hydrogenated with PtOa to give the
corresponding 3-ethyl compound (Malachowski and Wanczura, 1933), or
with Ni under pressure to yield more completely hydrogenated forms, cor-
responding to the uptake of 3-5 moles of Hg (Adkins et al., 1931).
Synthesis
Dehydroacetic acid was first obtained by Geuther (1866) by distilHng the
ethyl ester of acetoacetate, and Conrad (1874) found that a reasonable yield
could be obtained by heating this substance under pressure. The present
620 3. DEHYDROACETATE
method of synthesis involves heating ethyl acetoacetate with a small amount
of NaHCOg at 200o-210o for 7-8 hr with subsequent distillation at 1280-140°
in vacuo, the yield being over 50% (Arndt, 1955). It is also formed by the
tetramerization of ketene, and by suitable catalytic means may be almost
quantitatively obtained from diketene (Steele et al., 1949), the reaction
presumably proceeding through the enolized forms of the diketene. The
simplest method of purification of dehydroacetic acid is probably recrys-
tallization from ethanol.
Estimation
Two methods suitable for tissue analyses were developed by Woods et al.
(1950). A colorimetric test, based on the reaction of the acetyl group with
salicylaldehyde in alkaline solution to give a red-orange color, is sensitive
in the range 10-200 //g. The spectrophotometric test, based on absorption
at 312 m//, is approximately 10-fold more sensitive. Both tests depend on
the proper pre treatment and extraction of the tissue samples, since neither
test is particularly specific. The spectrophotometric tests give the best re-
coveries and are preferable for most analyses.
INHIBITION OF ENZYMES
Dehydroacetate at concentrations between 2.3 and 93 vaM progressively
depresses the oxygen uptake of slices and minces of cerebral cortex and
kidney respiring endogenously, but the excess oxygen uptake following ad-
dition of various substrates is inhibited only in the case of succinate (Seevers
et al., 1950). This suggested that dehydroacetate might inhibit succinate
oxidase and thus this enzyme was examined in some detail.
Succinate Oxidase
Inhibition of succinate oxidase is proportional to the logarithm of the
dehydroacetate concentration from 27 to 77% inhibition (Seevers et al.,
1950). Although no data were given, it was stated that the substrate con-
centration has little or no effect on the degree of inhibition, indicating it is
not competitive. If this is true (see page 621), if ^ would be around 11.6 xaM
indicating much less affinity of the enzyme for dehydroacetate than for mal-
onate. The site of inhibition in the succinate oxidase sequence was determin-
ed in two types of experiment. In the first, the cytochrome system was block-
ed by cyanide and cresyl blue added as a hydrogen carrier; dehydroacetate
inhibited this system to the same degree as the normal one, indicating the
action not to be on the cytochrome system (see accompanying tabulation).
In the second, the cytochrome system was studied directly and no inhibition
INHIBITION OF ENZYMES 621
Cyanide
Cresyl blue
Dehydroacetate
Oj Uptake
°/o Inhibition by
(mM)
(mM)
(mJ/)
{/a/30 min)
dehydroacetate
165
2
—
—
0
—
2
6
—
88.5
—
—
—
9.3
84
49
2
6
9.3
43.5
51
by dehydroacetate was observed. It was concluded that dehydroacetate acts
on succinate dehydrogenase. Although this is probably true it is evident
that, according to the modern elaboration of succinate oxidase, other sites
are possible.
Evidence that dehydroacetate does not inhibit by reacting with the SH
groups of succinate dehydrogenase was obtained. First, the inhibition is al-
most instantaneous and readily reversible, unlike inhibitions with most SH
reagents. Second, cysteine, glutathione, and dimercaprol are unable to pro-
tect the enzyme against dehydroacetate.* Third, no inhibition of urease
was observed, this enzyme being sensitive to most SH reagents; indeed
stimulation was observed. All in all, one must conclude that the possibility
of the reaction of dehydroacetate with SH groups has not been eliminated,
although there is little positive evidence for such a mechanism.
The possibility of competitive inhibition of succinate dehydrogenase can-
not be eliminated since no data were given for the statement, "Increasing
the substrate concentration does not appreciably alter the degree of inhi-
bition." If, for example, a succinate concentration of 50 mM was used
(which is the only concentration mentioned in the paper) with dehydro-
acetate at 9.3 mM, increasing the succinate to 100 or 200 m.M would not
be expected to reduce the inhibition markedly. It is interesting to speculate
that the dianionic forms of dehydroacetate have a basically similar charge
distribution to malonate. However, due to resonance the charge magnitude
H H
^ ^C^ CH, O^ X ^^
c c c c
o~ o" o' o"
Dehydroacetate Malonate
* Unfortunately the cysteine and glutathione were used at only about one fifth the
dehydroacetate concentration, so that even total reaction of the inhibitor would have
reduced the inhibition relatively little, actually about 5%. Cysteine reduced the inhi-
bition around 5% from the predicted value but glutathione did not. Cavallito and
Haskell (1945) mentioned that dehydroacetate does not react with cysteine.
622 3. DEHYDROACETATE
on dehydroacetate. might be lower than on malonate. The succinate dehy-
drogenase from calf thymus nuclei is inhibited somewhat more potently by
dehydroacetate at pH 6.6 than at pH 7.6 (see accompanying tabulation)
Yo Inhibition
I)
pH6.6
pH7.6
1
33
25
5
56
42
10
67
47
(McEwen et al., 1963 a). This does not fit the dianion inhibition theory very
well, since at pH 7.6 there should be more of the dianion than at pH 6.6.
One cannot attribute the change in inhibition to the ionization of enzyme
groups because malonate inhibits better at the higher pH from the limited
data provided. Unfortunately the succinate concentration was unvaried
from 20 mM and the formal nature of the inhibition remains unknown.
The mechanism of the inhibition of succinate dehydrogenase is thus at
present unsolved.
One further experiment deserves brief mention. It was claimed that al-
though malonate protects succinate oxidase from SH reagents, it does not
protect against dehydroacetate (see accompanying tabulation). First, one
Dehydroacetate (milf ) Malonate {mM)
% Inhibition
of O2 uptake
9.3
9.3
—
49
0.33
33
0.33
65
would not expect malonate at a concentration inhibiting only 33% to pro-
tect very much. Second, the inhibition given by both inhibitors is exactly
what would be predicted if both acted at the same site on the enzyme. No
conclusions as to the mechanism of inhibition can be deduced from this ex-
periment.
Other Enzymes
Dehydroacetate has no effect on cholinesterase up to 20 mM (Seevers et
al., 1950), or on pepsin, amylase, and trypsin at 8.5 mM (Bauer and La Sala,
1956), while urease is stimulated by concentrations up to 93 mM and pan-
EFFECTS ON RESPIRATION AND GLYCOLYSIS
623
creatic lipase is stimulated at 8.5 mM. The ATPase of pea mitochondria is
also stimulated by dehydroacetate, phosphate splitting being increased
around 200% by 1 mM (Forti, 1957), which might be related to the un-
coupling action reported by MarreeiaZ. (1956). Catalase is inhibited weakly
(K^ = 22 jnM) by dehydroacetate compared with other organic acids (Liick,
1957). The enzymes responsible for the destruction of mitomycin in Strepto-
myces mycelia are not affected by 4.8 mM dehydroacetate (Gourevitch et
al., 1961). It is difficult to explain some of the actions on metabolism with
this limited amount of information.
EFFECTS ON RESPIRATION AND GLYCOLYSIS
The effects of dehydroacetate on the endogenous respiration of minces of
various rat tissues are shown in Fig. 3-1. Brain and kidney respiration is
progressively depressed but muscle is anomalous in that marked stimulation
is observed at high dehydroacetate concentrations, while liver is stimulated
■3 -2
LOG (DEHYDROACETATE) ( Ml
Fig. 3-1. Effects of dehydroacetate on the respi-
ration of rat tissue minces measured over a period
of 2 hr. (From Seevers et al., 1950.)
moderately at all concentrations used. Brain slices respond as do the minces,
but liver slices are unaffected by dehydroacetate up to 9.3 mM and are
then depressed at higher concentrations. Mudge (1951) found 10 mM dehy-
droacetate to depress rabbit kidney slice respiration 25%, which is similar
to the inhibition reported by Seevers et al. (1950) in rat kidney. The stim-
ulation of respiration in muscle and liver might result from metabolism
624 3. DEHYDROACETATE
of dehydroacetate; this will be discussed later (page 629), but Shideman
et al. (1950 b) do not think it is likely. The endogenous respiration in such
minces and slices is poorly understood, so that it is difficult to interpret
these results, and the data when glucose is present are inconsistent. It is
also difficult to relate these changes in respiration to inhibition of succinate
oxidase (the effects of malonate and dehydroacetate are quite different),
and such statements as "It appears probable that the manifestations of
toxicity result largely, if not exclusively, from a specific type of chemical
(or physicochemical) action involving interference with oxidative or other
enzyme mechanisms which proceed by way of the Krebs cycle" (Shideman
et al., 1950 b) appear to have little basis, particularly since the effects of
dehydroacetate on the operation of the cycle (as in mitochondrial prepara-
tions) have not been studied. It is significant that dehydroacetate at 4.7-
9.3 TaM stimulates the anaerobic glycolysis in rat brain mince 40-50%
(See vers et al., 1950), an effect greater than any observed on respiration.
On the other hand, 50 mM dehydroacetate inhibits the formation of C^^Og
from glucose-6-C^* 87% in suspensions of isolated thymus nuclei, simultane-
ously the O2 uptake being depressed only 14% and the ATP level falling
27% (McEwen et al, 1963 b). Malonate at 10 mM has very little effect and
this was attributed to a failure to penetrate into the nuclei; dehydroacetate
either penetrates better than malonate or exerts an effect other than inhi-
bition of succinate oxidation.
EFFECTS ON TISSUE FUNCTIONS
Dehydroacetate in the whole animal produces changes in central nervous
system, cardiovascular, and renal functions. Only the renal effects have been
investigated in detail. In addition, the actions on the isolated intestine have
been studied relative to the metabolic disturbances produced.
Intestine
The contractile amplitude of isolated rabbit intestine is depressed slightly
by 1 mM and markedly by 10 vclM dehydroacetate (see tabulation below)
Dehydroacetate
(mM)
Substrate
% Inhibition
of amplitude
1
Acetate
2
1
Glucose
8
10
Acetate
72
10
Glucose
54
EFFECTS ON TISSUE FUNCTIONS 625
(Weeks et al., 1950). When the intestine is allowed to contract for 30-60 min
in the absence of substrate the amplitude is reduced to 15-35% of normal.
The addition of glucose, acetate, or pyruvate allows recovery, essentially
complete in the case of glucose. Dehydroacetate at 10 mM effectively pre-
vents this recovery with acetate and pyruvate, but only partially counter-
acts the effect of glucose. This was taken to mean that dehydroacetate blocks
the cycle preferentially. Malonate neither depresses the amplitude in the
presence of substrates, nor prevents the recovery of substrate-depleted in-
testine upon addition of substrates. This could be related to the poor pen-
etration of malonate into the cells. Dehydroacetate, even though it may
have a double negative charge, might penetrate better than malonate be-
cause the lipophilic fraction of the molecule is greater (sodium dehydroace-
tate is fairly soluble in a number of organic solvents).
Heart
Dehydroacetate given intravenously to dogs causes some slowing of the heart
rate at a dose of 300 mg/kg (Seevers et al, 1950), but in general the effects on
the cardiovascular system are minimal. Like many substances with the
— CH = CH — CO — grouping, dehydroacetate exerts a positive inotropic ac-
tion on hypodynamic cat papillary (Bennett et al., 1958). Although not so
potent as many other compounds, it is active at 5 mM and rated the same
as /3-angelicalactone. Isolated rat atria are depressed quite markedly by
5-10 mM dehydroacetate, and to some extent even by 1 mM, and this is
accompanied by rather unique effects on the membrane potentials (Webb
and Hollander, 1959). The resting and action potential magnitudes are re-
duced more strikingly than with most metabolic inhibitors. However, the
action potential duration is actually increased, due to a slowing of repolar-
ization, an effect observed with no other inhibitor. The depolarization rate
is unaffected, so that the moderate slowing of conduction noted must be
related to the reduced action potential. Malonate at higher concentration
scarcely alters the properties of such atria, again indicating a difference
either in penetration or in action.
Renal Transport
When dehydroacetate is injected into dogs in amounts sufficient to give
plasma concentrations of 20-25 mg% (1.2-1.5 mM), the renal tubular trans-
port of certain substances is markedly depressed — p-aminohippurate 81%,
phenolsulfonphthalein 80%, penicillin-G 60%, and iV-methylnicotinamide
77%, while the transport of glucose, creatinine, and phosphate is unaltered
(Shideman et al., 1950 b). Since diuresis is observed after dehydroacetate,
it is likely that water and ion transport is also affected to some extent. It
is thus clear that certain transport systems are inhibited and others are
626 3. DEHYDRO ACETATE
untouched by dehydroacetate. The mechanism is entirely tubular and no
changes in femoral flow, renal blood flow, or glomerular filtrate rate occur
(Shideman and Rene, 1951 b). The action is similar to that of carinamide
2)-(benzylsulfonamido)benzoate, a previously used blocker of penicillin ex-
cretion. Indeed, dehydroacetate at 0.5 g every 6 hr prolongs penicillin blood
levels in patients (Schimmel et at., 1956).
The active accumulation of phenolsulfonphthalein (Rathbun and Shide-
man, 1951), phenol red (Shideman and Rene, 1951 b), and p-aminohippu-
rate (Shideman and Rene, 1951 b; Farah and Rennick, 1956) in kidney
slices is readily inhibited by dehydroacetate. The effect on ^J-aminohippu-
rate uptake is shown in Fig. 1-18, from which it is seen that 50% inhibition
is given by 0.14 mM in dog kidney slices. The accumulation of tetraethylam-
monium ion is completely resistant to dehydroacetate, and it is believed that
the transport of this ion is not dependent on the cycle (Farah and Rennick,
1956; Farah, 1957).
It appears that certain renal transport mechanisms are more sensitive to
dehydroacetate than any other cell functions examined. The question as to
the relation of this inhibition to succinate oxidase or cycle depression is dif-
ficult to resolve. Dehydroacetate might block transport by acting directly
on the carrier system or by reducing the energy available for the transport,
Shideman and Rene (1951 b) incline to the latter view and attribute the
inhibition to an action on the cycle. The evidence for this comes partially
from the observation that high concentrations of acetate are able to coun-
teract the effects of dehydroacetate, both in vivo and in slices (Stoneman
et al., 1951; Rathbun and Shideman, 1951; Shideman and Rene, 1951 a).
However, the ability of acetate to reverse an inhibition is not evidence for
an action on the cyle, much less on succinate oxidase; indeed, the opposite
might be justifiably concluded. Furthermore, there is not a good correlation
between the activity in depressing p-aminohippurate transport and the in-
hibitory potency on succinate oxidation, especially when carinamide is con-
sidered, this substance being a weak succinate oxidase inhibitor but a more
potent transport inhibitor than dehydroacetate. Of course, different pene-
trabilities into the renal cells may account in part for this lack of correlation.
Ion transport in kidney slices is also inhibited by dehydroacetate (Mudge,
1951). Rabbit kidney slices were leached in 0.15 M NaCl to lower the in-
tracellular K+, and then incubated in 10 mM K+ with 10 vaM acetate as
the substrate, during which period the lost K+ is regained and the excess
intracellular Na+ is pumped out. Dehydroacetate at 10 mM inhibits this
K+-Na+ exchange 81%, while simultaneously the respiration is inhibited
only 25%. The action of dehydroacetate on the pH-regulating exchanges
of the kidney is not known, but may be important in contributing to the
acidosis observed in whole animals, and indirectly in the effects on certain
tissues such as the central nervous system.
EFFECTS ON THE WHOLE ANIMAL 627
EFFECTS ON THE WHOLE ANIMAL
The potential use of dehydroacetate as a food preservative led Spencer
et al. (1950 a,b) of the Dow Chemical Company, and Seevers et al. (1950) of
the University of Michigan, to investigate the effects on various animals and
man when administered by different routes for varying durations of time.
These studies were very thorough and our knowledge at this level of action
is better than for most inhibitors. We shall summarize only the more im-
portant aspects relative to the metabolic disturbances produced.
Acute Toxicity
The earliest evidence of toxicity in rats, dogs, and monkeys when de-
hydroacetate is given by any route is loss of appetite. As the dosage is in-
creased, various sjTnptoms related mainly to the central nervous system
appear: ataxia, salivation, emesis, incoordination, weakness, and stupor,
followed by muscle twitching and a gradual increase in muscle tone, passing
into clonic and tonic convulsions which persist until death, which is attri-
buted to respiratory i)aralysis. During the later phases, convulsions may be
initiated by excess sensory stimulation, indicating a general increase in the
reflex excitability. The effects are produced rather slowly and death occurs
usually after 24-72 hr. The acute oral toxicity in rats is given as: LD^.i =
0.52 g/kg, LDi6 = 0.80 g/kg, LD50 = 1.0 g/kg, LDg^ = 1.23 g/kg, and
LD99.9 = 1.92 g/kg.
The initial effects of intravenous injection (0.3-0.4 g/kg) are probably
related to the temporary disturbance in blood pH if the solutions are not
neutralized. The alterations in nervous system function may not be specific
and due to the direct action of dehydroacetate on the nerve cells, but de-
pendent on the developing acidosis. An initial respiratory alkalosis in dogs
is soon followed by a shift toward metabolic acidosis, compensated at first
but later becoming uncompensated. As the plasma pH suddenly drops to
levels approaching 7, with simultaneous decreases in plasma bicarbonate
and Pqq , convulsions occur. We have noted that renal transport is more
sensitive to dehydroacetate than is nerve or muscle respiration, especially
in the presence of glucose. The plasma levels of dehydroacetate in acute
poisoning are probably between 1 and 3 mM in most cases, which, coupled
with the possible concentration in the kidney, could easily disturb renal
function seriously. The concentration in the central nervous system during
poisoning appears to be quite low (see page 631). All of this points to an
indirect action on the nervous system and perhaps a primary renal site for
the toxicity.
Chronic Toxicity
Rats fed diets containing 0.02-0.10% dehydroacetic acid for 2 years
showed no obvious adverse effects on growth, mortality, hematology, organ
628
3. DEHYDROACETATE
weights, or tissue cytology, with the possible exception of an increase in
liver fat at the highest dose level. Monkeys maintained on oral doses of 50-
100 mg/kg/day for a year likewise showed no evidence of a toxic effect.
Even the maximum tolerated dose of 200 mg/kg/day, producing some toxic
effects in the monkeys, produced no pathological changes in the tissues. One
must conclude that dehydroacetate is a relatively nontoxic substance. Hu-
man subjects can tolerate 14-17 mg/kg/day for 26-48 days (approximately
1.2 g/day or a total of 30-60 g), maintaining a plasma level of 15-25 mg%
(around 1 mM), and experience only occasional anorexia and nausea. Hu-
man subjects are more sensitive to dehydroacetate than are experimental
animals on a dosage basis, but the plasma concentrations are approximately
the same, indicating that man either metabolizes or excretes dehydroacetate
less readily (see accompanying tabulation).
Tolerated levels
Nontolerated
levels
Species
Dose
(mg/kg)
Days
Plasma
cone.
(mg%)
Dose
(mg/kg)
Days
Plasma
cone.
(mg%)
Rat
50
730
11-
-17
Dog
50
199
12-
13
60-80
73
20-25
Monkey
50
378
12-
-16
100
397
20-30
Man
6-13
173
12-
-17
14-17
26-48
15-25
Urinary Excretion of Succinate
If dehydroacetate is able to inhibit succinate oxidase in the whole animal,
one would expect to find an accumulation of succinate in the body and an
increased excretion. This was indeed found, providing the best evidence
that such enzyme inhibition actually occurs in vivo. Rats given 600 mg/kg
of dehydroacetic acid orally show an increase in urinary succinate during
the first day from a control level of 3.2 mg/day to 24.8 mg/day (Seevers et
al., 1950). Dogs given 200 mg/kg orally for 3 days produce a maximal ex-
cretion of succinate on the second day (more than 175 mg/day compared to
a control of 28 mg/day), and the rise is maintained after cessation of
administration, being around 90 mg/day 4 days after the last dose. One
would also expect other acids to accumulate if the cycle is depressed and
the appearance of ketonemia, but this is not mentioned.
Antidotes
Seevers et al. (1950) attempted to combat the toxic effects of dehydroace-
tate in dogs by the administration of glucose, ammonium lactate, calcium
DISTRIBUTION AND METABOLISM 629
lactate, ammonium chloride, magnesium sulfate, sodium bicarbonate, and
dimercaprol [ ! ], but no protection or benefit was observed, which is not
very surprising. It would be interesting to know if either fumarate or malate,
the products of succinate oxidation and possible restorers of cycle activity,
is effective. Barbital controls the convulsions produced by dehydroacetate
and allows recovery from an otherwise fatal dose, indicating that the con-
vulsions must contribute to the death of the animals.
DISTRIBUTION AND METABOLISM
Whether given orally or parenterally, during acute or chronic administra-
tion, most of the dehydroacetate seems to be metabolized in the body, since
less that 25% is found in the urine (monkeys 10%, dogs 20%, man 22%)
and only around 5% in the feces (Shideman et at., 1950 a,b). Only insignifi-
cant amounts of conjugated dehydroacetate occur in the urine inasmuch as
2-4% more can be obtained on acid hydrolysis. It was not possible to de-
monstrate destruction of dehydroacetate by slices of rat liver, kidney, or
brain, or in muscle mince in incubations up to 4 hr, although, as pointed out,
the analyses may not have been specific enough to have detected certain
chemical modifications. No evidence could be found for the appearance of
2,6-dimethyl-l,4-pyrone or 6-methyl-2^-pyran-2,4(3/f)-dione, substances
formed in the chemical degradation of dehydroacetate. Nor were positive
tests for acetomalonate or acetoacetate, two possible metabolic products,
obtained in dog or human urine. The stimulation of liver and muscle respi-
ration by dehydroacetate might indicate metabolism of dehydroacetate in
these tissues, but this is not at all certain. However, it has been shown that
in animals with carbon tetrachloride liver damage the toxicity of dehydro-
acetate is increased 29%, pointing to the liver as at least one site for the
metabolism.
The oral administration to rats of small doses (60 mg/kg) of dehydroace-
tate, labeled in four of its carbon atoms, leads after 5 days to the following
distribution of the label: urine 23%, feces 19%, carcass 22%, and respira-
tory CO2 12.4% (Barman et al., 1961). The urine contains five labeled sub-
stances: unchanged dehydroacetate (4.7%), hydroxydehydroacetate (7%),
triacetic acid lactone (1.2%), urea (0.2%), and an unknown pyrone meta-
bolite (the figures in parentheses give the percentages of the dose). In ad-
dition, the imino derivatives of dehydroacetate and hydroxydehydroacetate
are found, since reaction with ammonia occurs in the urine. The major
metabolic pathway was postulated to be
Dehydroacetate -► hydroxydehydroacetate -> triacetic acid lactone ->
acetoacetate + acetate
The hydroxylation of the S-COCHg group occurs in slices of liver, but not
630 3. DEHYDROACETATE
in kidney, muscle, or testis (Barman et al., 1963). The rabbit apparently
metabolizes dehydroacetate more readily than the rat. The absence of glu-
curonides or ethereal sulfates in the urine was confirmed. The urinary pyrone
metabolite is probably the 3-carboxylate of triacetic acid lactone.
Dehydroacetate is excreted by the kidney quite slowly. This could be due
to the binding of a major fraction in the plasma to protein, or to active
resorption by the tubules; both actually occur. The fraction bound to plasma
proteins depends on the species and the dehydroacetate concentration
(Woods et al., 1950) (some averages are shown in the accompanying tabu-
lation). However, even when the renal excretion is corrected for the amount
Plasma
Total plasma
Bound de
hydroacetate
Species
protein
dehydroacetate
(%)
(mg%)
0/
/o
mg/g
Rat
5.58
5.7
90.5
0.93
Dog
4.28
4.0
58.5
0.62
Man
5.15
6.2
98.5
1.20
bound, it is evident that this is not the primary factor in the slow excretion.
Dehydroacetate is resorbed through the tubules to about the same extent
as water (98-99%). Since the ring structure of dehydroacetate is identical
to the pyranose form of glucose, it was felt that transport by the glucose
system might occur, but phlorizin, at a concentration that markedly in-
hibits glucose transport, does not alter dehydroacetate resorption. The re-
lationship between the transport of dehydroacetate and the effects of dehy-
droacetate on other transport mechanisms, if any, is not clear.
Both dehydroacetic acid and its sodium salt are absorbed rapidly when
given orally, peak plasma concentrations occurring in 90-120 min. It can
be detected in the blood 3 4 days after single doses, and when administered
chronically many days are required for the plasma level to drop to negli-
gible concentrations. The distribution of dehydroacetate in the tissues (Ta-
ble 3-1) and its variation with time illustrate the complexities of the factors
governing penetration and binding. The low concentration in the central
nervous system and the relatively high level in spinal fluid are surprising.
The biliary circulation of dehydroacetate may play a minor role in retain-
ing it in the body.
Dehydroacetate is secreted in the saliva at a reasonably high concentra-
tion, an injection of 5 mg intraperitoneally in the rat giving salivary levels
of 0.25-0.33 mM at 2-6 hr (Zipkin and McClure, 1958), roughly about half
the plasma concentration. Dehydroacetate has been incorporated into tooth-
pastes to inhibit bacterial growth and caries. However, it has been found
ANTIMICROBIAL ACTIVITY
631
Table 3-1
Distribution of Dehydroacetate in Tissues of the Dog"
Tissue
Dehydroacetate
Intravenously
(160 mg/kg)
Oral]
y (80 mg/kg/day)
At 1 hr
(mg°o)
At 5 hr
(mg%)
for 46 days
(mg%)
25
21
18
15
3.9
2.4
14
11
2.0
11
0.8
3.1
11
0
1.5
9.7
2.4
0.8
6.7
5.5
1.8
6.6
0
0.9
6.6
13
3.1
1.6
0
0
0
—
27
—
19
16
—
4.4
4.5
Blood
Kidney
Intestine
Heart
Spleen
Muscle
Liver
Cerebrum
Lung
Cerebellum -
Bile
Spinal fluid
Colon
medulla
° From Woods et al. (1950).
that dehydroacetate in the diet of rats markedly potentiates the develop-
ment of caries (Zipkin and McClure, 1957, 1958). Dehydroacetate at 0.1%
in a cariogenic diet or drinking water (corresponding to around 5 mg up-
take per day) increases significantly the frequency of caries. It is suspected
that the cariogenic action of dehydroacetate may be related to its secretion
in the saliva, but the mechanism is unknown.
ANTIMICROBIAL ACTIVITY
Dehydroacetate has been used widely the past few years as a food pre-
servative, especially against molds, and is certainly one of the safest and
most effective. This has stimulated extensive work to determine the mini-
mal growth inhibitory concentrations for various microorganisms, some of
the results of which are summarized in Table 3-2. Two things are imme-
diately evident from this table. Dehydroacetate is in general a rather weak
antimicrobial agent; it is of practical value because of its low toxicity. It
632
3. DEHYDROACETATE
Table 3-2
Antimicrobial Activity of Dehydroacetate
Organism
Bacteria
Aerobacter aerogenes
Alcaligenes faecalis
Bacillus anthracis
B. cereus
B. megaterium
B. mesentericus
B. suhtilis
Corynebacterium diphtheriae
Escherichia coli
Lactobacillus acidophilus
L. brevis
L. casei
L. fermenti
L. plantarum
Mycobacterium tuberculosis
Pseudomonas aeruginosa
Salmonella pullorum
Salmonella typhosa
Staphylococcus aureus
Vibrio cholerae
V. metchnikowii
Fungi
Aspergillus niger
Botrytis allii
Fusarium graminearum,
Penicillium digitalum
P. expansum
Rhizopus nigricans
Trichophyton interdigitale
T. mentagrophytes
Yeasts
Candida albicans
Saccharomyces cerevisiae
Minimal inhibitory
concentration
Reference "
(mM)
17.8
(6,7)
23.8
(6)
59.5
(1)
17.8
(6)
17.8
(6)
17.8
(6)
17.8
(6)
3.0
(1)
23.8
(6)
59.5
(2)
12-60
(2)
12-24
(2)
12-60
(2)
12
(2)
5.9
(6)
5.9
(5)
23.8
(6)
17.8
(6)
12
(6)
108
(4)
10
(5)
17.8
(7)
59.5
(1)
5.9
(1)
3.0
(6)
0.3
(3)
0.47
(3)
2.4
(3)
1.8
(6,7)
0.6
(6)
2.4
(6,7)
0.3
(6)
0.59
(4)
0.3
(6)
1.9
(4)
5.9
(6,7)
" References: (1) Brodersen and Kjaer (1946); (2) Fitzgerald and Jordan (1953);
(3) McGowan et al. (1948); (4) Stedman et al. (1954); (5) Ukita etal. (1949); (6) Wolf
(1950); (7) Wolf and Westveer (1950).
ANTIMICROBIAL ACTIVITY 633
is clear that fungi are more sensitive to dehydroacetate than are bacteria.
The means of the concentrations in the table are, of course, not quantita-
tively significant, but show well the difference: 26 mM for bacteria and
1.1 mM for fungi. The mechanism of growth inhibition is completely un-
known. There is no obvious correlation between cycle activity in these organ-
isms and susceptibility to dehydroacetate. The fungi behave differently than
bacteria with regard to so many drugs that one must assume basic differ-
ences in metabolism or permeabilities, and it would be impossible at this time
to attribute the greater sensitivity to dehydroacetate to any one factor.
Permeability seems to be of importance in the action of dehydroacetate,
as indicated by the effects of pH. A decrease in activity with increasing pH
has been generally noted (Shibasaki and Terui, 1953; Bandelin, 1958), with
the exception of Salmonella and Staphylococcus (Wolf and Westveer, 1950).
The results of Bandelin on several fungi are typical (see accompanying ta-
bulation). A 100- to 200-fold increase in activity as the pH is raised from
Minimal inhibitory concentration (mM)
Organism
pH 3
pH 5
pH 7
pH 9
AUernaria solani
0.059
0.12
1.18
11.8
Aspergillus niger
0.12
0.30
2.36
11.8
Chaetowium globosum
0.059
0.30
2.36
5.9
Penicillium citrinum
0.059
0.30
2.36
11.8
3 to 9 is observed. The only obvious explanation is that the anionic forms
of dehydroacetate do not penetrate well. The major decrease in activity
occurs between pH 5 and 7, correlating with the pK^'s near 5.2,
CHAPTER 4
SULFHYDRYL REAGENTS
A substance which can react with sulfhydryl groups and thus alter en-
zymic, metabolic, or functional processes is generally called a sulfhydryl
reagent. Such substances represent a very important group of inhibitors
and have been used extensively to determine if enzymes or metabolic reac-
tions depend in any way on intact sulfhydryl groups. In addition, they are
often used to estimate the number and reactivity of sulfhydryl groups on
proteins, or to histochemically localize the sulfhydryl groups in cells or tis-
sues. The next few chapters will be concerned with sulfhydryl reagents, and
in this chapter we shall discuss several general aspects of inhibition result-
ing from modifications of sulfhydryl groups and some of the problems en-
countered in work with these substances. This is one phase of inhibition
that has recently received considerable attention, and several reviews cover-
ing certain aspects of the problem are available. The articles by Boyer (1959)
and Putnam (1953), and the books "Glutathione" (1954) and "Sulfur in
Proteins" (1959), are particularly recommended.
The terminology to be adopted attempts to follow the most recent usage.
The sulfhydryl group (= mercapto group) will for brevity be designated
as an SH group. Compounds containing SH groups will be designated as
thiols (elsewhere occasionally called sulfhydryl compounds or mercaptans).
A sulfhydryl reagent wiU be termed an SH reagent. The designation as a
sulfhydryl enzyme has often been meant to imply that the catalytic activity
of the enzyme is dependent on SH groups, i.e., that the SH groups actually
participate in the enzyme reaction. As Boyer (1959) has pointed out, not
a single enzyme has been definitely shown to involve protein SH groups
in the catalysis, and we shall see that the inhibition of an enzyme by an
SH reagent does not prove that the SH groups are functional. Hence, a
more practical definition of a sulfhydryl enzyme at the present time is an
enzyme that shows a loss of activity when some or all of its SH groups
are modified.
635
636 4. SULFHYDRYL REAGENTS
ROLE OF SH GROUPS IN METABOLISM AND FUNCTION
Cellular components containing SH groups may be conveniently grouped
in three categories: (1) low molecular weight thiols, such as the cofactors li-
poate, coenzyme A, and glutathione, or various amino acids and related
compounds, such as cysteine, homocysteine, 2-thiolliistidine, ergothioneine,
and thioglycolate, (2) nonenzyme proteins, probably including most of the
cytoplasmic proteins (e.g., those involved in movement, such as actomyosin,
ciliary proteins, and proteins of the mitotic spindle), plasma membrane pro-
teins, and structural proteins, and (3) enzymes of all types and catalyzing
every variety of reaction. Modification of or reaction with any of these SH
groups may directly or indirectly alter cellular metabolism and function.
Even reaction with nonenzyme protein SH groups may disturb metabolism,
because of the role such proteins may play in the structural organization of
the metabolic units or in the permeabilities of cells. In addition to the free
SH groups, many proteins and enzymes contain disulfide (S — S) groups
that, in the case of enzymes, are probably not involved directly in the ca-
talysis but in the structural stability. These disulfide groups can under cer-
tain circumstances be reductively cleaved to form free SH groups, with
simultaneous loosening of the protein structure, or can perhaps react di-
rectly with certain agents to form mercaptides.
In the early days of interest in thiols, it was believed that SH reagents
altered metabolism, and were sometimes lethal, as a result of reaction with
glutathione or other low molecular weight thiols, but it was soon realized
that enzyme SH groups are a much more important site of attack. Even
today the role that such small thiols play in metabolism and the importance
of their reaction with SH reagents are not well understood, except in the
case of coenzyme A and lipoate. The ubiquitous glutathione plays at pres-
ent an indeterminate role in metabolism, except for its likely participation
in the reactions of phosphoglyceraldehyde dehydrogenase, glyoxalase, mal-
eate isomerase, maleylacetoacetate isomerase, formaldehyde dehydrogenase,
and indolylpyruvate tautomerase, and in transpeptidation and folate split-
ting. Low molecular weight thiols have also been supposed to regulate me-
tabolism by redox equilibria with enzyme SH groups, maintaining a cer-
tain fraction of these in the reduced or active state.
The SH groups of enzymes have been considered to bind cofactors or
coenzymes to the apoenzyme, or to form acyl or phosphoryl complexes
with intermediates derived from substrates, or to function directly as redox
couples in electron transfer, but there is little evidence for any of these, as
likely as they may be. The SH group readily donates electron pairs and
thus is one of the most reactive enzyme groups with regard to the formation
of covalent bonds, so it would not be surprising if covalent intermediate
complexes occur. Whatever the role SH groups play in enzyme catalysis,
CHEMICAL PROPERTIES OF SH GROUPS 637
their modification often abolishes activity and, since metabolism depends
on sulfhydryl enzymes, it is evident that most important metabolic path-
ways would be sensitive to SH reagents. In addition, coenzyme A, lipoate,
and glutathione function in key metabolic positions. Thus glycolysis, the
tricarboxylate cycle, fatty acid oxidation, photosynthesis, phosphate trans-
fer, and various synthetic pathways are inhibitable by SH reagents. Many
effects of thiols on metabolism have been observed but no detailed mechan-
isms emerge. Brain respiration and glycolysis in vivo proceed at only a frac-
tion of their maximal rates; it has long been known that glutathione stim-
ulates aerobic glycolysis in the brain, and thus it has been implicated in
the regulation of cerebral metabolism. Mcllwain (1959) reported that the
aerobic glycolysis of brain slices is stimulated by glutathione, cysteine,
homocysteine, 2-mercaptoethanol, and other thiols, although the effects on
respiration are rather slight. However, the respiratory stimulation by 50 m.M
KCl is depressed by these thiols, as is the excess respiration in the presence
of dinitrophei|ol. The glycolytic stimulation is accompanied by a decrease
in creatine-P and a rise in inorganic phosphate, these changes being corre-
lated with the metabolic changes. If the respiratory augmentation produced
by increased functional activity were mediated through glutathione or
similar thiols, there would have to be a fairly large change in their concen-
trations, or in the ratios of the oxidized and reduced forms, which is not
observed. The metabolic relations are clear but it is not known by what
mechanism the thiols reduce brain creatine-P.
Studies of the effects of SH reagents on cell function are complicated by
the fact that undoubtedly some of the proteins of the functional systems
contain SH groups, and may even be dependent upon them. This has been
investigated principally in the proteins involved in motility; for example,
the polymerization of G- to F-actin, the interaction of actin and myosin,
the ATP-induced contractions of glycerinated flagella, the round-up of cul-
tured fibroblasts, the formation of the mitotic apparatus, and many other
phenomena appear to be dependent on free SH groups. Cell excitability
and impulse conduction, based on ionic fluxes and a specific membrane
^ructure, must also involve SH groups in the membrane. Thus effects of
SH reagents on cell function cannot be immediately interpreted in terms
of an enzymic or metabolic site of action.
CHEMICAL PROPERTIES OF SH GROUPS
Only a few characteristics of the SH group that are particularly important
in enzyme inhibition will be discussed. A brief and excellent summary of
sulfur chemistry is that of Calvin (1954) and much of interest may be found
in "Organic Sulfur Compounds" edited by Kharasch (1961), as well as in
the general references given earlier in this chapter.
638 4. SULFHYDRYL REAGENTS
The reactions of most SH reagents with thiols depend on the pH and this
undoubtedly relates to the ionizations of both SH reagent and the SH groups
reacted. In most cases, as with the mercaptide-forming reagents and the al-
kylating agents, the rate and degree of reaction increase with increasing
pH, and it is likely in these cases that the ionized mercapto anion, R — S",
is more reactive than the un-ionized R — SH form. One may visualize some
of these reactions as a competition between the SH reagent and a proton
for the R — S~ group. The reaction with a heavy metal ion, for example,
may be written as:
R— SH + Me+ ^ R— S— Me + H+
and it is obvious that increase of the pH will favor the formation of the
mercaptide complex, or, to put it in another way, that the reaction:
R— S- + Me+ ^ R— S— Me
will proceed more readily. On the other hand, reactions with double bonds
(as with maleate or quinones) or oxidations to the disulfide may occur more
readily when the SH group is un-ionized. In any event, the state of ioniza-
tion of the SH group is important in enzyme inhibition and may account
partly for the different reactivities of protein SH groups. The ionization of
SH groups is well discussed by Edsall and Wyman (1958), mainly on the
basis of work by Benesch and Benesch (1955). The ionization microcon-
stants for cysteine are given in Table 1-14-4. It is clear that the pK^ of an
SH group is markedly dependent on the electric field present, that is, on
the vicinal ionic groups. One might roughly estimate the p^,/s for an SH
group as shown in the following tabulation:
pK„
Near a + charged group 7.2-8.5
No electric field 8.5-9.2
Near a — charged group 9.2-10.2
On the surface of a protein the electric field will be the resultant of all the
contributions of the ionic groups. Enzyme SH groups must therefore exhibit
a wide range of ionizations at any designated pH, and in most cases will
exist mainly in the un-ionized form at physiological pH. The piii^ of the SH
groups on aldolase in 4 M urea is around 8.66, but the native enzyme has
6 exposed SH groups with \)K„ values near 10.5 and buried SH groups with
an apparent p^„ of 11.5 (Donovan, 1964).
CHEMICAL PROPERTIES OF SH GROUPS 639
Many of the atomic and bond properties will be found in the tables of
Chapter 1-6. The bond dipole moments are fairly high (C — S 1.73 and S — H
0.68, corresponding to fractional atomic charges of 0.20 and 0.11, respec-
tively) and the bonds with sulfur are readily polarized (the molar refrac-
tions are C — S 4.43, S — H 4.62, and S — S 7.41) compared with most other
bonds occurring in proteins. Furthermore, the bond energies are uniformly
low compared with the corresponding oxygen bonds, except for the disulfide
bond, which is a good deal stronger than the peroxide bond (see accompa-
nying tabulation). These fundamental properties account for many of the
Bond energies (kcal/mole)
C— S 54
C— 0 80
S— H 87
0— H 105
S— S 66
0—0 34
characteristic reactions of the SH group and the relatively unique role of
these groups in enzyme activity and inhibitition. The inherent dipole mo-
ments and the high polarizability of sulfur bonds may play an important
role in the interactions of enzymes with substrates and inhibitors, whereas
the bond energies are involved in determining ionization tendencies, oxida-
tion-reduction potentials, and the equilibria between SH groups and disul-
fide structures.
The thiol-disulfide equilibria are important for enzyme structure in all
probability but, in addition, may well be determining factors in the states
and reactivities of the SH groups. It has been shown recently that the reac-
tion of a thiol with a disulfide is not a simple oxidation-reduction but an
exchange reaction involving a two step ionic displacement (Eldjarn and
Pihl, 1957 a, b; Parker and Kharasch, 1959; Foss, 1961), often with the
formation of mixed disulfides:
X— SH + Y— S— S— Y ±^ X— S— S— Y + Y— SH
X— S— S— Y -f X— SH ±? X— S— S— X + Y— SH
Low molecular weight thiols, such as glutathione or cysteine, could thus
interact with enzyme SH and disulfide groups to form mixed disulfides. Par-
ticularly in the cell, where such thiols occur, these interactions may be im-
portant in regulating enzyme activity, and could easily affect the reactivity
of enzymes with SH reagents. That this can actually occur with proteins
was shown by the use of a colored disulfide, with which seralbumin and /?-
lactoglobulin react to form mixed disulfides (Klotz et at., 1958). If an SH
enzyme and oxidized glutathione (GSSG) are allowed to react, one would
640 4. SULFHYDRYL REAGENTS
have ESH, ESSG, ESSE, GSSG, and GSH (where E represents the enzyme)
present, perhaps only ESH being catalytically active and the forms ESSG
and ESSE protected from SH reagents. Although it is usually thought that
only the SH form can react with most SH reagents, it is possible that di-
sulfides are occasionally reactive; aryl arsinites, for example, can exert a
nucleophilic displacement on the disulfide bond:
OH
RSSR + R'AsO(OH)- ±^ RSAsR' + RS~
O
However, such reactions are probably slower than with free SH groups.
The new reagent, dithiothreitol (HS— CH2— CHOH— CHOH— CHg— SH),
which has a low redox potential ( — 0.33 v at pH 7), is highly water-soluble
and reduces protein disulfide groups (Cleland, 1964). It was suggested that
it might be valuable in protecting enzyme SH groups, having several ad-
vantages over the ones commonly used, and could also be applied for the
purpose of maintaining enzymes in the SH state for inhibition studies.
Hydrogen bonding by sulfur should be mentioned since it must play a
role in both intra- and intermolecular interactions of the SH group, but
most of the data we have derives from studies of the small thiols and there
is very little information on hydrogen bonding of protein SH groups. Boyer
(1959) has presented the evidence for the occurrence of hydrogen bonds to
sulfur in a variety of compounds. Sulfur does not form hydrogen bonds as
readily as oxygen or nitrogen, since it is less electronegative (as indicated
by the smaller dipole moment of the S — H bond compared to the 0 — H and
N — H bonds) (Table 1-6-1). However, there is evidence for intramolecular
S — H • • • 0 and S — H • • • N bonds in cysteine and its peptides, and Benesch
et al. (1954) have advanced hydrogen bonding to explain some of the dif-
ferent reactivities of simple thiols. It is possible that hydrogen bonding of
enzyme SH groups can modify their susceptibilities to various SH reagents.
Detection and Determination of Enzyme SH Groups
Valuable reviews of the general methods for the determination of SH
groups in proteins and enzymes have been provided by HeUerman and
Chinard (1955) and R. Benesch and R. E. Benesch (1962). Some of the
most reliable methods involve the use of mercurials (to be discussed later,
pages 762 and 766). Here we shall mention only a few of the more recently
developed reagents which may be applicable in inhibition studies.
Bis(2?-nitrophenyl)disulfide reacts with thiols at pH 8 to form 1 mole of
p-nitrophenol per mole of thiol, and this anion, being highly colored, can
be used to determine the thiol concentration (Ellman, 1959). However, this
CHEMICAL PROPERTIES OF SH GROUPS 641
reagent is poorly soluble in water, so carboxyl groups were introduced to
solubilize it; this compound is 5,5'-ditliiobis(2-nitrobenzoate) and reacts
0,N {' V — S— S — V ) — NO;
5, 5'-Dithiobis (2-nitrobenzoate)
with thiols and SH groups in the blood and tissues. Another water-soluble
reagent for free SH groups is 2,2'-dicarboxy-4,4'-diiodoaminoazobenzene,
which was shown to react only with the SH groups on denatured meromyosin
(Fasold et al., 1964). The number of SH groups reacted can be determined
coo'
I— CHi— CONH-Y^ \^N=N^/ ^V-NHCO— CH2— I
"ooc
2, 2'-Dicarboxy-4, 4'-diiodoaminoazobenzene
spectrophotometrically because of the chromogenic azo link. A yellow SH
reagent, A^-(4-dimethylamino-3,5-dinitrophenyl)maleimide, was introduced
by Witter and Tuppy (1960) and found to react with the free SH groups
of seralbumin. The treated protein could be hydrolyzed with pepsin and
the A'-(4-dimethylamino-3,5-dinitrophenyl)succinimido-cysteine peptides
isolated by means of their yeUow color. This reagent was used by Gold and
Segal (1964) to obtain information on the nature of the active site of 3-
phosphoglyceraldehyde dehydrogenase. Following pepsin treatment the sin-
gle hexapeptide - Ala-Ser-(DDPS-Cys)-Thr-Thr-AspNH2 - w^as found to
contain essentially aU the color. This provides evidence that the three active
sites on the enzyme are similar in structure, at least in part, and that the
reactive SH groups are not those of glutathione, which occurs on the enzyme.
O.N
0,N
A/-(4-Dimethylamino-3, 5-
dinitrophenyl) maleimide
Such reagents would not be particularly useful for the inhibition of SH en-
zymes because of their bulky structure and the presence of a variety of
642 4. SULFHYDRYL REAGENTS
groups, but they could be applied to the determination of changes in the
SH content after treatment of the enzymes with the usual inhibiting re-
agents.
TYPES OF SH REACTION IMPORTANT IN INHIBITION
The reactions of most of the important SH reagents have usually been
classified into four types. The SH groups have been written as un-ionized
in all cases, not implying that this is necessarily the only reactive form.
The mechanisms of these reactions will be discussed in the chapters devot-
ed to the individual inhibitors.
(I) Oxidation of SH growps
2R-SH ^ X •* > R-S-S-^R ^ XH,
R = (SH), + X •* » R^l + XH.
S
(X may be an acceptor of either hydrogen atoms or electrons.)
Examples: o-iodosobenzoate, porphyrindin, porphyrexide, iodine, alloxan
(not the only mechanism), ferricyanide, oxidized glutathione, tetrathionate,
sulfite, performic acid, and oxygen (catalyzed by metal ions).
(II) Mercaptide formation
R-SH * X" ^ jT R-S-X - H
2R— SH + X^ • -^ ^ R-S— X-S— R ■ 2H
. /S^
R=(SH), + X ' "* >: R^ X + 2H
Examples: HgClg, organic mercurials, arsenite, organic arsenicals, and
various heavy metal ions (Cu++, Pb++, Cd++, Ag+, etc.).
(III) Alkylation of SH groups [alkyl transfer)
R— SH + X— R' ±? R— S— R' + X
Examples: iodoacetate, iodoacetamide, S- and N-mustards, chloraceto-
phenone, chloropicrin, bromobenzylcyanide, and fluoropjTuvate.
FACTORS •DETERMINING THE REACTIVITIES OF SH GROUPS 643
(IV) Addition of SH groups to double bonds
CH— R' R— S— CH— R'
R— SH +11 ±? I
CH— R" CH2— R"
OH
R— SH + 0 = C— R' ±^ R— S— C— R'
(This may also be considered as a type of alkylation reaction.)
Examples: maleate, iV-ethylmaleimide, quinones, acrolein, acetoacetate,
and methylglyoxal.
FACTORS DETERMINING THE REACTIVITIES OF SH GROUPS
The SH groups of various simple thiols, peptides, and proteins differ mark-
edly in reactivity with SH reagents. Although this has been known for many
years, the molecular basis for this differential reactivity is poorly understood.
In general the reactivity is maximal in simple thiols and minimal in proteins,
but in proteins there are usually reactive and unreactive SH groups. Barron
(1951) classified SH groups as (a) freely reacting, (b) sluggish, and (c) mask-
ed, depending on whether they react readily, slowly, or not at all. Although
such a division is often useful in discussing SH groups, there is actually a
continuous sequence of groups from highly reactive to unreactive. If a pro-
tein is allowed to react with an SH reagent under approximately physiolo-
gical conditions, one generally finds that the SH groups disappear at differ-
ent rates, perhaps several reacting completely before others are affected. A
graded response is clearly seen in the reaction of aldolase with p-chloromer-
curibenzoate (Swenson and Boyer, 1957). Ten SH groups react relatively
rapidly, a few more slowly, and the rest not at all unless the enzyme is un-
folded by high concentrations of urea. Furthermore, the reaction of the first
10 SH groups does not alter the enzyme activity, indicating that these
groups are not part of, or even too near, the active center, whereas disap-
pearance of the more slowly reacting groups abolishes the activity. A similar
situation has been observed with urease, which has 5 cysteine residues per
molecule, one freely reacting and 4 sluggish, the catalytic activity being
affected only by modification of the latter (Hellerman, 1939; Hellerman et
al., 1943). Thus porphyrindin, iodoacetamide, and iodosobenzoate react with
one SH group but do not inhibit (except at very high concentrations),
whereas p-chloromercuribenzoate can combine with another SH group abol-
ishing the activity. These examples — and we shall have occasion to discuss
many others — illustrate four most important principles: (1) the differential
reactivity of enzyme SH groups, (2) the increase in reactivity of many of
the SH groups following denaturation, (3) the different reactivities of var-
644 4. SULFHYDRYL REAGENTS •
ious SH reagents, and (4) the lack of correlation between the reactivity of
SH groups and their relationship functionally or spatially to the active
center. These principles are central to the problem of inhibition by SH re-
agents and require some general discussion relative to the possible mecha-
nisms involved.
The various, and mostly obvious, hypotheses to explain the differential
reactivity of SH groups have frequently been presented with a prolixity
inversely proportional to the amount of evidence available. Indeed, at the
present time there is little, if any, positive evidence for any explanation,
but there are a number of factors that must be of some importance, and
these can be enumerated. It should be emphasized that differential reactivity
should be based on accurate spectrophotometric or argentimetric titrations
of the enzyme SH groups under various conditions. The fundamental prob-
lem is to determine the cause for the slow reactions of all or a fraction of
an enzyme's SH groups, the total number of such groups being determined
by quantitative titrations of the enzyme after complete unfolding. The
theories assume either that (I) free SH groups are present in the native cat-
alytically active enzyme, but are for some reason unable to react readily
with SH reagents, or that (II) the unreactive SH groups are so modified
that they are no longer free.
(I) Free SH groups present in native enzyme
A. Steric factors impede reaction: the reagent is simply unable to approach
the SH group because it is located in a pit or crevice of the enzyme, or ac-
tually within the protein structure.
B. Electrostatic factors impede reaction: the SH group is in the electric
field of surrounding groups, this discouraging reactions with reagents of
the same charge sign as these groups.
C. Ionization state impedes reaction: if either the SH or the S~ form reacts
preferentially but is not significantly present at the experimental pH, the
reaction will be appreciably slowed.
(II) SH groups are not free in the native enzyme
A. Present as disulfide groups
B. Hydrogen bonded to adjacent groups
C. Present in thiazolidine or thiazoline rings
D. Reacted with some component of enzyme reaction: for example, acylated,
phosphorylated, or complexed with some metal ion.
The fact that complete opening up or unfolding of the protein structure
invariably increases the susceptibility of certain SH groups to attack does
not necessarily imply that the groups are in some way within the enzyme,
FACTORS DETERMINING THE REACTIVITIES OF SH GROUPS 645
since denaturation can also dissolve disulfide bonds, hydrogen bonds, ring
structures, and other possible chemical interactions of the SH groups. As
long as one studies only a single SH reagent, it is easy to postulate a reason-
able mechanism for the unreactivity of particular SH groups. For example,
if one finds that iodoacetate does not alkylate an enzyme SH group, the
group may be thought of as sequestered within the protein structure, but
if subsequent work shows that p-chloromercuribenzoate reacts readily with
this group, this hypothesis must be abandoned inasmuch as j^-chloromer-
curibenzoate is a larger molecule than iodoacetate. Likewise, postulating
that negative charges surround the SH group, preventing the approach of
iodoacetate, will not be valid if p-chloromercuribenzoate is effective, since
both of these reagents are negatively charged, as has been pointed out by
Boyer (1959) in perhaps the best discussion of differential SH group reac-
tivity. It is certainly likely that steric and electrostatic factors are occa-
sionally important, but one must demonstrate some correlation of the un-
reactivity of the SH groups with the properties of a variety of SH reagents.
Haurowitz and Tekman (1947) believed that protein SH groups are often
inaccessible to reagents because of the tightly folded nature of the polypep-
tide chains, rather than chemically combined, because unfolding is accom-
panied by the appearance of reactivity in other than SH groups, e.g. phe-
nolic groups. Although this is suggestive, it is not proof for the inaccessibi-
lity theory. Unreactivity due to an unfavorable ionization state has per-
haps been insufficiently considered, particularly for inhibitors such as iodo-
acetate, and there is no question but that the very low concentration of S~
near neutrality for some SH groups must be important.
Turning to the second group of theories, no one denies that disulfide
groups occur in some enzymes, but that this generally cannot explain the
differential reactivity of SH groups is obvious. Indeed, one finds a wide
range of reactivities in simple thiols where cryptic exclusion or disulfide
bonding may be eliminated. Benesch et al. (1954) not only demonstrated
different nitroprusside reaction rates with various biologically important
thiols, but showed that urea increases the reactivity of the more slowly
reacting SH groups, just as it does in proteins. This was interpreted in
terms of the breaking of hydrogen bonds and thus the initial sluggishness
of reaction as due to hydrogen bonding of the SH groups to adjacent amino
or peptide groups. This may occur in a cysteine peptide in the following
way:
H,C "H H,C H
^11 "I i
R,— CONH-HC— CO— N— R, R— CONH— HC— CO— NH— R,
the hydrogen donator depending on the pH. We have seen that SH groups
form only weak hydrogen bonds (page 640), so that it is likely that this
646 4. SULFHYDRYL REAGENTS
alone cannot depress the reactivity too greatly. However, in addition the
hydrogen bond may bring an adjacent side chain into the region of the SH
group and this second steric factor may further reduce the reactivity. Ben-
esch and Benesch (1953) compared the peptides, phenacetyl-L-cysteinyl-
glycine (PCG) and phenacetyl-L-cysteinyl-D-valine (PCV), with respect to
the polarographic reduction of their mercaptides with mersalyl and Ag+,
and found that a relative suppression of the ability of the PCV SH group
to react with these reagents is evident. This was interpreted as due to the
steric interference of the isopropyl group of valine in PCV, brought into
the proximity of the SH group by hydrogen bonding, whereas in PCG there
is no such side chain. In proteins the interference may be even greater.
This concept thus involves both a reaction of the SH group and steric
factors.
The hydrogen-bonded structures could further lose water to form more
stable thiazolidine or thiazoline rings. Linderstrom-Lang and Jacobsen
(1941) found that 2-methylthiazoline can hydrolyze under certain con-
ditions to release an SH group and a peptide linkage, such as occurs
during protein denaturation. Thus in a cysteine peptide, where the hy-
drogen bonding is now to the keto oxygen, the following structures can
be written:
S— H
/ \
H,C O HX S
Ri — HC— NH— C— R, R— HC— NH— C — Rs
OH
H-bonded form Thiazolidine form
H,C S
"I I
R — HC— N=C-R2
Thiazoline form
Such transformations have been more recently discussed by Calvin (1954)
in connection with the structure of glutathione and the reactivities of SH
groups in enzymes. Indirect evidence for these rings was obtained by com-
paring the absorption spectra of thiols with 2-methylthiazoline (although
at high acidity so that the situation near neutrality is still not clear). If
such structures occur in enzymes, they could unquestionably account for
unreactivity.
INTERPRETATION OF INHIBITIONS BY SH REAGENTS 647
It is quite possible that many or all of these mechanisms contribute in
various situations to the differential reactivity of SH groups, and that we
should not be too eager to argue for a single dominant factor. After all,
there is a graded scale of reactivity, which in itself implies multiple mecha-
nisms. It would probably aid in the characterization of SH groups if some
standard method for designating the reactivity could be used, rather than
designating them by terms such as "sluggish," etc. The time for 50% reac-
tion, where determinable, might be the simplest and most useful, although
some form of reaction rate constant would be preferable.
INTERPRETATION OF INHIBITIONS BY SH REAGENTS
The SH reagents are used most commonly to determine whether a parti-
cular enzyme is an "SH enzyme" or not. What this means depends on one's
definition of "SH enzyme." If we take the definition proposed earlier (page
635) that an SH enzyme is one that is inhibited by SH reagents, not a great
deal has been achieved by proving that an enzyme belongs to this class.
In the past, many workers have been satisfied to stop at this distinction,
and the designation of an enzyme as an SH enzyme has been deemed suffi-
cient without further discussion as to the significance of the observation.
On the other hand, some have assumed immediately that inhibition by SH
reagents indicates a catalytically functional role for an SH group at the
active center, and this, as we have seen, is entirely unjustifiable. It is thus
important to determine as far as possible what such inhibition means and
what valid conclusions may be drawn from the use of SH reagents.
The various mechanisms by which SH reagents can inhibit pure enzymes
may be classified in the following way.
(A) The SH group reacted is at the active center and is functional. The SH
group may be involved in the binding of substrate, coenzyme, or activator
to the apoenzyme, or it may participate in the transfer of groups of elec-
trons.
(B) The SH group reacted is at the active center but is nonfunctional. It is
possible that an SH group occurs at the active center but is unrelated to
the catalytic process.
(C) The SH group reacted is vicinal to the active center. The SH reagent
introduces a new structure on the enzyme, and if this is near enough to
the active center it may either sterically or electrostatically modify the
reactions proceeding at the active center.
(D) Reaction of the SH groups alters the enzyme protein structure. This
could also apply to reaction with disulfide groups or other complexes form-
ed by SH groups. The change in protein structure would then reduce the
648 4. SULFHYDRYL REAGENTS
rate of the enzyme reaction, particularly if it included the region of the
active center.
(E) Tlie SH group reacted is on the substrate. This is a possibility especially
in the case of proteolytic enzymes, the modification of the peptide or protein
substrate preventing normal reaction with the enzyme.
(F) The SH reagent interferes in a manner unrelated to SH groups. Many
SH reagents are not entirely specific for SH groups; e.g., iodoacetate also
reacts with amino groups and with heavy metal ions such as Cu++, and can
often form complexes with protein groups other than SH, especially amino
and carboxylate groups. Also the SH reagent may inhibit because it is
structurally similar to the substrate and can compete with it for the active
site; e.g., p-chloromercuribenzoate may act like a substituted benzoate on
certain enzymes rather than as a mercurial.
Other mechanisms can be visualized in special cases and particularly for
those enzymes comprising several units and catalyzing complex reactions,
since the SH reagents can conceivably dissociate the functionally related
units, just as /j-chloromercuribenzoate can split the relatively simple muscle
phosphorylase a into four equivalent fractions (Madsen and Cori, 1956).
It is very difficult to distinguish between the first four possibilities. In-
deed, proof of the functional role of SH groups usually must come from
evidence other than inhibition. Protection of the enzyme against SH re-
agents by the substrate does not provide adequate evidence that the react-
ed SH group is part of the active center, since the substrate could also slow
down or prevent reaction with vicinal groups as well, and could also sta-
bilize the protein structure around the active center. The secondary alter-
ation of protein structure brought about by reaction of SH groups cannot
always be detected by reversal experiments because the changes may, like
certain types of denaturation, be reversible. One must therefore conclude
that the demonstration of inhibition by SH reagents indicates at best (as-
suming the mechanisms (E) and (F) have been eliminated) only that one
or more SH groups are sufficiently near the active center to interfere with
the catalysis, either directly or by structural changes, when they are react-
ed. It must be admitted that such a conclusion is not very informative,
especially when it is considered that most enzymes contain 5 to 30 SH
groups per molecule and that statistically one would expect one or more
of these to be near the active center. Indeed, it is rather surprising that in
some instances a fair number of SH groups can be reacted without altering
the catalytic activity.
One characteristic of inhibition by SH reagents which has been often
neglected is that the reaction with the enzyme SH groups in most cases
introduces a new side chain onto the protein. The inhibition may be as
much related, if not more, to the properties of this side chain as to the
INTERPRETATION OF INHIBITIONS BY SH REAGENTS 649
disappearance of a free SH group. These new groupings have varying sizes
and frequently electrical fields. It is quite possible that, all else being equal,
a smaller reagent of a particular type may exert less inhibition, due simply
to the fact that it exerts less steric hindrance to the catalytic process. Thus
the inhibition by methylmercuric chloride may be different from that by
p-chloromercuribenzoate for this reason. The groups introduced by iodo-
acetate and p-chloromercuribenzoate are negatively charged, whereas those
from iodoacetamide and phenylmercuric acetate are uncharged, and this
could well be responsible for some of the differences observed between these
inhibitors. This is one reason why many studies with SH reagents would
profit from a quantitative comparison of the effects of a large number of
inhibitors of different types.
Another factor of some importance may be the influence of the reaction
of certain SH groups on the reactions of other SH groups. Further reaction
apparently may be either depressed or accelerated. In phosphorylase a the
reaction of the first SH group seems to facilitate the combination of the
remaining 18 groups with p-chloromercuribenzoate (Madsen and Gurd, 1956).
On the other hand, reaction of one SH group on hemoglobin prevents the
further reaction of one or two other groups with Ag+, implying that the SH
groups here occur in clusters (Ingram, 1955). The number of molecules of
SH reagent bound to the protein may thus not be equivalent to the number
of SH groups.
Boyer (1959) has emphasized that insufficient consideration has usually
been given to the possible secondary structural changes induced in enzymes
by reaction with SH reagents. If some SH groups are unreactive because
of steric blocking or chemical combination, these hindrances must be over-
come in order to react these groups, and this could imply a modification
in the protein structure that in itself might be inhibitory. It has been ob-
served frequently that the poorly reacting SH groups are more important
in the enzyme structure than the free readily reactive ones. One of the first
statements of the importance of structure in the inhibition by SH reagents
resulted from work on urease by Desnuelle and Rovery (1949). Phenyliso-
cyanate reacts with certain SH groups rapidly but this does not inhibit;
inhibition begins when the unreactive SH groups are attacked, and this
was attributed to a reversible change in the enzyme structure. Similarly,
various properties of aldolase change as the SH groups are progressively
reacted with p-chloromercuribenzoate: after 3-5 are reacted, the enzyme
begins to be more readily attacked by trypsin; after 10 are reacted, the tur-
bidity increases, denoting marked structural changes, and inhibition is ob-
served (Szabolcsi and Biszku, 1961). There is a progressive labilization of
the tertiary structure, accompanied by appearance of previously masked
SH groups, with further reaction and eventual denaturation. In many cases,
the initial reactions must reduce the protein stability, perhaps only locally.
650 4. SULFHYDKYL REAGENTS
and this spreads and progresses rapidly as further groups are attacked, just
as in other types of denaturation. The blocking of the SH groups of phos-
phoglyceraldehyde dehydrogenase changes the optical rotation and the in-
trinsic viscosity, the latter increasing linearly with the equivalents of p-
chloromercuribenzoate reacted (Elodi, 1960). Reversibility with cysteine
varies with the time of exposure to the mercurial, at first the effects being
completely reversible and eventually irreversible, again indicating a pro-
gressive breakdown of the protein structure. Elodi postulated three phases:
(1) a reversible reaction with certain SH groups, (2) an unfolding of the
polypeptide helices as a result of the alteration of the SH groups, and (3)
precipitation due to intermolecular bridges formed between the new groups
appearing on the protein surface. Ribonuclease is perhaps another example
of structural changes resulting from the scission of disulfide bonds, of which
there are 4 in the native enzyme: breaking 1 does not alter the activity,
breaking 2 inhibits about 20%, breaking 3 inhibits about 40%, and then
suddenly the activity drops to zero as the last disulfide is split (Resnick
et al., 1959). The — S — S — bonds were believed to be of importance in pro-
viding stability to the secondary structure of the enzyme, their breaking
leading to progressive unfolding.
PROTECTION AND INHIBITION REVERSAL BY THIOLS
Some of the problems involved in protection and reversal experiments
with SH reagents and thiols were discussed at some length in Volume I
(pages 622-626). A few of the conclusions reached there will be briefly sum-
marized. (1) Protection or reversal by a thiol depends on the relative affin-
ities of the SH reagent for the enzyme SH groups and the thiol, and the
relative concentrations of the components, and hence every degree of re-
versibility of SH-inhibited enzymes may be observed. (2) Protection or
reversal by a thiol does not provide conclusive information on the mecha-
nism of the inhibition or the enzyme groups attacked. (3) Irreversibility is
brought about not only by very tightly bound reagents, but by progressive
structural changes in the enzyme, as discussed above. This type of irrever-
sibility thus increases with the concentration of the SH reagent and the
time of exposure. (4) The amount of useful information relative to the
mechanism of inhibition obtained from such studies is much less then
commonly believed.
The stability of the product formed by reaction of an enzyme SH group
with an SH reagent varies with many factors, most of which have been
mentioned in connection with the differential reactivity of these groups.
Sometimes the product is completely stable for all experimental purposes
and the reacted enzyme is permanently altered; such would be the case
with most of the alkylating agents. Then the mercaptide complexes vary
GENERAL CONSIDERATIONS OF THE USES OF SH REAGENTS 651
greatly in stability, so that in some instances they can be split at a rate
too rapid to be technically measurable, while in others the rate is too slow
to measure. It is not necessary that this stability be correlated with struc-
tural changes in the enzyme or irreversible inactivation. The inability to
reactivate an enzyme inhibited by an SH reagent can be attributed to a
variety of factors, some of which are listed below.
(A) The binding of the SH reagent to the enzyme is stronger than to the
reversor; one must use the proper reversor and concentration (e.g., dimer-
caprol will reverse some inhibitions untouched by cysteine).
(B) The enzyme is chemically altered by the SH reagent so that it is not
a question of a tightness of binding; reversal can occur only by a chemical
transfer of the attached group to another radical.
(C) The enzyme is structurally altered irreversibly (denatured) by the
blocking of the SH groups.
(D) The SH reagent has caused a splitting off of some coenzyme or co-
factor, which must be added back following restoration of the SH group
for activity to be evident.
(E) The reversor may in some manner inhibit the enzyme, even though
restoring the free SH groups initially, as when the SH groups are oxidized
by disulfides formed from the oxidation of the added thiols.
In order that irreversibility be correctly attributed to protein denaturation,
these other possibilities must be ruled out. Complete reversibility is more
easily interpretable, and one can quite confidently say that at least no per-
manent derangements in the enzyme structure have been induced by the
SH reagent.
Protection experiments, in which some thiol is added previous to, or with,
the SH reagent, are, as has been emphasized earlier, of little value, since all
one is doing is reducing the concentration of the free SH reagent (assuming
that it reacts with the thiol). Actually, it is a little difficult to speak of this
as protection, inasmuch as one usually would not call a reduction in inhib-
itor concentration a type of protection. It is very unlikely that worthwhile
information can be obtained from such experiments, aside from the practical
determination of the ability of substances to reduce the toxic effects of SH
reagents.
GENERAL CONSIDERATIONS OFTHE USES OF SH REAGENTS
Numerous types of reagent are available for satisfactorily specific reaction
with SH groups but no single one is adequate for all purposes. The most
useful information on the nature of enzyme SH groups, their locations and
652 4. SULFHYDRYL REAGENTS
relation to the catalysis, can be obtained by the proper use of several types
of SH reagent. It is also advisable to use different concentrations of the
reagents (it is not very informative to report that 1 mM of some SH reagent
inhibits 100%) and calculate a K, that characterizes the potency of the in-
hibition and the affinity of the enzyme for the reagent. In order to present
the kinetics properly, it is necessary to determine if the inhibitions under
the experimental conditions used are reversible, and for this purpose it is
best to perform the reversal study in nitrogen. It is also useful to determine
the degree of inhibition and the number and type of SH groups reacted
simultaneously in order to correlate reactivity and relationship to the active
center. Finally, at least some simple rate studies should be done to deter-
mine if the inhibitions observed are for equilibrium conditions. In many
reports one finds only that the enzyme was incubated with the SH reagent
for a certain period (even this information is frequently omitted) and it is
impossible to determine if the inhibition observed is maximal or not. SH
reagents have perhaps been used during the past several years more com-
monly than any other type of enzyme inhibitor and yet they have been
used with little concern for the many complexities involved in the interpre-
tation of the results, with a few notable exceptions.
Despite the generally good specificity of these reagents for SH groups,
they are not specific inhibitors from the metabolic standpoint in most cases.
Since SH groups are present not only in many enzymes but in most other
proteins of the cell, one must expect that in complex systems there will be
many components reacted. Whether some of these reactions will be of im-
portance in what is measured will depend on the nature of the work. It is
probably justifiable to suggest that the use of most SH reagents be restrict-
ed at the present time to enzyme studies for the purpose of determining the
nature of the enzyme SH groups. As the complexity of the system increases,
the value of SH reagents diminishes, at least if one is trying to correlate
some enzymic or metabolic process with over-all cellular metabolism or
function. In this connection it is interesting to note that although the most
reactive SH reagents are generally best for pure enzyme work, this is not
necessarily true for more complex systems. What one usually requires in
metabolic or functional investigations is specificity with respect to a partic-
ular enzyme or metabolic pathway. Thus iodoacetate, although it is a rather
poor reagent for the detection of SH groups and has frequently been ma-
ligned for this purpose, is actually more valuable in cellular work than most
of the others since it has the ability, if used properly, of inhibiting the phos-
phoglyceraldehyde dehydrogenase and glycolysis without affecting other
systems significantly, whereas a more reactive inhibitor, such as p-chloro-
mercuribenzoate, is valueless for producing specific metabolic blockade. The
choice of SH reagent to be used should always be made on the basis of the
type of work to be done. Another factor to be considered in work with eel-
GENERAL CONSIDERATIONS OF THE USES OF SH REAGENTS 653
hilar preparations is the penetrabihty of the SH reagent, and those reagents
should be chosen that have the most likelihood of reaching the systems to
be attacked. Thus iodoacetamide is often a better choice than iodoacetate
for intracellular inhibition because it is uncharged and probably enters cells
more readily.
The treatment of the individual SH reagents in the following chapters
must be eclectic in view of the immense amount of reported work, partic-
ularly during the past few years. The attempt will be made to select the
results of those investigations done most carefully and thoroughly, and to
include work on the most important or interesting facets of inhibition by
SH reagents. A third aim is to present all the available accurate data that
may aid in the assessment of the specificity of these inhibitors in order to
use them more profitably in complex systems.
CHAPTER 5
OXIDANTS
Many reagents have been used to oxidize protein and enzyme SH groups
for the purpose of either estimating these groups or determining the relation-
ship of the groups to the enzyme activity. Most of these oxidants at present
are of little importance in the study of enzymes or metabolism, mainly be-
cause of their lack of specificity for SH groups. General over-all oxidation
of an enzyme, involving several types of group and ending in partial or com-
plete denaturation, provides no useful information. If oxidants are to be
used for the specific modification of SH groups it is necessary that the choice
of oxidant and the experimental conditions be made very carefully. The
oxidizing activity of the reagent must be neither too high nor too low (i.e.,
its oxidation-reduction potential must be in the proper range relative to the
SH groups under the selected conditions) and the ability of the substance
to react in other ways with the enzyme must be minimal. Of the factors
determining the rates of oxidation of SH groups and the specificity of an
oxidant, the pH and the temperature are the most important. Some of the
oxidants that have been abandoned in enzyme work might well be applicable
in certain studies if the optimal conditions for their use were known.
The formation of enzyme disulfide groups during oxidation requires SH
groups that are close enough to link together in S — S bonds, or are so located
as to be able to approach each other readily. The SH groups may be on the
same enzyme molecule or on different molecules:
SH S
Intramolecular oxidation: R, ^ Xqx >'- R
SH
\
^red
Intermolecular oxidation:
2 R— SH -r Xox •* * R— S-S — R ^ Xred
The hydrogen atoms may be transferred directly to the oxidant, or may
form H+ ions, the oxidant accepting only electrons. A lone SH group on an
enzyme, although fully exposed, may not be oxidized if sterically it cannot
associate with another SH group. The formation of enzyme aggregates, or
655
656 5. OXIDANTS
actual precipitation, upon oxidation has occasionally been taken as evidence
for intermolecular disulfide bonding, but this is perhaps not always valid,
since the oxidation may bring about a dissolution of the protein structure
leading to such intermolecular reactions as occur during any type of de-
naturation. If reversal of aggregation can be induced by reducing agents, it
is more likely that disulfide bonds are responsible. One factor of primary
importance in determining whether a disulfide bond can be formed is the
steric relationship between the interacting groups. The C — S — S — C group-
ing is not linear, or even planar; the S — S — C bond angle is around 107° and
the dihedral angle between the two C — S bonds is close to 90° (due to the
electrostatic repulsion between unbonded electron pairs). Thus such bonds
will be formed readily only when the residues to which the sulfur atoms are
attached can assume the proper orientations.
The thiol-disulfide equilibrium:
R— SH ^ 1/2 (R— S— S— R) + H+ -f e-
has not been easy to determine, due to abnormal electrode reactions and
the usual sluggishness of such systems, and hence values for the oxidation-
reduction potential vary with the method used. It appears that Eq at pH 7
for various low molecular weight thiols generally lies between —0.35 and
0.0 (Calvin, 1954; Clark, 1960, p. 486). The values of E^' for protein SH
groups are not known, but it is likely that they would lie mainly in this
range also. It is possible, however, that some SH groups, due to their par-
ticular molecular environment, may have positive potentials, i.e., would be
less easily oxidized than most SH groups of the smaller compounds. It is
certainly true that certain SH groups on proteins, although readily acces-
sible to alkylation or mercaptide formation, are not oxidized readily, but
whether this is due to an especially high oxidation-reduction potential or
steric factors, as discussed above, is not known. The values of Eq depend
strongly on the pH, which must be taken into account when experiments
are run at pH's varying from neutrality. In any event, the oxidant should
have a rather high potential (probably 0.2 or higher) in order to oxidize
the susceptible SH groups to virtual completeness. On the other hand, it is
usually desirable to oxidize the SH groups only to the disulfide stage. Strong
oxidants can occasionally not only oxidize SH groups to sulfonate but at-
tack enzyme groups other than SH so that specificity is lost. Thus iodate
oxidizes gluten and thiolated gelatin mainly to the disulfide stage:
6 R— SH + lOg^ -► 3 R— S— S— R + 1+3 H^O
but further oxidation also occurs simultaneously:
R— SH + IO3- -> R— SO3- + H+ + I-
OXIDANTS 657
(Hird and Yates, 1961). o-Iodosobenzoate usually oxidizes only to the di-
sulfide stage at pH 7, but if too high a concentration is used, or the pH is
much below 7, sulfinate or sulfonate groups are produced (Hellerman et at.,
1941). In addition, methionine residues may be oxidized to the sulfoxide
stage. Sizer (1942 a,b, 1945) studied the effects on enzymes of many oxida-
tion-reduction systems over a wide range of Eq and found that as the Eq
is increased from around — 0.5 there is little effect on activity until a crit-
ical value is reached, which is +0.6 for /?-fructofuranosidase, +0.35 for
intestinal phosphatase, and +0.58 for chymotrypsin, inactivation increas-
ing rapidly above these values. Of course, it is not entirely a matter of the
Eq , the nature of the oxidant being also very important. The enzymes
used are not those containing SH groups most easily oxidized, but it does
indicate that rather strong oxidants must be used with many enzymes.
Oxidants can inhibit enzymes by mechanisms other than oxidation of SH
groups. They may (1) oxidize other enzyme groups, (2) be chemically in-
corporated into the enzyme (e.g., the iodination of tyrosine residues by
iodine), or (3) inhibit reversibly by any of the mechanisms observed with
nonoxidizing inhibitors. Other enzyme groups susceptible to oxidation are
the hydroxy! groups of tyrosine and serine, the hydrocarbon chain of leu-
cine, the indole ring of histidine, and perhaps the amino, guanidine, and
peptide groups. A few examples will be mentioned and others will be dis-
cussed in the sections on the individual oxidants. Lieben and Bauminger
(1933 a) showed that several amino acids are attacked by ^permanganate.
During the oxidation of casein, the arginine content falls, urea and dixan-
thylurea appearing. Haas et al. (1951) emphasized the importance of tyro-
sine and tryptophan in the actions of permanganate on proteins. Although
phenylalanine is quite refractory, tyrosine and tryptophan are oxidized, as
shown by changes in the ultraviolet spectra. The spectra of insulin and pep-
sin treated with permanganate (0.1 mM at pH 2) change in a manner sim-
ilar to that of the free amino acids, so it is likely that oxidation of these
amino acids occurs when they are part of the protein structure. Oxidation
of several proteins by periodate releases formaldehyde, which probably arises
from hydroxylysine (Desnuelle and Antonin, 1946). One mole of ovalbumin
reduces 30 moles of periodate to iodate, the protein losing all of its cysteine
and cystine, one third of its tryptophan, and a small fraction of its tyrosine
(Desnuelle et al., 1947). Oxidation of seralbumin by periodate results in
destruction of certain amino acids, producing changes in spectral and elec-
trophoretic properties (Goebel and Perlmann, 1949). Periodate releases acet-
aldehyde from chymotrypsin, arising from terminal threonine, although this
is not responsible for the inhibition of the enzyme inasmuch as it occurs
before inactivation starts, and in this case no ultraviolet spectral changes
are observed (Jansen et al., 1950, 1951). Nitrous acid not only oxidizes cer-
tain enzyme groups, such as SH, but attacks free tyrosine and amino groups
658 5. OXIDANTS
(Philpot and Small, 1938; Weill and Caldwell, 1945 a). Hypochlorite oxidizes
a number of amino acids, only glycine being resistant, and spectral changes
occur with proteins indicating oxidation of tyrosine and tryptophan residues
(Lieben and Bauminger, 1933 b); in addition it chlorinates amino groups
(Wright, 1926). Sizer (1942 b) noted in his work with many oxidants that
SH groups are by no means the only susceptible groups on enzymes, the
tyrosine residues being particularly oxidizable. These results point to the
importance of exercising great caution in the choice of oxidants and condi-
tions for treatment of enzymes if specific oxidation of SH groups is desired.
The need for characterizing well the enzyme changes — e.g., disappear-
ance of SH groups as determined by the standard methods, or alterations
in the ultraviolet spectrum — upon oxidation is also indicated.
Oxidation of Enzymes by Molecular Oxygen
Thiols and enzyme SH groups are not oxidized by Og unless certain metal
ions are present. Thus papain is oxidized by Og in the presence of Cu++ or
Fe+++ and the consequent inactivation of the enzyme is readily reversed by
glutathione (Hellerman and Perkins, 1934). Enzymes such as papain and
urease were the earliest studied with respect to the effect of oxidation on
their catalytic activities, and this work led to the concept wherein the redox
state of SH groups is an important regulating mechanism in cell metabolism
(Hellerman, 1939). The initial over-all reaction may be written as:
2 R— SH + O2 ±5 R— S— S— R + H2O2
but the hydrogen peroxide can produce further oxidation:
2 R— SH + H2O2 ^ R— S— S— R + 2 H^O
or it can oxidize other components present. The kinetics of the Cu++- and
Fe+++-catalyzed oxidations are complex and the mechanism is not comple-
tely understood. One theory involves the formation of a Fe++-thiol radical
from a Fe+++-thiol complex; two such radicals would combine to form the
disulfide and free Fe++, which is reoxidized by O2 (Williams, 1956). A sec-
ond theory postulates a Fe++ (thiol )2 chelate complex, which is oxidized
by O2 to the ferric complex, within which electron transfer occurs to form
the disulfide and Fe++ (Martell and Calvin, 1952). Since some type of com-
plex between metal ion and thiol must occur, it is evident that the suscepti-
bility of various SH groups to this type of oxidation must vary greatly.
It should be noted that the rates of such oxidations depend on the nature
of the buffer used and the pH.
The toxic effects of high tensions of Og on cell metabolism may depend
on the oxidation of enzyme SH groups. Brain respiration is slowly inhibited
by O2 and there is increasing inability of the tissue to oxidize glucose, pyru-
OXIDANTS 659
vate, lactate, fructose, and succinate (Dickens, 1946 a). It was suggested
that the most sensitive system is perhaps the pyruvate oxidase due to the
involvement of SH groups. This inactivation is not mediated through the
H2O2 formed, inasmuch as the concentrations are never great enough due
to the catalase present. The following enzymes are inactivated by high Og
tensions: succinate dehydrogenase, phosphoglyceraldehyde dehydrogenase,
choline oxidase, phosphoglucomutase, and other SH enzymes (Dickens,
1946 b). Lactate dehydrogenase, malate dehydrogenase, D-amino acid oxi-
dase, and yeast hexokinase are resistant. It is interesting that malonate
(1 mM) and Mn++ (0.25 mM) protect succinate dehydrogenase against oxi-
dation by O2, and that NAD protects phosphoglyceraldehyde dehydrogen-
ase, indicating that either O2 or some intermediate must react directly
with the enzyme SH groups. Haugaard (1946) also found a good correlation
between the SH nature of enzymes and their susceptibility to O2, the follow-
ing sensitive enzymes being added to the list above: a-ketoglutarate oxidase,
pyruvate oxidase, glutamate dehydrogenase, and xanthine oxidase. Dickens
stated that succinate dehydrogenase is irreversibly inactivated by 0^, but
Haugaard found reactivation by cysteine or glutathione, indicating a simple
disulfide formation. Inactivation of certain enzymes during extraction and
purification is probably due to oxidation by Og since metal-chelating agents,
such as ethylenediaminetetraacetate (EDTA), are able to protect against
such inactivation.
The cytochrome system may be involved in the inactivation of enzymes
by O2. Since cysteine is oxidized by O2 through the cytochrome system to
form cystine, and since cystine will in turn oxidize certain enzyme SH groups
(see page 661 ), enzyme extracts in which cysteine is present may be unstable.
Thus cysteine inhibits succinate dehydrogenase (Potter and DuBois, 1943),
but in mouse kidney homogenates the inhibition shows a lag period which is
interpreted as due to the necessary oxidation of cysteine to cystine by the
cytochrome system (Ames and Elvehjem, 1944 a, b).
Various Minor Oxidants
A number of strong oxidants, such as permanganate, perchlorate, dichro-
mate, and related compounds, have been used in the past to oxidize enzyme
groups. Most of these have dropped out of use because they were felt to
lack specificity toward SH groups, or other groups. Actually none of these
oxidants has been studied thoroughly with respect to what enzyme groups
are oxidized, or to the optimal conditions for achieving specificity. It is
quite possible that, at certain pH's and concentrations and temperatures,
these reagents may be specific oxidants. Certainly some enzymes are quite
susceptible and others very resistant, and it would be interesting to know
the reasons. For example, permanganate at 10 mM inhibits a-amylase
> 75% (Di Carlo and Redfern, 1947), at 0.05 mM inhibits /^-amylase 85%
660 5. OXIDANTS
(Ghosh, 1958), at 1 mM inhibits green gram flavokinase 48% (Giri et at.,
1958), at 0.1 mM inhibits /?-fructofuranosidase 94% (Sizer, 1942 a), at
10 mM inhibits /5-glycerophosphatase 100% (Rao et al, 1960), at 2 mM
inhibits Aerobacillus hydrogenlyase 23% (Crewther, 1953), at 0.01 mM in-
hibits intestinal phosphatase 95% (Sizer, 1942 b), at 1 mM inhibits Pseudo-
monas proteinase 100% (Morihara, 1963), and at 0.1 and 1 mM inhibits
beef liver urocanase 13% and 87%, respectively (Feinberg and Greenberg,
1959). Since these experiments were done at different pH's, temperatures,
and incubation times, it is difficult to compare the results accurately. In-
deed, no thorough investigation of the effects of pH or temperature on such
oxidations has been made. The moderate inhibition (12%) of liver arginase
by 5 mM permanganate was believed due to an effect on the Mn++ cofactor
rather than on the enzyme (Greenberg et al., 1956). Although yeast /?-fruc-
tofuranosidase is so sensitive to permanganate, it is inhibited only 16% by
10 mM dichromate (Sizer, 1942 a) and only 43% after 90 min incubation
with 123 mM periodate (Myrback, 1957 b). Dichromate is also less effective
than permanganate on /5-glycerophosphatase (Rao et al., 1960) and /?-amyl-
ase (Ghosh, 1958). In this connection, it must be remembered that the prod-
ucts of the reduction of the oxidant may also be inhibitory, e.g., the MnOg
or Mn++ from permanganate. The results of Taylor and Gale (1945) on E.
coli amino acid decarboxylases are interesting in that the effects of perman-
ganate were found to depend on the substrate used. For example, 0.1 mM
permanganate inhibits the decarboxylation of histidine 15%, arginine 17%,
glutamate 41%, ornithine 98%, lysine 100%, and tyrosine 100%. Whether
there are different enzymes or different effects with the various substrates
is not known. Permanganate and other strong oxidants can occasionally
act on substrates or other components of the reaction. This is illustrated in
the effects of permanganate and p-benzoquinone on the growth of Fusarium
conidia (Braune, 1963). Both are inhibitory alone but when present together
nullify each other and may actually stimulate growth. This was shown
not to be due to some oxidation product of jo-benzoquinone or reduction
product of permanganate. The formation was postulated of a substance X
which protects against p-benzoquinone and heavy metal ions. Indeed, treat-
ment of maleate or tartrate with permanganate gives rise to substance X.
Although such effects in cellular systems are complex, related actions must
be expected in certain enzyme systems.
Results with nitrous acid are difficult to interpret, but in all cases the
inhibition progresses very slowly (Myrback, 1926). Whereas the of-amylase
from B. suhtilis cannot be reactivated by HgS after inhibition by nitrite
(Di Carlo and Redfern, 1947), the /9-amylase of barley is completely reac-
tivated (Weill and Caldwell, 1945 a). In the former case it was concluded
that SH groups are not involved in the inhibition, and in the latter case
that they are. Similarly, inhibitions by redox dyes may or may not be at-
DISULFIDES 661
tributed to SH oxidation, but because of the molecular complexity of most
dyes it would be very unlikely that specific effects on SH groups could be
obtained. Inhibitions by dyes will be discussed in a separate chapter.
DISULFIDES
Cystine oxidizes certain protein SH groups and was first used for the de-
termination of these groups by Mirsky and Anson (1935). They found that
those protein SH groups reacting with nitroprusside are completely oxidized
by cystine. It was stated by Mirsky and Anson, and has been restated by
others, that cystine is one of the most specific oxidants of protein SH groups.
One disadvantage of cystine is its low solubility; dithioglycolate was sug-
gested as superior in this regard but has been used very little. Another dis-
advantage is perhaps that the oxidation-reduction potential of the cystine-
cysteine couple is not high enough to oxidize all the SH groups, which, if it
is assumed that the mean potential of protein SH groups is similar to free
cysteine, must be true. A further complication is the formation of mixed
disulfides (see page 639):
E— SH + R— S— S— R ±5 E— S— S— R + R— SH
which is not the simple oxidation of enzyme SH groups previously suppos-
ed, a portion of the disulfide being bound to the enzyme.
A few results on enzyme inhibition are summarized in Table 5-1. The in-
hibitions are not to be taken quantitatively because the conditions and the
incubation times vary greatly. Particularly important is the duration of
contact between the enzyme and the disulfide, since in almost all instances
the reaction has been found to proceed very slowly. Rapkine (1938) found
that phosphoglyceraldehyde dehydrogenase requires up to 5 hr for maximal
inhibition by GS,SG, and Hopkins et al'. (1938) showed that the inhibition
of succinate dehydrogenase develops steadily over 2 hr, and probably con-
tinues after that time. Whether this is due to the slowness of the oxidation
of enzyme SH groups or to secondary factors is not known. One reason for
the lack of inhibition of certain enzymes by disulfides may well be that suf-
ficient time was not allowed for reaction. Such slow reactions limit the use
of the disulfides for either SH group determination or enzyme studies. In-
sufficient examination of the reversibility of disulfide inhibitions by reduc-
tion also makes it difficult to evaluate the mechanism. Rapkine (1938) found
that both GSH and cysteine reactivate GSSG-inhibited phosphoglyceral-
dehyde dehydrogenase, and Hopkins and Morgan (1938) obtained similar
results with succinate dehydrogenase, but reversibility has not been at-
tempted in most work. Many enzymes require SH groups for activity but
others are active only when disulfide bonds are formed. Cytochrome oxidase
662
5. OXIDANTS
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DISULFIDES 663
normally contains disuL&de groups and can be inactivated by cysteine, GSH
and other thiols; reactivation occurs with cystine or GSSG (Cooperstein,
1963). This may also be the case with lens aminopeptidase, which is inhib-
ited rather strongly by cysteine and GSH but not at all by GSSG (Spec-
tor, 1963). Another reason for the lack of response to a disulfide is that
the environment of the enzyme SH group may be unfavorable for the ap-
proach of the disulfide or may affect the redox potential in such a manner
as to deter the interaction. If seralbumins are incubated with excess cystine,
the SH content as determined by p-mercuribenzoate titration drops to zero;
half the SH groups are lost from bovine hemoglobin (Isles and Jocelyn,
1963). GSSG has no effect on either type of protein. Incidentally, the reac-
tion of cystine with bovine seralbumin leads to the disappearance of 1 mol-
ecule of cystine for each pair of SH groups lost, so it is likely that the
cysteine formed is reoxidized and mixed disulfides are formed according to:
2 Cy— S— S— Cy - 2 Prot— SH ±-. 2 Cy— SH - 2 Trot— S— S— Cy
2 Cy— SH ^ Cy— S— S— Cy i- 2 H+ + 2 e-
Cy— S— S— Cy f 2 Prot— SH ^-. 2 Prot— S— S— Cy + 2 H^ -f 2 e-
Some evidence for the formation of mixed disulfides during the inhibition
of enzymes has been reported. Beef liver catalase contains 8 SH groups ti-
tratable with p-chloromercuribenzoate. Reaction with cystine-S^^ results in
the oxidation of 4 of these groups to disulfides and the formation of 4 mixed
disulfides {YM et al, 1961):
SH^ SH SH SH X— S— S S— S S — S— X
\ I I / _^ \ I I /
catalase^ -^ C— S — S— C >.: catalase
/ I I \ / I I \
SH SH SH SH X— S— S S— S S— S— X
This inhibition is spontaneously reversible; cystine at 0.037 mM, pH 7, and
370 inhibits maximally 17% around 5 min and then the inhibition decreases.
Cystamine monosulfoxide likewise appears to form mixed disulfides with
phosphoglyceraldehyde dehydrogenase, and this is reversible by thiols (Pihl
and Lange, 1962). Cystamine itself inhibits glucose utilization in erythro-
cytes and this block is believed to be at hexokinase (Eldjarn and Bremer,
1962). The inhibition is reversible by thiols, and mixed disulfide formation
was postulated; however, there was no direct evidence for this as opposed
to simple oxidation. Dithioglycolate and cystine were reported to form
mixed disulfides with myosin ATPase, but inhibition of the enzyme activity
does not occur until the last 2 or 3 SH groups are altered (Barany, 1959).
The optimal temperatures and pH's for reaction of enzyme SH groups
with disulfides are difficult to determine in our present state of knowledge.
Apparently the temperature coefficient is quite high; succinate dehydrogen-
664 5. OXIDANTS
ase is inhibited 20% at 20°, 78% at 30°, and 93% at 40^ by 100 mM GSSG
after 4 hr incubation (Hopkins et al., 1938). Thus one might expect incon-
veniently slow reactions at low temperatures. Yet Barany (1959) ran his
experiments with ATPase at 0^, although up to 15 hr was sometimes re-
quired for satisfactory reaction. Increase in the pH favors the oxidation of
enzyme SH groups to disulfides, since the — S~ form presumably reacts
more readily. Hence, incubation of the enzyme with the disulfide at pH's
around 9 may be useful where possible.
PORPHYREXIDE AND PORPHYRINDIN
Porphyrindin was originally synthesized by Piloty and Schwerin (1901 a,
b, c; Piloty and Vogel, 1903) but only much later attracted attention,
when it was studied by Kuhn et al. (1934) as the first clearly demonstrated
double free radical, and shown by Kuhn and Franke (1935) to have one of
the highest oxidation-reduction potentials among organic substances. It was
introduced as a reagent for the determination of protein SH groups by Kuhn
and Desnuelle (1938) because of its high potential and applied particularly
by Greenstein (1938; Greenstein and Edsall, 1940; Greenstein and Jenrette,
1942) for this purpose. Meanwhile its synthesis was improved by Porter and
Hellerman (1939). For the past 20 years it has been used sporadically to
inactivate enzymes by oxidation of the SH groups. Possibly it has been
neglected in enzyme studies since, although it is probably not as specific
for SH groups as is o-iodosobenzoate, it is certainly more selective than
most oxidants and, furthermore, reacts more rapidly and more completely.
Chemistry
The structures of porphyrindin and porphyrexide may be written in sev-
eral different ways because of resonance. Both in the crystalline state are
paramagnetic, the values indicating one unpaired electron in porphyrexide
and two in porphyrindin (Kuhn et al., 1934). The paramagnetism, however,
increases with temperature (Miiller and Miiller-Rodloff, 1935), suggesting
equilibria between diamagnetic and paramagnetic forms. Thus the resonance
structures for porphyrindin may be written as:
HoC O" ~0 CH, HjC O' b CH3
1 L .11 I 1+ +11
H3C-C— N N— C— CH3 H3C-C— N N-C— CH,
I \\ //I \ ■\ /-I
C— N=N— C -« >- C=N— N = C
I / \ I I / \ I
HN— C— N N— C = NH HN=C— N N— C = NH
H H H H
(diamagnetic) (paramagnetic)
Porphyrindin
PORPHYREXIDE AND PORPHYRINDIN 665
while for porphyrexide only the free radical form is possible:
H3C 6
I U
H,C— C— N
I *\
C = NH
HN=C— N
H
Porphyrexide
The diamagnetic form is more stable than the paramagnetic by about 0.56
kcal/mole. Magnetic studies have indicated the free electrons to be fairly
well localized and not diffusely distributed over the molecule. Possibly these
free electrons contribute to the color of these substances: porphyrexide is
red and porphyrindin a deep blue. Upon reduction the color disappears
(leucoporphyrindin may be slightly yellow), this being the basis for the col-
orimetric titration of SH groups. Porph>Texide has an absorption maximum
at 460 m// and porphyrindin at 653 m// (Kuhn and Franke, 1935).
Reduction involves the change from
O- O- OH
I I I
— N+= or — N+— to — N —
and a disappearance of free radicals. The oxidation-reduction potentials at
pH 7 and 18° are:
Porphyrexide: £"„' = + 0.725 v
Porphyrindin: E^' = + 0.565 v
SO that porphyrexide approaches the oxygen potential and both are well
above most systems commonly seen in biological work. The oxidation of
simple thiols is very rapid, cysteine and glutathione reacting almost instan-
taneously.
Porphyrindin is not very stable and the solid crystalline dye should be
stored at low temperatures and desiccated. Greenstein (1938) pointed out
that porphyrindin solutions are stable enough for an hour but the activity
then decreases. Brand and Kassell (1940) reported that solutions at 0^ show
3% deterioration in 1 hr, 5% in 2 hr, and 9% in 4 hr. At 25^ deterioration
occurs at a rate of about 0.5% per minute. Reactions of enzyme SH groups
can usually be carried out at 0°, but in work with tissues at physiological
temperatures this spontaneous decomposition must be borne in mind. Por-
phyrindin should be quantitatively determined in all accurate work since
it is seldom pure; this may be done by titrating with asorbic acid (Chinard
and Hellerman, 1954).
666 5. OXIDANTS
Spiro analogs of both porphyrexide and porphyrindin were synthesized
by Porter and Hellerman (1944) and found to have high oxidation-reduc-
O" O 0
N— C— NH / \ N=C— N=N— C=N,
C— NH ' ' C— NH HN— C
II II II
NH NH NH
Spiroporphyrexide Spiroporphyrindin
tion potentials. The Ef^ at pH 7 for spiroporphyrexide is + 0.69 v, the po-
tential increasing at lower pH's. The spiroporphyrindin is quite insoluble
and may be more polymerized than indicated. An interesting aspect of spiro-
porphyrexide is that it does not inhibit urease, despite its high potential,
indicating possible steric effects of the cyclohexyl ring. This is a good exam-
ple that not only is oxidation-reduction potential important in the oxida-
tion of SH groups on enzymes but that structural configuration is a factor,
as in any inhibition.
Oxidation of Thiols and Amino Acids
Porphyrexide and porphyrindin react very rapidly with thiols at neutral-
ity and the end-point is generally quite sharp; cysteine, glutathione, and
cysteinylcysteine are titrated quite comparably (Greenstein, 1938). The oxi-
dation of cysteine at pH 7.2 is complete within 3-60 sec and glutathione is
oxidized only slightly more slowly (Brand and Kassell, 1940). No reaction
under ordinary conditions is seen with cystine, cysteate, tryptophan, hydro-
xyproline, histidine, methionine, serine, phenylalanine, or threonine. Tyro-
sine, however, is oxidized slowly with the formation of a pink-orange color,
the reaction taking around 30 min for completion (2 equivalents of porphy-
rindin per mole of tyrosine) at pH 7.2 and 0°. Tyrosine and other phenols
are oxidized more rapidly in alkaline solutions and at higher temperatures,
but in most cases the reaction is much slower than the oxidation of SH
groups (Greenstein and Edsall, 1940). At pH 7.33 and 25^ the oxidation of
tyrosine may be fairly rapid, and even tryptophan may be slowly reacted
(half-reaction time around 2 hr) (Barron et al., 1941). Porphyrindin can
also oxidize a variety of other substances, such as ascorbate or thiamine
(Kuhn and Desnuelle, 1938), and in complex systems or cellular prepara-
tions the effects may not be due entirely to SH group oxidation.
Oxidation of Proteins
Kuhn and Desnuelle (1938) showed that native ovalbumin does not react
with porphyrindin (in common with other SH reagents) but that following
PORPHYREXIDE AND PORPHYRINDIN 667
heat denaturation, titration gives results comparable to other methods for
total cysteine. This was confirmed by Greenstein (1938), who used urea and
guanidine for denaturation. The rapidity of SH oxidation was noted. Brand
and Kassell (1940), on the other hand, did not find good end-points with
denatured ovalbumin and, because of the pink color developed, felt that
tyrosine groups are also oxidized. This was criticized by Greenstein et al.
(1940) on the basis that far too much porphyrindin was used, and they em-
phasized that such high concentrations are probably not specific and should
be avoided. The method was somewhat improved (Greenstein and Edsall,
1940; Greenstein and Jenrette, 1942) by reducing the reaction time and
lowering the pH to 6.4-6.8 and, using myosin, seralbumin, ovalbumin, and
tobacco mosaic virus, it was believed that accurate titration of the SH
groups could be achieved with little interference from tyrosine oxidation.
Barron et al. (1941) treated scarlet fever toxin with 1 mM porphyrindin for
1 hr at pH 7 and found that the activity of the toxin, as determined by skin
tests, is abolished. Since other SH reagents do not inactivate the toxin, it
was felt that oxidation of groups other than SH is involved. However, the
SH reagents used (iodoacetate, iodoacetamide, hydrogen peroxide, alloxan,
and Cu++) are not the most satisfactory for the demonstration of SH groups,
so this evidence is not conclusive. In order to achieve specificity toward SH
groups it is advisable to (1) avoid alkaline conditions, (2) reduce the reac-
tion time with porphyrindin as much as possible, (3) use as low concentra-
tions of porphyrindin as possible, and (4) determine the disappearance of
SH groups by some secondary titration.
Inhibition of Enzymes
Results of treating enzymes with these oxidants are shown in Table 5-2;
one cannot help but be surprised that so little use has been made of these
substances, especially during the past few years. It is evident that quite
low concentrations are needed for those enzymes which have susceptible
SH groups and that porphyrindin and porphyrexide are among the most
potent oxidant inhibitors.
Balls and Lineweaver (1939 b) attempted to titrate papain with por-
phyrindin but found that no clear end-point could be obtained at room
temperature, and at 2-3° no bleaching of the dye occurred during several
minutes when dilute concentrations were used. Higher concentrations pro-
duce a pink coloration, even at pH 4.6. The native papain SH groups are
thus not reactive with porphyrindin; neither are they reactive with nitro-
prusside. On the other hand, iodoacetate and iodoacetamide react and in-
hibit; papain treated with these alkylating agents still gives rise to the pink
coloration, indicating that tyrosine is oxidized, although not necessary for
enzyme activity. E. L. »Smith (1958) concluded that the SH group which is
668
5. OXIDANTS
M
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PORPHYREXIDE AND PORPHYRINDIN 669
at or near the active center of papain is in a high energy state, perhaps as a
thiol ester, and this may explain why it is resistant to porphyrindin.
The reactive SH groups of urease are rapidly oxidized by porphyrindin
and the nitroprusside test becomes negative (Hellerman, 1939). However,
no inhibition occurs. Porphyrindin-treated enzyme is inhibited by jo-chloro-
mercuribenzoate, so that certain SH groups required for activity are resist-
ant to porphyrindin. If an excess of porphyrindin is used, inactivation oc-
curs slowly and is irreversible. Oxidation past the disulfide stage or oxida-
tion of other groups is possible. Since, p-chloromercuribenzoate protects the
enzyme from high concentrations of porphyrindin, it appears that SH groups
are indeed involved. It was established later that there are two types of SH
group in urease: reactive a groups not necessary for enzyme activity, and
less reactive b groups at the active center. Porphyrindin reacts with the
former but only slowly with the latter at higher concentrations (Hellerman
et al., 1943). If urease is denatured with guanidine, many more SH groups
appear and react with porphyrindin.
Xanthine oxidase is inhibited readily by porphyrindin but this is not re-
versible with cysteine (Harris and Hellerman, 1956). The inhibition by o-
iodosobenzoate is also irreversible. This problem comes up repeatedly with
inhibitions by oxidants and seems on the surface to indicate that a simple
oxidation to the disulfide stage does not occur. However, it is also possible
that (1) the oxidation-reduction potential of the groups involved is such
that cysteine will not reduce them, (2) the enzyme structure is altered by
the formation of disulfide bonds, (3) oxidation past the disulfide stage has
occurred, or (4) the formation of intermolecular disulfide linkages prevents
the access of cysteine to the group. It is impossible to distinguish at this
time between these different possibilities.
Effects on Tissue Function
Porphyrindin at 0.37 0.75 mM produces an increase in the contractile
amplitude of the frog heart and this effect can last for as long as 90 min
(Mendez, 1946; Mendez and Peralta, 1947). Higher concentrations of 3.7-
7.5 mM bring about a progressive contracture, the heart stopping in systole
in around 30 min. The rate is simultaneously slowed. The atria continue to
beat after the ventricles have stopped. These effects can be prevented by
glutathione but not reversed, as expected. In these respects the heart re-
sponds to porphyrindin much as it does to other SH reagents. The site or
sites of action are not known, and it is useless to speculate since the relative
sensitivities of the possible enzymes involved are undetermined.
A few miscellaneous and unrelated observations will be mentioned. The
short-circuit current and potential of frog skin are altered by oxidants and
reductants, such as quinones, dyes, and iodine, but there is little effect of
porphyrindin at 1 mM; the potential may drop temporarily but the cur-
670 5. OXIDANTS
rent is unaffected (Eubank et al., 1962). Porphyrindin injected at a dose of
200 mg in a pregnant mouse produced neuroblastic necrosis in the fetus,
but not as much as p-chloromercuribenzoate, oxophenarsine, or o-iodosoben-
zoate, these damaging neuroblasts in a pattern similar to radiation (Hicks,
1953). SH reagents usually cause blebbing of Sarcoma 37 ascites cells. Por-
phyrindin has no effect at 2 nxM but produces symmetrical blebs at 8 roM
(Belkin and Hardy, 1961). Such blebbing involves a raising of the entire cell
membrane and presumably is due to some disturbance in water transport.
FERRICYANIDE
Ferricyanide has been used widely as a fairly specific oxidant for the de-
termination of protein SH groups. Furthermore, it is commonly used as an
electron acceptor in various dehydrogenase systems, since it is reduced by
some of the components in electron transport before cytochrome c, and fer-
rocyanide has often served as an electron donor in studying the cytochrome
system. When ferricyanide or ferrocyanide is used for such purposes, it is
important to consider the possibility of inhibition of the enzymes involved,
particularly the dehydrogenases, and to use as low concentrations as pos-
sible. The first use of ferricyanide for the determination of SH groups was
by Flatow (1928) and this was simplified by Mason (1930) so that the ferro-
cyanide formed could be colorimetrically estimated after transformation to
Prussian blue, this being suggested by Folin's ferricyanide method for blood
glucose. This reaction has been used for the histochemical localization of SH
groups but is not very satisfactory. Anson and Mirsky (1931) noted that
hemoglobin treated with ferricyanide yields a globin that no longer reacts
with nitroprusside, but it remained for Schiiler (1932) to show that ferri-
cyanide oxidizes more than the heme group and that globin itself reacts
after separation from heme. Mirsky and Anson during the next 10 years
elucidated the nature of the reactions between ferricyanide and proteins,
and applied their results to determination of protein SH groups.
Chemistry
The oxidation of thiols may be written as:
2 Fe{CN),^- + 2 R— SH ±^ 2 Fe(CN)6''- + R— S— S— R + 2 H+
When the SH groups are on different molecules, the kinetics are complex
and the detailed mechanism of the reaction is not understood. The ferro-
cyanide formed is usually determined by addition of Fe+++, forming Prus-
sian blue, but in work with proteins it is advisable to determine also the
disappearance of SH groups with nitroprusside or ??-chloromercuribenzoate,
since groups other than SH may be oxidized. The oxidation-reduction po-
FERRICYANIDE 671
tential of the ferricyanide-ferrocyanide couple is + 0.36 v and does not
change from pH 4 to 10. The potential is, however, rather strongly depen-
dent on ionic strength. The potential is thus sufficiently high for SH groups
to be oxidized completely if they are available to the ferricyanide, and the
reaction is generally quite rapid.
Most commercial preparations of ferricyanide contain ferrocyanide, which
may be detected by the Prussian blue method, and the latter should be re-
moved by addition of a little bromine water if SH determinations are done
by the colorimetric technique. Solutions of ferricyanide should be stored in
the cold and dark to avoid changes.
Oxidation of Thiols and Proteins
The reactions of ferricyanide with proteins have direct bearing on the
effects of ferricyanide on enzymes so that it is necessary to discuss the re-
sults in some detail. Although titration of cysteine and glutathione with
ferricyanide is rapid and provides good end-points, reactions with proteins
may not be so clear-cut. The conditions for the reaction are very important.
There is a marked effect of pH, as shown originally by Mirsky and Anson
(1936 a) for hemoglobin (see tabulation). Indeed, at pH 6.8 one may spe-
pH Total SH groups oxidized (%)
6.8
0
7.3
28
9.0
44
9.5
65
cifically oxidize the heme iron to form methemoglobin without affecting SH
groups. The conditions for reaction were: 83 mill ferricyanide incubated with
the protein for 30 min at room temperature — all reactive SH goups are
oxidized, as shown by titration of SH groups in denatured globin. Kolthoff
and Anastasi (1958) have also observed that oxidation of the SH groups of
denatured seralbumin is faster at pH 9 than 7. They noted that the reac-
tion is accelerated by Cu++, and Katyal and Gorin (1959) found in a study
of ovalbumin that iodide also catalyzes the oxidation by ferricyanide.
Ferricyanide is not specific for SH groups, however, unless the conditions
are rigorously controlled, as shown early by Mirsky and Anson (1936 b) in
proteins not containing cysteine (zein and serum globulin) but nevertheless
reducing ferricyanide, or in proteins previously treated with cystine to oxi-
dize all the available SH groups. These groups are oxidized more slowly than
the SH groups and are more difficult to oxidize (e.g., milder oxidants than
672 5. OXIDANTS
ferricyanide will not oxidize them), but their total reducing capacity (the
amount of ferricyanide they can reduce) is often greater than for the SH
groups. Furthermore, the rate and degree of oxidation of these non-SH
groups depend on the same factors as reaction with SH groups; thus, the
rate is accelerated by rise in pH, rise in temperature, and denaturation,
Native ovalbumin is not oxidized at all bj^ ferricyanide, but denatured oval-
bumin treated with cystine to remove SH groups reduces ferricyanide. In
other words, these other groups become available during unfolding of the
protein. /5-Lactoglobulin, which contains 2 SH grovips per molecule (molec-
ular weight of 37,000), reacts very slowly with ferricyanide in the native
state but rapidly in the presence of urea or guanidine (Leslie et al., 1962 a).
The stoichiometry indicates that the SH groups are oxidized beyond the
disulfide stage. Since the reaction was carried out under fairly mild condi-
tions (0.1-0.8 mM ferricyanide, pH 7, 37^, and 30-45-min incubation), it
is evident that one cannot generally assume the simple formation of disul-
fides from the actions of ferricyanide on enzymes. Ferricyanide can oxidize
tyrosine and tryptophan, but not histidine, and the characteristics of the
oxidation parallel oxidations of proteins. Mirsky and Anson suggested that
tyrosine and tryptophan are the residues responsible for ferricyanide re-
duction. Gelatin, which contains no (or very little) tyrosine and no tryp-
tophan, scarcely reacts with ferricyanide, supporting this view. Anson
(1939 b) observed that at pH 9.6, where much previous work had been
done, the oxidation is nonspecific, but that at pH 6.8 combined with the
treatment of the protein with Duponol PC the SH groups react rapidly and
specifically if not too much ferricyanide is used (2-5 mM is best); under
these conditions there is no reaction with cystine, tyrosine, tryptophan, or
proteins that do not contain cysteine. The specificity of SH group oxida-
tion could also be shown by pretreatment of denatured ovalbumin with
iodoacetamide, following which ferricyanide is no longer reduced by the
protein. Katyal and Gorin (1959) also demonstrated such specificity by
blocking SH groups with p-chloromercuribenzoate. Mirsky (1941) discussed
the method in detail and showed that when properly run the oxidation oc-
curs within 1 min. Barron (1951) has also reported in detail his modifica-
tion of the method. Various oxidations by ferricyanide have been reviewed
by Thyagarajan (1958).
Inhibition of Enzymes
One must conclude from the results with proteins that application of ferri-
cyanide to enzymes cannot be done haphazardly if specific effects on SH
groups are to be anticipated. Unfortunately most studies have used ferri-
cyanide along with numerous other inhibitors under the same conditions
of pH, temperature, and incubation time, without considering that rather
stringent conditions have been proposed for the use of ferricyanide. Some
FERRICYANIDE 673
inhibitions are summarized in Table 5-3. Certain enzymes which possess SH
groups reactive with other reagents, e.g. urease, are not inhibited by even
high concentrations of ferricyanide. One might imagine ferricyanide to be
unable to gain access to the SH groups. Ferricyaiade is not only a fairly
large ion but has a strong negative charge. If the enzyme SH group occu-
pied a region of high negative charge, this might repel the ferricyanide and
reduce the reaction. Indeed, one must always consider the possibility that
ferricyanide inhibits certain enzymes by mechanisms other than oxidation,
and related more to its charge and structure. For example, it would not be
so surprising if ferricyanide inhibits succinate dehydrogenase to some ex-
tent because it can interact with the cationic groups normally binding suc-
cinate. One notes also that ferrocyanide generally inhibits aconitase more
strongly than does ferricyanide, and here redox reactions may be of no sig-
nificance (Rahatekar and Rao, 1963). On the other hand, some enzymes
are inhibited just as rapidly and completely by ferricyanide as by the more
commonly used SH reagents; myosin ATPase is one example (Singer and
Barron, 1944). The inhibitions of papain and aldolase are quite reversible
with cysteine, indicating that reversible oxidation is the mechanism of the
inhibitions. Oxidation of coenzymes or cof"ctors can also occur. Ferricyanide
can directly oxidize NADH but the rate is slow (Schellenberg and Heller-
man, 1958). In the case of homogentisicase it may well be the Fe++ that is
oxidized but, on the other hand, there appears to l)e a tyrosine phenolic
group at the active site (Tokuyama, 1959).
It is interesting that Weill and Caldwell (1945 b) report /5-amylase to be
not readily inhibited by either ferricyanide or Cu+^ alone but strongly in-
hibited when both are present, even when the ferricyanide is at a concen-
tration noninhibitory by itself (see accompanying tabulation). Could this
Ferricyanide Cu++
(mM) (mM)
% Inhibition
0.2
—
12
—
0.32
4
0.02
0.32
93
relate to the observation of Katyal and Gorin (1959) that Cu++ accelerates
the action of ferricyanide ? Or does the Cu"^+ in some manner alter the en-
zyme structure so that ferricyanide can attack the SH groups more easily?
Effects on Cellular Metabolism and Function
Mendel (1937) reported that Balogh mouse tumor glycolysis is markedly
depressed by 10 mM ferricyanide and that the inhibition is maintained
674
5. OXIDANTS
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5. OXIDANTS
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FERRICYANIDE 677
when the ferricyanide is washed out. Ferricyanide slowly injected intraven-
ously into mice (1.5 g/kg of sodium salt) produces no disturbance of tumor
metabolism, but when the various tissues are treated with re++"'", only the
tumor turns blue. The anaerobic glycolysis of no other tissue is depressed
by ferricyanide, which in this respect differs from other glycolytic inhibitors
(Mendel and Strelitz, 1937). Renal medulla was studied particularly because
it has a significant rate of aerobic glycolysis and ferricyanide was found to
have no effect (and in some cases even stimulated somewhat). This action
was not investigated further until Birkenhager (1959, 1960) attempted to
locate the site of inhibition. He confirmed that 10 mM ferricyanide does
indeed inhibit both aerobic and anaerobic glycolysis in Walker and Crocker
tumors, but not in Ehrlich ascites cells, and further showed that it inhibits
the extra glycolysis brought about by dinitrophenol. The respiration in the
presence of glucose rises 30-60% in the presence of ferricyanide in the two
former tumors, but not in the ascites cells. The problem of what happens
to the glucose taken up, since this is not depressed as much as lactate for-
mation, remains unsolved. A small accumulation of pyruvate was found
under aerobic conditions but not anaerobically, and no other glycolytic in-
termediates could be detected. Use of glucose- 1-C^* and glucose-6-C^* point-
ed to the conclusion that ferricyanide either directly or indirectly inhibits
glycolysis at the level of phosphohexose isomerase or phosphohexokinase;
this would make more hexose phosphate available for the pentose phosphate
shunt. However, aldolase was found to be very sensitive to ferricyanide
(88% inhibition at 0.5 mM) and addition of aldolase to a tumor extract in
which glycolysis has been abolished by ferricyanide leads to recovery. Phos-
phoglyceraldehyde dehydrogenase is not sensitive to ferricyanide nor does
its addition reverse the glycolytic inhibition. Birkenhager ascribed the dif-
ference between cells and extracts in susceptibility to ferricyanide as due
to permeability factors. Certainly one might expect an ion such as ferri-
cyanide to enter cells with difficulty. However, the initial observation of
Mendel that tumor tissue is specifically sensitive to ferricyanide remains to
be explained. If such a difference exists, it must be due to ferricyanide
penetrating into tumor cells more readily, since none of the enzymes con-
sidered to be the point of attack differs markedly in tumor cells compared
with normal tissues.
Inasmuch as ferrocyanide is presumably formed in tissues during the
reduction of ferricyanide, the effects of ferrocyanide on the tricarboxylate
cycle may play a role in any over-all action. Martin (1955) noted that
growth of Aspergillus niger is inhibited by ferrocyanide at concentrations
below 0.002 mM. However, acid production may not be simultaneously in-
hibited, and is depressed 50% only at 0.4 mM. An accumulation of citrate
is actually observed at 1 mM ferrocyanide and, at this concentration, iso-
citrate dehydrogenase is inhibited 100% (Ramakrishnan et al., 1955). In
678 5. OXIDANTS
A. tereus 0.1 mM ferrocyanide has no effect on glucose utilization but in-
creases the yield of itaconate, due presumably to the piling up of citrate
(Bentley and Thiessen, 1957). The uptake and metabolism of itaconate are
inhibited by ferrocyanide, which is reasonable on the basis of the inhibition
of isocitrate dehydrogenase, and the assumption that itaconate feeds into
the cycle (Shimi and Nour El Dein, 1962). There has been very little work
on the effects of either ferro- or ferricyanide on cycle enzymes, but it ap-
pears likely that ferrocyanide blocks tricarboxylate steps selectively, where-
as ferricyanide less potently inhibits pyruvate oxidation. More work should
be done on these effects since no other specific inhibitor of isocitrate dehy-
drogenase is known.
A few miscellaneous observations on ferricyanide may be interpreted
when more is known of the basic actions. Thus, 10 mM ferricyanide inhibits
P^2 incorporation into phospholipids 19% while stimulating respiration 17%
in M. tuberculosis, in this way acting more like the uncoupling agents (azide,
dinitrophenol, and arsenate) than the common SH reagents (Tanaka, 1960).
Whereas other oxidants and SH reagents frequently cause mitochondrial
swelling, ferricyanide is without effect, which could scarcely be due to per-
meability factors (Rail et al., 1962). Porphyrin synthesis from porphobili-
nogen is strongly inhibited by 1 mM ferricyanide; this may be partly the
result of direct oxidation of the porphyrins (Rimington and Tooth, 1961).
The eggs of Urechis and Hemicentrotus are very sensitive to ferricyanide,
0.02 mM elevating the fertilization membrane in the former and causing
cytolysis in the latter (Isaka and Aikawa, 1963). The dorsal-ventral modifi-
cation i^roduced hy ferricyanide in Dendraster eggs, whereby either dorsal
induction or ventral inhibition is manifest, is similar to that produced by
iodoacetate or iodine, but is unexplainable since the factors involved in
bilaterality are completely unknown (Pease, 1941). The effects of ferricy-
anide on the naturally occurring quinones must occasionally be important.
For example, ferricyanide potentiates very markedly the growth-inhibiting
activity of menadione on yeast, due to the fact it reoxidizes the reduced
menadione and hence maintains the naphthoquinone in the active form
(Kiesow. 1960 b).
IODINE
Iodine has been used more frequently than the oxidants previously dis-
cussed for the oxidation of protein 8H groups and in enzyme studies, and
yet it seems under most conditions to be less specific than the others. Al-
though it is quite a potent inhibitor of many enzymes, unless one can de-
termine if a particular protein group is oxidized, or otherwise attacked, the
information obtained is negligible. Another complication in the use of iodine
is the multiplicity of forms in solution and the difficulty in characterizing
the nature of the oxidation reaction.
IODINE 679
Chemistry
Iodine is soluble to the extent of 1.33 mM in water at 20°, which is much
less than the other halogens. There is interaction with the water, which
initially was written as a hydration:
I, + H,0 ^ I, ■ H^O
but evidence pointing to the highly polarized state of iodine in the complex
has suggested the following reaction:
I, + H3O t:, 1+ . H,0 + I-
The equilibrium constant for this reaction has been estimated as roughly
lO^^*'. Iodine may also undergo hydrolysis:
I2 + H2O ^ I- + HOI + H+
the equilibrium constant being 3 X 10~^^. The hypoiodous acid formed has
a p^^ of 12.3 so is little ionized at physiological pH's. The hypoiodous acid
can also go to iodate, especially in alkaline solution:
3 HOI -> IO3- + 2 I- + 3 H+
A third reaction of iodine is with the iodide ion:
I, + I- ±^ I3-
to form the triiodide, which is the principal reason for the greater solubility
of iodine in KI solutions. The equilibrium is given by:
(I2) (I-)
1.38 X 10-3
(I3-)
Thus iodine would be soluble in 50 mM KI solution to the extent of 46 mM,
an appreciable increase over the 1.33 mM in water. The production of nas-
cent oxygen by the reaction:
I2 + H2O -> 2 H+ + 2 I- + O
which has been believed to be involved in protein oxidations, does not occur
with iodine, although the other halogens react to some extent in this way.
In most biological work, iodine is dissolved in fairly strong KI or Nal so-
lution. This not only serves to increase the solubility, but limits the fraction
of the iodine in other forms (HOI, IO3", and I+'HaO); the principal form
here is presumably the Ig" anion. However, there will always be significant
concentrations of iodine present. The relative importance of the Ig and Ig"
680 5. OXIDANTS
forms in the oxidation of SH groups is not known. The oxidation-reduction
potential for iodine varies with the type of reaction in which it participates
and the pH, but is usually sufficiently high to oxidize any accessible SH
groups. It is important in certain enzyme studies to realize that iodine may
disappear fairly rapidly from solution, independently of reaction with or-
ganic materials; such is favored by lack of iodide and high pH.
Reaction of Iodine with Thiols
Iodine is able to oxidize SH groups to four different states: the disulfide
(S — S), the sulfenate (SO"), the sulfinate (SOg"), and the sulfonate (SO3-).
Apparently it is quite easy to oxidize beyond the disulfide state with iodine.
The stoichiometry of a particular reaction will depend not only on the state
of oxidation of the SH groups, but also on the degree of reduction of the
iodine, this varying with the pH. For example, the following reactions can
be written for oxidation to the disulfide state:
I2 + 2 R— SH -> R— S— S— R -I 2 I- + 2 H+
2 I2 + 2 R— SH + H2O -> R— S— S— R + HOI + 3 I" + 3 H+
4 I2 + 2 R— SH + 3 H2O -> R— S— S— R + IO3- + 7 I- + 8 H+
However, it is likely that near neutrality the first reaction is dominant. In
the oxidation of cysteine, 3 equivalents of iodine 'are taken up to form
cysteate:
3 I2 + R— SH + 3 H2O -► R— SO3- + 6 I- + 7 H+
It is interesting that, at pH 3.2, iodine oxidizes cysteine well, but does not
react with cystine, tyrosine, or histidine. This indicates that the first prod-
uct in the oxidation of cysteine is not cystine but free radicals, which can
either combine to form disulfides or be further oxidized to sulfonate groups
(Anson, 1940). At pH 6.8, cystine is the major product. In most instances,
especially with proteins, several reactions will occur and mixed products
will be found. In addition to these straightforward oxidations, we shall see
that there is now evidence for the formation of sulfenyl iodide groups (SI),
so that a certain fraction of the iodine can be incorporated into the thiol
molecule.
Reactions of Iodine with Proteins
The SH groups of denatured ovalbumin are oxidized by iodine within 5
min at pH 3.2 and 37° (Anson, 1940). The rate of the reaction decreases as
the pH is raised to 6.8. In acid media iodine does not react with tyrosine or
proteins containing tyrosine (e.g. pepsin), whereas at neutrality it readily
iodinates tyrosine. By proper choice of pH and iodine concentration it is
possible to oxidize the SH groups of denatured ovalbumin without appre-
IODINE 681
ciably altering tyrosine residues (Anson, 1941). Iodine reacts only with the
tyrosine residues of seralbumin, and denaturation accelerates the formation
of diiodotyrosine (Li, 1945). The rate of the reaction is, however, quite slow
in native seralbumin (half-reaction time around 100 min). Human seral-
bumin iodinated at low temperature takes up 36 atoms of iodine per mole
of albumin, but only 12 diiodotyrosyl groups are found (Hughes and Straes-
sle, 1950). The remainder was believed to be incorporated into histidyl re-
sidues. Some oxidation of cysteinyl residues also occurs and this presumably
is beyond the disulfide stage, since 2.2 moles of iodine are taken up per SH
group. In any particular case, the amount of disulfide formed will depend
to a large extent on steric factors, i.e., how readily the sulfhydryl radicals
can combine; the seralbumin molecule is fairly large and, not surprisingly,
disulfide groups are not found after oxidation. Although no degradative
changes in seralbumin are observed, protein structure is certainly modified
by treatment with iodine, since the water binding capacity is increased
(Jensen et at., 1950) and the rates of pepsin and trypsin digestion are de-
creased (Raghupathy et al., 1958).
We have noted that 2 to 3 atoms of iodine are occasionally utilized for
each protein SH group. This might indicate (1) oxidation of SH beyond
the disulfide state, (2) reduction of the iodine beyond the iodide state, or
(3) some substitution of iodine in the cysteinyl residue. This problem was
studied by Fraenkel-Conrat (1955) with tobacco mosaic virus protein. It
is possible that sulfenate or sulfenyl iodide groups might be produced, but
it has always been thought that such groups are quite unstable and cannot
exist for appreciable time. However, the virus protein SH groups react with
2 atoms of iodine fairly rapidly, and this was shown to be accompanied by
the formation of sulfenyl iodide groups:
V— SH + I2 -> V— SI + I- + H+
This group appears to be stable in this particular protein. Fraenkel-Conrat
pointed out that further reaction with thiols can form mixed disulfides:
V— SH + R— SH ±^ V— S— S— R + H+ -f- I"
Such reactions have been studied further in /5-lactoglobulin by Cunningham
and Nuenke (1959, 1960, 1961), using a spectrophotometric method. This
protein reacts with 4 equivalents of iodine to form 2 sulfenyl iodide groups
per mole of protein:
P(— SH), + 2 I, -> P(— SI), + 2 I- + 2 H+
Ovalbumin reacts similarly but 6 equivalents of iodine are taken up. The
sulfenyl iodide groups are quite stable in these proteins, but can react with
simple thiols (e.g., glutathione, cysteine, and others) to form mixed disul-
682 5. OXIDANTS
fides. Intermolecular disulfide formation was ruled out for these proteins.
Ovalbumin has 4 SH groups and 2 S — S groups; the protein treated with
iodine has 2 SH groups and 2 S — S groups, and has incorporated 1 iodine
atom (Winzor and Creeth, 1962). Since 5 atoms of iodine are taken up, it
is not a simple oxidation to disulfide. It was suggested that the following
reactions occur:
2 P(— SH)2
HS-P— S— S-P-SH
IS-P-S-S-P— SI
+ 2 I2 + 3 HjO
IS-P— SO + bzS-P-SI
where P represents that portion of the protein not reacting with iodine.
Further oxidation of the sulfenate group to sulfinate may occur to give a
homogeneous product. Therefore the formation of sulfenyl iodide groups
and oxidation of SH groups to sulfenate and sulfinate must be considered
as likely possibilities in enzymes treated with iodine.
Inhibition of Enzymes
Many enzymes have been found to be readily inhibited, often by low con-
centrations of iodine (Table 5-4). It is impossible to know in most cases
whether the inhibition is due to reaction with SH groups or to iodination
of tyrosine. The fact that most studies have been done at pH's around
neutrality implies that both SH and tyrosyl groups could be reacted, so
that the relative importance would depend on the accessibility of the groups
and their location with respect to the active center. Fixation of iodine into
an enzyme does not imply inhibition; an example is Aspergillus protease
(Dhar and Bose, 1962). The inhibition of certain enzymes by iodine is
probably related to oxidation of SH groups: papain, creatine kinase, urease,
aldolase, lactate dehydrogenase, succinate dehydrogenase, pyruvate decar-
boxylase, and adenosinetriphosphatase. Other enzymes, such as pepsin or
peroxidase, are inhibited through tyrosine iodination, and in some instances
a mixed mechanism is probable.
One way of determining if SH group oxidation is responsible for enzyme
inhibition is to attempt reversal with thiols. Complete reversal certainly
implies such a mechanism, but negative results can be interpreted in various
ways. Even oxidation to disulfide groups is not necessarily reversed by
thiols if steric factors prevent reaction, and oxidation past the disulfide
IODINE 683
stage would not be expected to be reversed. No reactivation of /3-galactosi-
dase (Knopfmacher and Salle, 1941 ) or a-amylase (Di Carlo and Redfern,
1947) is observed; however, in both cases there is some reason for believing
that SH groups are involved. Partial reactivation of /5-amylase (Weill and
Caldwell, 1945 b) and phosphoglyceraldehyde dehydrogenase (Rapkine,
1938) was taken to mean that at least SH group oxidation is responsible
for the inhibition. Essentially complete reactivation with thiols has been
found for urease (Hellerman, 1939), papain (Hellerman and Perkins, 1934),
cholinesterase (Nachmansohn and Lederer, 1939), and lactate dehydrogen-
ase (Nygaard, 1955) so that an SH mechanism seems assured for these.
The mechanism of the inhibition of /5-fructofuranosidase is still unknown,
although it is the first enzyme studied with iodine. The enzyme is inhibit-
ed fairly rapidly to the extent of 45 50%, but further inactivation proceeds
very slowly (Myrback, 1926). A "Jodsaccharase" was assumed, but the
iodine must not be fixed at the active center since there is no decrease in
the affinity for the substrate. There is no reactivation by reduction (Myr-
back, 1957 a) so there is little evidence for SH group oxidation. Sulfenyl
iodide groups may be involved.
There has been little study of the disappearance of SH groups during in-
hibition by iodine. Cardiac lactate dehydrogenase SH groups are rapidly
oxidized by iodine, as determined with Ag+ and spectrophotometrically with
p-mercuribenzoate, and the inhibition develops in parallel fashion (Nygaard,
1956). The lactate dehydrogenase from rabbit muscle, on the other hand,
incorporates iodine at 0^ and pH 8 over many hours; when 1 atom of iodine
is incorporated per molecule of enzyme, the inhibition is 30%, and the in-
hibition increases until 21 atoms of iodine are incorporated. Both NAD and
oxalate protect the enzyme against iodination. Although the results with
iodoacetamide indicate an SH group at the active site, one cannot be cer-
tain if this is the initial point of attack for the inhibition (Dube et al., 1963).
It is likely in situations like this that both SH group oxidation and iodina-
tion of tyrosine occur.
Pepsin is not an SH enzyme but is inhibited by iodine, and here it is
highly probable that tyrosine iodination occurs. The activity of pepsin de-
creases with the amount of iodine incorporated; it is inactive when 35-40
atoms of iodine are bound (Herriott, 1937). 3-Iodotyrosine has been isolat-
ed from inhibited pepsin (Herriott, 1947), confirming the importance of
tyrosine for the enzyme activity. Of the 6 tyrosyl groups in ribonuclease,
3 are unreactive, and the problem of where these are in the polypeptide
chain was studied with iodine (Cha and Scheraga, 1961 a,b). At pH 9.4 and
10° — conditions favorable for tyrosine iodination with minimal effects on
other groups — 3 tyrosyl residues are iodinated; the others can be iodinated
only very slowly. The iodinated tyrosyl residues were located in the amino
acid sequence. Such techniques will undoubtedly become more common
684
5. OXIDANTS
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IODINE 685
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IODINE 687
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688 5. OXIDANTS
when more enzymes are susceptible to sequential analysis. Another interest-
ing approach to elucidating enzyme binding groups with iodine is illustrat-
ed by the study of the old yellow enzyme by Theorell (1956). Flavin is
bound to the apoenzyme through its imino group and fluorescence is quen-
ched; Weber had suggested that a tyrosine hydroxyl group might bind this
imino group. This was examined by reaction of the apoenzyme with iodine;
since no SH groups are present, this is relatively easy. It was found that
very low concentrations of iodine decrease the coupling rate of FMN to the
apoenzyme, and 90% of the iodine which disappears is recovered as diiodo-
tyrosine.
Some results on the variation of inhibition with pH appear to point to
the importance of tyrosine iodination. The iodination of pepsin is very slow
below pH 4.5 and rises suddenly as the pH is increased to become maximal
around pH 5.5 (Herriott, 1937). This is essentially the same pH dependence
as found for glycyltyrosine. The inhibition of /3-fructofuranosidase by iodine
is minimal at pH 5.14 and much faster at pH's above 6 (Myrback, 1926),
which might support the importance of tyrosine iodination in the inhibition.
Although one might expect the effect of iodine on cathepsins to be mainly
through reaction with SH groups, Maver and Thompson (1946) found
greater inhibition by 0.25 mM iodine at pH 7 (71%) than at pH 3.5 (20%).
Results with various SH reagents do not favor the importance of SH
groups. This may well be a case where there is a mixed mechanism for the
inhibition.
An interesting situation occurs in the reaction of the exopenicillinase of
B. cereus with iodine (Citri and Garber, 1960, 1961). This enzyme can exist
in two antigenically different states — a and y — and these differ in re-
sponse to iodine, although the enzyme activity is the same for both. a-Pen-
icillinase is quite resistant to iodine whereas ^-penicillinase is inhibited
by 0.5-1 mM iodine. The enzyme must be flexible since in the presence of
the competitive inhibitor, 6-(2,6-dimethoxybenzamido)penicillanic acid, it
becomes sensitive to iodine. Pretreatment with this inhibitor, followed by
its removal, does not alter hydrolysis of benzylpenicillin, so that any struc-
tural change that occurs is not permanent. It is very difficult to understand
how a rather large competitive inhibitor, which must cover the active cen-
ter, could allow reaction of any group at the active center with iodine, un-
less the reacted group is just vicinal to the active center and either SH oxi-
dation or iodination alters the structure.
Effects on Cellular Metabolism and Function
The uncoupling of oxidative phosphorylation by iodine has been claimed
to relate to the effects of thyroxine. Klemperer (1955) reported that al-
though iodide exhibits no uncoupling activity, iodine is quite effective in
rat liver mitochondria with /5-hydroxybutyrate as substrate (see accom-
IODINE 689
panying tabulation). Iodine appears to fulfill the requirements of an un-
coupler, in that it can reduce the P:0 ratio significantly without depressing
the respiration, although, to completely uncouple, the O2 uptake must be
KI
(mM)
I2
{mM)
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inhibited. Middlebrook and Szent-Gyorgyi (1955) found an uncoupling in
mitochondria when Cl~ is partially replaced with I^; at 25 mM I~, phos-
phorylation is almost abolished without depression of respiration. It is not
known if this is due to I" itself, to reduction in CI", or to iodine formed
from I~. Iodine causes mitochondrial swelling at a concentration as low as
0.005 raM, and in this it resembles thyroxine (Rail et al., 1962). Iodine is
able to oxidize NADH but addition of NADH does not reverse the swelling.
Other oxidizing agents do not duplicate this effect. Furthermore, agents that
inhibit thyroxine-induced swelling also inhibit that caused by iodine. De-
spite the superficial similarities in the actions of iodine and thyroxine, it is
difiicult to understand the nature of any relationship. It is very unlikely
that thyroxine releases its iodine, and since thyroxine is always more potent
than iodine, not enough iodine could be released in any event. It is possible
that iodine does not act directly, but iodinates tyrosine or some protein,
and that this product is the active uncoupler.
One might expect iodine to be an effective inhibitor of glycolysis, inas-
much as this pathway involves a number of SH-dependent steps. Yeast
fermentation is indeed quite sensitive to iodine, 0.017 mM inhibiting 22%
and 0.085 roM inhibiting 100% (Schroeder et al, 1933 b). There is simul-
taneously a loss of GSH and probably other SH groups. This does not
prove that the glycolytic inhibition is related to SH groups, but is sugges-
tive. The locus of action is not known; one thinks of aldolase, because of
its great sensitivity to iodine, but most of the glycolytic enzymes have not
been tested. That iodine can oxidize SH groups in cells was shown by Ca-
fruny et al. (1955 a). Kidney sections incubated with iodine exhibit 85%
loss in SH groups in the proximal and distal tubules. However, so little
work has been done on cell metabolism with iodine that it is impossible to
predict if any pathways are inhibited selectively; it would appear to be
unlikely, unless glycolysis proves to be more susceptible than other systems.
690 5. OXIDANTS
Iodine can interfere with the transport of substances across cell mem-
branes. Hemolysis by glycerol and other nonelectrolytes is quite potently
inhibited by iodine at 0.08 mM (LeFevre, 1947, 1948) as it is by various
SH reagents. The effects are readily reversed by thiols. This was taken to
indicate that SH groups are in some manner involved in the transport of
these substances into the erythrocytes. However, it is not necessarily evi-
dence for an active transport, since membrane permeability could be af-
fected directly or indirectly. Iodine also inhibits the transport of phosphate
into staphylococci and it was claimed that this process involves SH groups
(Mitchell, 1954). Finally, iodine at 0.1 mM reduces the short-circuit current
and electrical potential of frog skin (Eubank et al., 1962), but there is no
evidence as to the site or mechanism of this action.
Iodine has been studied a great deal in connection with its germicidal
activity (Gershenfeld and Witlin, 1950) but not a great deal has been done
from the metabolic standpoint. The effects of pH on the ability of iodine
to kill bacteria, fungi, or spores are, however, of interest, since they would
presumably apply to work with any cells. It has generally been considered
that at lower pH's there is more free iodine, and hence greater penetrability
into cells and greater activity. It is true that more iodate would be formed
in alkaline solutions and, in the absence of much iodide, more hypoiodous
acid. Wyss and Strandskov (1945) found the bactericidal activity to de-
crease at higher pH's and attributed this to a greater formation of HOI
and lOg". When iodide is present, the formation of HOI is suppressed, and
the pH does not affect the activity. It was also observed, as would be ex-
pected, that the action of iodine is strongly dependent on temperature, re-
quiring about 4 times as long to kill Bacillus metiens spores for each 10°
drop in temperature.
PEROXIDES
Hydrogen peroxide and other peroxides occasionally depress enzymes and
metabolism potently but little is known about the specificity with respect
to SH groups. In comparison with other oxidants, no thorough studies of
the effects of hydrogen peroxide on proteins have been made. Mirsky and
Anson (1935) mention that hydrogen peroxide is convenient to use in the
oxidation of SH groups, but it has never been widely applied for this pur-
pose. The interesting effects of hydrogen peroxide on glycolysis and a few
enzymes justify a brief discussion.
Chemistry
Hydrogen peroxide is a nonlinear molecule that is quite miscible with
water:
H2O2 + H2O -? H3O+ + OOH- K = 2Ax 10-1''
PEROXIDES 691
The ion product (H+) (00H-) is 1.55 X 10-^2 at 20°. Thus it is a very weak
acid and the ion OOH" is probably unimportant in its reactions. Hydrogen
peroxide can function as both oxidant and reductant. It is a strong oxidiz-
ing agent in both acid and alkaline media, but a relatively poor reductant.
Although the oxidation-reduction potential would be more favorable for
oxidation in acid medium, the rate of oxidation is often greater in alkaline
conditions. Hydrogen peroxide, of course, is an unstable substance, especi-
ally in the presence of organic material, and this must be considered in its
use. Despite its instability ( — AF = 23.4 kcal/mole), it is rather stable in
pure solution, but its decomposition is catalyzed by heavy metal ions, and
is more rapid in alkaline than acid media.
Inhibition of Enzymes
The few results summarized in Table 5-5 are not comparable with each
other because the conditions were quite different in the various studies.
However, there is no doubt that some enzymes are very sensitive to hydro-
gen peroxide. The inhibition develops very slowly in some cases; with yeast
/5-fructofuranosidase the inhibition by 2.9 M hydrogen peroxide is 0% at 1
min, 10% at 30 min, 47% at 3 hr, and 100% at 21 hr (Myrback, 1957 b).
Of course, at this very high concentration one has no idea of the mechanism
of the inhibition, and can only marvel at the resistance of this enzyme. The
inhibition of ATPase depends on the pH at which the reaction is run: Thus
the enzyme was incubated with hydrogen peroxide at pH 7 for 15 min, and
the inhibition was found to be 51% when the ATPase reaction was tested
at pH 6.3 and 95% when tested at 9.2 (Mehl, 1944). The reason for this
strange behavior is unknown. One factor that has not been generally consid-
ered is the possible presence of heavy metal ions in the hydrogen peroxide.
Holmberg (1939) believed that the inhibition of uricase he observed was
due to traces of Cu++, inasmuch as diethyldithiocarbamate prevents the
inhibition. It is also possible that some metal ion may be necessary to
catalyze the oxidation of the enzyme and that the inhibition is not due
to the Cu++ itself.
The inhibition of /5-galactosidase by hydrogen peroxide is completely re-
versible by HgS or cyanide, while that by iodine is not, indicating that here
one may oxidize the SH groups more specifically with the peroxide (Knopf-
macher and SaUe, 1941). Reactivation of ATPase inhibited by hydrogen
peroxide was observed by both Mehl (1944) and Ziff (1944), using cysteine
or glutathione, so that specific oxidation of SH groups may occur with this
enzjTne. Simultaneously there is a suppression of the interaction of actin
and myosin, which is believed to depend on SH groups (Bailey and Perry,
1947). Papain inhibited up to 90% by hydrogen peroxide can also be reac-
tivated by cysteine, but beyond this there is apparently oxidation beyond
the disulfide stage (Sanner and Pihl, 1963). Blocking the SH groups with
692
5. OXIDANTS
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694 5. OXIDANTS
p-mercuribenzoate prevents their reaction with hydrogen peroxide. The
single SH group of papain is about 5 times as susceptible to oxidation by
hydrogen peroxide as the SH group of phosphoglyceraldehyde dehydrogen-
ase. a-Chymotrypsin has two methionine residues, one being 3 residues
away from the active site serine and the other 15 residues removed. The
Met-3 is oxidized by hydrogen peroxide specifically, whereas both SH groups
are oxidized in the urea-denatured enzyme (Schachter et at., 1963). It is
obvious that disulfide bonds cannot be formed intramolecularly in the na-
tive enzyme and it was shown that methionine sulfoxide is the product.
Glutathione is also oxidized to the sulfoxide by hydrogen peroxide (Utzin-
ger et al., 1963).
Hydrogen peroxide generated during oxidation in enzyme preparations
or cells is sometimes inhibitory and for many years it has been assumed
that at least one function of catalase is to protect cells against it. Dixon
(1925) was the first to demonstrate this with a purified enzyme system.
When purines are oxidized by oxygen in the presence of xanthine oxidase
there is a progressive inactivation of the enzyme, which is due to hydrogen
peroxide formed, since it can be prevented by catalase. Hydrogen peroxide
was claimed to stimulate xanthine oxidase at very low concentrations (be-
tween 0.00001 mM and 0.01 mM) (Table 5-5), and to inhibit above 0.1 mM
(Bernheim and Dixon, 1928). The rate of activation is slow, maximal effects
of 0.001 mM hydrogen peroxide occurring in around 100 min. The mecha-
nism for this is unknown; metal impurities seem unlikely because they would
be at extremely low concentrations. The inhibition of Aspergillus aconitase
depends on the strain of the organism, the sensitivity varying over at least
a 4-fold range of concentration (Bruchmann, 1961 a). Certain strains tend
to accumulate citrate under specified conditions and this is believed to be
due to the hydrogen peroxide formed and the particular sensitivity of aco-
nitase in these strains (Bruchmann, 1961 b). Adding exogenous hydrogen
peroxide augments the accumulation of citrate (Bruchmann, 1961 c).
Succinyl peroxide is a radiomimetic substance and has been found to
inhibit several SH enzymes while having much less action on non-SH en-
zymes (Wills, 1959). The enzymes inhibited are amylase, /5-fructofuranosi-
dase, urease, succinate dehydrogenase, phosphoglyceraldehyde dehydrogen-
ase, papain, tyrosinase, and cholinesterase. Urease, for example, is inhib-
ited completely in 8 min by 0.033 mM succinyl peroxide. Reversal of inhi-
bitions by cysteine is obtained only if the period of exposure to the peroxide
is brief. It is questionable if the action is exerted by succinyl peroxide itself,
since it is immediately hydrolyzed to persuccinate in aqueous solution, and
peracids have long been known to oxidize SH groups (Freudenberg and
Eyer, 1932; Swan, 1959). The inhibition of catalase by monoethyl peroxide
is probably not due to SH group oxidation but to an analog type of inhi-
bition (Blaschko, 1935).
PEROXIDES 695
Effects on Metabolism
Hydrogen peroxide inhibits brain respiration and especially the oxidation
of succinate (Dickens, 1946 a). If one compares the effects on various tis-
sues, the sensitivity to hydrogen peroxide depends on the relative concentra-
tions of catalase; the more catalase, the less inhibition. In brain, 75 roM hy-
drogen peroxide inhibits respiration 36% and succinate oxidation 95% over
a 60 min period. It was thought initially that the toxic effects of high
oxygen tension on brain might be due to hydrogen peroxide released, but
this was shown not to be true.
If Lactobacillus is grown anaerobically, the cells lose their iron enzymes
and catalase; if they are then exposed to oxygen, hydrogen peroxide is form-
ed and the cells are killed (Warburg et al., 1957). Since cancer cells possess
an anaerobic type of metabolism and contain much less catalase than nor-
mal cells, it was postulated that this may be the cause of the greater sensi-
tivity of cancer cells to hydrogen peroxide. It was found that 1 mM hy-
drogen peroxide has no effect on the aerobic or anaerobic glycolysis of em-
bryo tissue, but inhibits both almost completely in ascites cells. Since the
catalactic activity of embryo tissue is around 10-fold that of the ascites
cells, this could explain the differential susceptibility. Inasmuch as radia-
tion of cells can induce hydrogen peroxide formation, this may be one reason
for the more selective effects of radiation on cancer cells. Holzer and Frank
(1958) extended these observations in ascites cells to show that hydrogen
peroxide at 0.056 mM not only inhibits glycolysis 86%, but simultaneously
reduces the NAD concentration very markedly (0.31 to 0.05 //moles/ml).
Triose-P and fructose-diP rise, indicating a block of the phosphoglyceral-
dehyde dehydrogenase. However, they found the extracted enzyme to be
inhibited only 37% by 0.079 mM hydrogen peroxide, so that concentra-
tions effectively blocking glycolysis would have little effect on this enzyme
(assuming the same sensitivities of the intact and extracted enzymes). They
thus postulated that the inhibition is due to a reduction of NAD and that
this suppresses the oxidation of triose-P. Nicotinamide can protect both
NAD and glycolysis from hydrogen peroxide, and Pantlitschko and Seelich
(1960) showed that it could overcome the inhibition when added 1 hr after
the hydrogen peroxide. Baker and Wilson (1963) confirmed the inhibition
of anaerobic glycolysis in Ehrlich ascites carcinoma ceUs, although the ef-
fects were not as marked as observed previously — some inhibition at 0.3
mM, around 50% at 0.9 mM, and 80% at 2 mM — and further showed
that during the oxidation of unsaturated fatty acids some hydrogen per-
oxide is formed and may depress glycolysis. Piitter (1961) studied the
possible relationship between glycolysis and transplantability of ascites
cells, but encountered the difficulty that the hydrogen peroxide used to
inhibit glycolysis is fairly rapidly decomposed so that the inhibition dis-
appears. Thus it requires above 1 mM to interfere with transplantability.
696 5. OXIDANTS
Hydrogen peroxide is by no means the ideal glycolytic inhibitor for this
type of work.
Effects on Tissue Function and in Whole Animals
The spontaneous motility of the rat intestine is extremely sensitive to
hydrogen peroxide inasmuch as 10% stimulation of the amplitude occurs
with 0.00057 mikf (Goodman and Hiatt, 1964). At 0.057 mM hydrogen per-
oxide the stimulation of contraction by acetylcholine is blocked and 67%
of the total SH groups of the tissue are reacted. Although hydrogen per-
oxide at 0.001 vaM has no definite effect on the spontaneous contractility,
it reduces the effect of acetylcholine somewhat. Other SH reagents act sim-
ilarly and it appears that the response to acetylcholine is dependent on
SH groups. The contractility of nonconducting rabbit psoas muscle is block-
ed by 300 mM hydrogen peroxide after 7 min exposure, and this is not
reversible with cysteine (Korey, 1950). However, little can be learned from
concentrations of this magnitude.
An interesting relationship was discovered by Feinstein et al. (1954), in
that a sublethal dose of iodoacetate (20 mg/kg) and a 20% fatal dose of
hydrogen peroxide (15 meq/kg) given together kill all the animals. Inas-
much as iodoacetate also potentiates the lethality of X-irradiation in mice,
this was considered as evidence that radiation may produce some of its
effects by the release of hydrogen peroxide. It was noted that the toxicity
of hydrogen peroxide is markedly increased by treating the animals with
azide, a catalase inhibitor; however, hydroxylamine, which is a better cat-
alase inhibitor, does not augment the effects of hydrogen peroxide.
TETRATHIONATE
Tetrathionate appears to be a fairly specific oxidant for SH groups under
the proper conditions, but has been used very little in enzyme work. It was
found to be capable of antagonizing cyanide poisoning in dogs at doses of
500 mg/kg (Chen et al., 1934), and today we might interpret this as due to
methemoglobin formation. Tetrathionate has been used in a method for
the determination of protein methionine, which is demethylated to homo-
cysteine and then oxidized (Baernstein, 1936). It has been used clinically
in thromboangiitis obliterans, supposedly for an effect on the blood, an in-
crease in the oxygen capacity being observed (Theis and Freeland, 1940).
It is surprising that the blood glutathione increases after injection of tetra-
thionate. It has been applied occasionally to the reduction of the cytochrome
components of the respiratory chain, since the initial work of Keilin and
Hartree (1940), the tetrathionate apparently being oxidized to sulfite. It is
thus, like hydrogen peroxide, both an oxidant and a reductant, which makes
TETRATHIONATE 69?
its effects in complex systems more difficult to interpret. The first use of
tetrathionate as a reagent for protein SH groups was by Anson (1941), who
demonstrated that it would titrate denatured ovalbumin, although the reac-
tion is slower than with ferricyanide, porphyrindin, or p-chloromercuriben-
zoate, not being complete in 3 min at neutrality. It has not been used ex-
tensively for this purpose, but it may well be a valuable reagent in certain
types of work; a detailed description of the method is given by Chinard
and HeUerman (1954).
Chemistry and Reaction with SH Groups
Sodium tetrathionate is prepared from the thiosulfate by oxidation with
iodine in 90% ethanol. The precipitate is purified by redissolving it in an
equal weight of water and filtering it into absolute ethanol, in which it re-
precipitates. It is washed with ethanol and dried in vacuo. Sodium tetra-
thionate crystallizes with 2 waters of hydration. When it is kept at 0° in
the dark, both the solid and 0.1 M solutions are stable for many weeks
(Pollock and Knox, 1943), but it is unstable when kept under ordinary
conditions. For all accurate work it is necessary to be certain that it is free
of appreciable thiosulfate and other impurities, and it should be recrystal-
lized as above.
Tetrathionate rapidly oxidizes simple thiols, such as cysteine, homocys-
teine, and glutathione, according to the reaction:
2 R— SH + S^Og" ^ R— S— S— R + 2 S^Og" + 2 H+
In titrations of SH groups the thiosulfate is determined iodometricaUy.
According to Baernstein (1936), it is specific for SH groups and does not
react with other amino acid groups. It is not a strong oxidant, since the
standard oxidation-reduction potential for the reaction
2 §203= ±? 8406= + 2 e-
is + 0.08 V. Although it has always been assumed that tetrathionate oxi-
dizes SH groups to disulfide, Pihl and Lange (1962) have obtained evidence
that sulfenyl thiosulfate groups may be formed:
R— SH + 8406= -> R— 8— 8^03= + 8^03= + H+
Incubation of phosphoglyceraldehyde dehydrogenase with tetrathionate-S^^
leads to the appearance of S^^ bound to the protein, and the binding of each
S^^ is associated with the disappearance of one SH group.
One of the few thorough kinetic studies of SH group oxidants was made
by Goffart and Fischer (1948). It was shown that tetrathionate oxidizes
protein SH groups more slowly than cysteine or glutathione (Fig. 5-1).
698
5. OXIDANTS
Furthermore, the reaction is initially rapid but in most instances slows
down suddenly, which is difficult to explain for the simple thiols. The oxi-
dation proceeds much faster at pH 7 than at pH 5 (see accompanying
tabulation).
Time for complete reaction
Protein
(min)
pH 5 pH 7
Lens protein
140 30
Ovalbumin (denatured)
120 12
Myosin (denatured)
140 50
06
0 5
GSH ^_______
-rSTEINt
0.4
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0.3
OVAL BUM
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01
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Fig. 5-1. Rates of reaction of
10 mil/ tetrathionate with the
SH groups of thiols and proteins
at pH 5. (From Goffart and
Fischer, 1948.)
Inhibition of Enzymes and Metabolism
Succinate dehydrogenase is inhibited around 90% by 0.1 mM tetrathi-
onate (Keilin and Hartree, 1940). This is an effect on the dehydrogenase SH
groups according to these authors and PhiHps et al. (1947), who confirmed
the inhibition on succinate dehydrogenases from several tissues. No inhi-
bition of ascorbate oxidation, and hence of the cytochrome system, is ob-
served even with 10 mM. Succinate protects the enzyme; when tetrathionate
is 0.5 mM, succinate reduces the inhibition from 96% to 39%, and when
it is 0.1 mM from 79% to 10% (pigeon breast enzyme). The inhibition is
only partially reversible with glutathione or cysteine. In work with suc-
cinate dehydrogenase it may be well to consider the possibility that some
of the inhibition results from a competitive action of the tetrathionate, since
it has negative charges appropriately separated. Choline dehydrogenase
from rat liver is also quite sensitive to tetrathionate, 38% inhibition re-
TETRATHIONATE 699
suiting from 0.2 mM and 93% from 0.6 mM (Gordon and Quastel, 1948).
The only clear-cut demonstration of reaction with enzyme SH groups is
that of phosphoglyceraldehyde dehydrogenase (Pihl and Lange, 1962). Here
tetrathionate inhibits as well as /)-chloromercuribenzoate, i.e., when 3 moles
of inhibitor are reacted per mole of enzyme, the activity is reduced to zero
in both cases. Also the inhibition is fully reversible with thiols. However,
as mentioned above, the reaction does not appear to be a simple oxidation,
but involves the formation of sulfenyl thiosulfate groups. The enzyme is
very sensitive, since 0.005 mM inhibits completely (enzyme = 0.0005 mM)
within 5 min.
In view of the potent inhibition of phosphoglyceraldehyde dehydrogenase
one might anticipate tetrathionate to be a glycolytic inhibitor. Goffart and
Fischer (1948) attempted to demonstrate a Lundsgaard effect in muscle,
i.e., a typical contracture such as produced by iodoacetate and certain other
SH reagents. Following injection into rabbits, the extremities become weak
but the muscles remain elastic and the reflexes normal; if the gastrocne-
mius is stimulated, it does not go into contracture. Intraarterial injection
produces a temporary contracture (or at least some inhibition of relaxation).
Injection into frogs does not give an iodoacetate-like effect and the isolated
frog rectus abdominis muscle gives only a temporary contracture-like reac-
tion. It is doubtful if true contractures are observed, and in any case the
tetrathionate concentration must be quite high. It is possible that the phos-
phoglyceraldehyde dehydrogenase in the muscle is protected by a perme-
ability barrier to the doubly charged inhibitor, and by the presence of NAD
and substrate on the enzyme. MacLeod (1951) found inhibition of glycolysis
in human spermatozoa, but the tetrathionate concentration was 10 mM and
the inhibition progressed very slowly, so that even after 3 hr the glycolysis
is not completely depressed (around 50%). The motility decreases simul-
taneously with the reduction in glycolysis. One of the pitfalls of reversibility
experiments is well illustrated here, for when cysteine is used there is a rapid
toxic effect on the spermatozoa, this being due to the cystine arising as the
result of the oxidation by tetrathionate.
Tetrathionate is reduced to thiosulfate by reaction with SH groups and
it has been supposed that the rapid conversion into thiosulfate in rabbits
and dogs is due to this (Gilman et al., 1946). While this must be true in part,
there is some evidence for enzyme systems catalyzing this reaction. Thus in
various bacteria tetrathionate is readily reduced, while in others no reaction
at all occurs (Pollock and Knox, 1943). Postgate (1956) has isolated cell-free
systems reducing tetrathionate, thiosulfate, and sulfite from the anaerobe
Desulfovibrio desulfuricans, the cytochrome system acting as an electron
carrier for the tetrathionate reductase. Indeed, it is likely that tetrathio-
nate can be oxidized through the cytochrome system in most cells. Thus
tetrathionate must generally be rather labile in most biological situations.
700 5. OXIDANTS
Nephrotoxic Action
Intravenous injection of around 0.5 g/kg of sodium tetrathionate into
dogs leads to the development of anuria within 30-60 min, a rapid reduc-
tion in creatinine clearance, and the appearance of proximal tubular ne-
crosis (Gilman et al., 1946). At the time of death there is no evidence of
toxicity, symptomatic or histological, in any other tissue from such min-
imally lethal doses; higher doses, however, can cause a long-lasting ataxia,
and in rabbits some difficulty in muscular relaxation. Inasmuch as simul-
taneous reduction of the tetrathionate to thiosulfate occurs, it was assumed
that the renal damage is related to the oxidation of SH groups, the toxicity
thus being related to that produced by the mercurials. Nevertheless, nephro-
toxic doses in rabbits do not inhibit kidney succinate dehydrogenase at aU
in vivo, and yet this enzyme is very sensitive to tetrathionate (Philips et al.,
1947). Large doses of tetrathionate (1 g/kg of the sodium salt) in rabbits
lead to a 77% loss of glutathione in the kidneys, 28% in the blood, 30%
in the liver, and 20% in muscle after 90 min, most of the change occurring
within 30 min (Goffart and Fischer, 1948). These results might indicate that
tetrathionate has a greater effect on renal SH groups than those of other
tissues, without implying that the toxicity is due to the loss of glutathione.
CHAPTER 6
o-IODOSOBENZOATE
The most commonly used oxidant for enzyme SH groups at present is
o-iodosobenzoate because it is probably the most selective for these groups.
For this reason it deserves a somewhat more detailed treatment than the
other oxidants and a separate chapter. o-Iodosobenzoate was first prepared
by Meyer and Wachter (1892) and studied biologically by Heinz (1899) in
Germany. The early interest stemmed from the use of iodine and organic
iodine compounds in superficial infections. Indeed, Heinz was mainly con-
cerned with the administration of sodium iodide and o-iodosobenzoate to-
gether so that by the oxidation of the iodide it would be possible to form
"nascent" iodine in the tissues. Consequently there w^ere early investiga-
tions on the antibacterial activity (Jahn, 1914) and the effects on phagocy-
tosis (Arkin, 1912). The initial pharmacological study was by Loevenhart
and Grove (1909, 1911) at the University of Wisconsin, but the results did
not engender much clinical enthusiasm and, inasmuch as the actions at
that time were not related to any metabolic site of attack, o-iodosobenzoate
was little used by biochemists until it was introduced by Hellerman et al.
(1941) for the estimation of protein SH groups. The application to enzyme
characterization was slow but during the past several years it has come to
be one of the most useful SH reagents. It differs from the arsenicals, the
mercurials, and the alkylating agents in not introducing new groups or side
chains onto the enzymes, since it is generally believed that the primary
action is an oxidation of the SH groups to the disulfide state. However, the
use of o-iodosobenzoate, like most SH reagents, in complex systems or cel-
lular preparations is limited because of the number of components affected
and the inherent difficulty in the interpretation of the results.
CHEMISTRY
The structures of the different oxidation states of the iodinated benzoates
may be written as:
701
702
coo
o-Iodobenzoate
6. 0-IODOSOBENZOATE
coo'
I— O"
o-Iodosobenzoate
COO
o-lodoxybenzoate
It is possible that the aryl iodoso compounds, cp — 1+ — 0~, can add a proton
to form 9? — 1+ — OH since the corresponding protonated iodoxybenzene is
known, but the ionization constant is unknown. Indeed, the carboxyl p^^
is not accurately known, but is probably around 6.0 to 6.5, in contrast to
the piiig for o-iodobenzoic acid (2.86), and o-iodosobenzoic acid may be
precipitated from solution by passing COg through solutions of the sodium
salt. o-Iodosobenzoic acid may be easily prepared by oxidation of o-iodo-
benzoic acid with permanganate, crystallization by cooling, and recrystal-
lization with COg, and can be determined iodimetrically (Loevenhart and
Grove, 1911; Chinard and HeUerman, 1954). Commercial samples should
probably be repurified for accurate work. The m- and p-iodosobenzoates
are also strong oxidizing agents and might possibly have certain advantages
over o-iodosobenzoate for particular purposes, but they have been almost
completely ignored. It is interesting that p-iodosobenzoate reacts like o-iodo-
sobenzoate with the SH groups of L-glutamate dehydrogenase, but no de-
tailed comparison was made (Hellerman et al., 1958). The o-iodoxybenzoate
is also a potentially useful reagent but essentially nothing is known of its
actions on proteins or enzymes.
The oxidation of SH groups by o-iodosobenzoate results in the formation
of o-iodobenzoate. Whether the disulfide link is intra- or intermolecular de-
pends on the thiol reacted; with cysteine or glutathione it is obviously in-
termolecular, but what evidence exists for proteins suggests that intramo-
lecular oxidation is dominant. Oxidation apparently does not proceed be-
yond the disulfide stage:
COO
COO
I— O
SH
SH
+ R
/'
+ H,0
at pH 7, and under proper conditions accurate titration of SH groups can
be achieved. However, at lower pH's or in the presence of excess of o-iodo-
sobenzoate, further oxidation to the sulfinate or sulfonate stages may occur,
and groups other than SH may be attacked. Whether a free radical mecha-
nism is involved in the oxidation of SH groups here is not known, but if so,
one might postulate the following types of reaction:
REACTION WITH PROTEIN SH GROUPS 703
-S— S— R
R— SH ^R-S-= '"' > R— S— I — <)— coo"
Reaction (1) would involve combination with another R — S-, while reac-
tions (2) and (3) would involve additional o-iodosobenzoate. Compounds of
the type R — S — I — R' are generally unstable, but on proteins such link-
ages may occasionally be more stable, as in the formatiom of sulfenyl thio-
sulfates (page 697).
At neutrality, 25°, and 1-5 loM o-iodosobenzoate, only SH groups are
significantly oxidized; cystine, methionine, glucose, and ascorbate are not
oxidized appreciably under these conditions. However, ascorbate is oxi-
dized slowly by o-iodosobenzoate at pH 4.6 when both are present at 1 mM
in acetate buffer, half-reaction time being around 80 min (Caraway and
Hellerman, 1953). The nature of the buffer is important inasmuch as it is
an acid-catalyzed reaction. It is interesting that m- and p-iodosobenzoates
oxidize ascorbate almost instantaneously. NADH is not oxidized by o-iodo-
sobenzoate at pH 4.6 (Schellenberg and Hellerman, 1958). The oxidizability
of tyrosine phenolic groups by o-iodosobenzoate has not been thoroughly
examined but there is no evidence from work with proteins that this reac-
tion proceeds readily. The tyrosine groups of /5-amylase seem to be resistant
to o-iodosobenzoate since there is no change in absorption at 280 mju
(Englard et al, 1951).
REACTION WITH PROTEIN SH GROUPS
In order to titrate selectively protein SH groups with o-iodosobenzoate,
it is necessary to control the conditions carefuUy, as with all SH reagents.
It is usual in titrations with o-iodosobenzoate to add a slight excess of the
reagent and determine the amount not reduced by addition of KI and sub-
sequent titration of the released Ig with thiosulfate (Chinard and Heller-
man, 1954). The test is best run at pH 7 and between 15° and 25°. The
required reaction time varies with the protein tested but is usually less
than 30 min. Ovalbumin denatured with guanidine is titrated quite satis-
factorily and all of the SH groups are oxidized. Only a fraction of the SH
groups of native ovalbumin or other proteins is oxidized.
The specific oxidation of protein SH groups to disulfide may be accom-
panied by changes in the protein structure, which could have important
bearing on the mechanism of enzyme inhibition. Evidence for such struc-
tural alteration is given by the increased water-binding capacity of gels
formed from serum proteins previously treated with o-iodosobenzoate (Jen-
sen et al., 1950). At about equimolar ratios, o-iodosobenzoate changes the
704 6. O-IODOSOBENZOATE
nature of the clots from soft and opaque to firm, elastic, and almost trans-
parent, and simultaneously the water binding increases from 14.3 to 36.5
g/g. It is quite possible that more linear proteins, which may be reasonably
flexible, can be altered quite markedly by the formation of disulfide bonds,
and that particular regions on the surface may be made unavailable for
other reactions.
Further evidence for structural changes induced by o-iodosobenzoate is
the decrease in the viscosity of G-actin brought about by 2 mJf of the
reagent acting for 30 min at 25^ and pH 7.8-8 (Barany et al., 1962). This
is interpreted as an inhibition of polymerization. Simultaneously there is a
decrease in the ability to bind Ca++, as shown by the loss of G-actin-bound
Ca^^ upon tretment with o-iodosobenzoate. Although there appears to be
some correlation between changes in viscosity and Ca++ binding as pro-
duced by various SH reagents, the mechanisms involved are not yet under-
stood.
INHIBITION OF ENZYMES
Most SH enzymes are inhibited by o-iodosobenzoate (see Table 6.1) and
in a few instances the inhibition is very marked. Enzymes without SH
groups at or near their active centers, as shown by failure to be inhibited
by SH reagents in general, are not affected by o-iodosobenzoate, except
possibly in the single instance where it has been claimed to act as a com-
petitive inhibitor on D-amino acid oxidase, although in such cases the con-
centration would usually have to be a good deal higher than for the oxi-
dation of susceptible SH groups (Frisell et al., 1949) (see page 342). Inas-
much as o-iodosobenzoate is used up in the reaction, inhibition is of a titra-
tion type and spontaneously irreversible; thus the degree of inhibition will
often depend on the amount of enzyme present, or the amount of some SH
containing impurity that also reacts with the reagent. In most complex
systems, many enzymes will be inactivated to varying degrees, and probably
little specificity is possible. However, the remarkable sensitivity of creatine
kinase — definite inhibition at 0.00001 mM and complete inhibition at
0.00013 mM — might well make it possible to block this enzyme selectively
(Ennor and Rosenberg, 1954). This inhibition may explain the observation
of Bailey and Marsh (1952) that the fall in creatine phosphate in muscle
homogenates is almost completely prevented by o-iodosobenzoate. It might
be worthwhile to consider the use of o-iodosobenzoate in glycerinated and
similar muscle preparations to determine the role of creatine kinase in the
initiation of contraction or relaxation. Phosphoglyceraldehyde dehydro-
genase seems to be less sensitive to o-iodosobenzoate than to iodoacetate or
iodoacetamide, so that a specific block of glycolysis at this step would be
impossible. It may be noted that several dehydrogenases are quite well
inhibited by o-iodosobenzoate, notably the xanthine, malate, and aldehyde
INHIBITION OF ENZYMES
705
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INHIBITION OF ENZYMES
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714 6. O-IODOSOBENZOATE
dehydrogenases. At the present time, o-iodosobenzoate is more useful in the
study of pure enzymes as an indicator of SH groups than in cellular systems,
but has been little investigated in the latter and may possess potentialities
as a metabolic blocker if applied properly.
Titration of Enzyme SH Groups
Muscle phosphoglyceraldehyde dehydrogenase contains 11 cysteine resi-
dues and reacts rapidly with 1 1 moles of ^^-chloromercuribenzoate per mole
of enzyme. Segal and Boyer (1953) reported that 7.3-7.45 moles of o-iodo-
sobenzoate react with each mole of this enzyme, indicating 14.6-14.9 re-
ducing groups. Theoretically one would expect 5.5 moles of o-iodosobenzoate
to be reduced by each mole of enzyme, assuming that all the SH groups
are oxidized to the disulfide level. Segal and Boyer thus suggested that some
of the SH groups may be oxidized beyond the disulfide stage. Actually only
10 of the 11 SH groups could form intramolecular disulfide bonds, so the
extra SH group must either remain unoxidized, form a disulfide link with
another molecule of the enzyme (which is unlikely), or be oxidized to some
state other than the disulfide. Since oxidation to the S — 0~ or SOg" state
would require 2 molecules of o-iodosobenzoate for each SH group, 7 moles
of o-iodosobenzoate would react with each mole of enzyme if oxidation of
the extra SH group occurred in this way. Rafter (1957) investigated this
problem further and, under his conditions, found 10-11 moles of o-iodoso-
benzoate to react with each mole of the enzyme, indicating 20-22 reducing
equivalents. Furthermore, the o-iodosobenzoate-treated enzyme still pos-
sesses 30% of its initial SH groups, as determined by reaction with p-chloro-
mercuribenzoate. These results point to reaction of o-iodosobenzoate with
groups other than SH groups, or to oxidation of a fraction of the SH groups
beyond the disulfide stage. There is no evidence for reaction with other
groups and the enzyme is completely reactivated by cysteine. Thus one
might assume that 4 SH groups are oxidized to the disulfide level, 4 are
oxidized beyond this, and 3 remain unreacted; this would require 10 moles
of o-iodosobenzoate per mole of enzyme. Another possibility is that some
of the SH groups form E — S — I — cp — C00~ residues. It is interesting that
although o-iodosobenzoate abolishes the usual phosphoglyceraldehyde dehy-
drogenase activity and the arsenolysis reaction, it simultaneously increases
phosphatase activity 6-fold, this phosphatase activity being dependent on
NAD. The esterolytic activity with p-nitrophenylacetate as substrate is in-
hibited completely in 10 min when 4-5 moles of o-iodosobenzoate have react-
ed per mole of enzyme (Olson and Park, 1964). No substrate protection was
observed. Yeast phosphoglyceraldehyde dehydrogenase contains fewer SH
groups than the muscle enzyme — 2-4 per molecule — and reacts with 6
molecules of o-iodosobenzoate, so that here too an anomalous effect is seen.
Barron and Levine (1952) report 11.9 SH groups in yeast alcohol dehydro-
INHIBITION OF ENZYMES
715
genase by o-iodosobenzoate titration and 9.3 by an amperometric method,
and thus the value with o-iodosobenzoate is a little high in this case also.
Kinetics of Inhibition
The rate of oxidation of simple thiols by o-iodosobenzoate is usually quite
rapid, but protein or enzyme SH groups vary greatly in the rapidity with
which they react with this reagent. The inhibition of succinate oxidase by
o-iodosobenzoate at 0.2 uiM and 37° requires about 30 min to become max-
imal (Slater, 1949). This is shown in Fig. 1-12-12, where the maximal inhi-
bition of around 35% indicates that insufficient o-iodosobenzoate was pres-
ent for reaction with all of the enzyme and nonenzyme material. At 16° it
is evident that there are two phases, one complete within 10 min and
the other incomplete after 2 hr (Fig. 1-12-13). Although the first phase
undoubtedly represents oxidation of SH groups, it is not clear if the slow
reaction is further oxidation of other SH groups or secondary inactivation
of the enzyme.
The inhibition of ribonuclease by o-iodosobenzoate also shows two phases
(Figs. 6-1 and 6-2), one a fairly rapid reaction inhibiting around 20% and
Fig. 6-1. Rates of inhibition of ribo-
nuclease by o-iodosobenzoate at pH 7.
The times indicate the duration of the
incubation of the enzyme and inhib-
itor. (From Ledoux, 1954.)
the other a much slower one that is linear at least over 1-2 hr (Ledoux,
1954). The rate of inhibition during the second phase is dependent on the
concentration of o-iodosobenzoate and, since this phase starts from essen-
tially the same degree of inhibition for each concentration, it seems that
716
6. O-IODOSOBENZOATE
the slow phase is a further reaction with enzyme groups rather than a sec-
ondary inactivation. It has been emphasized several times with other en-
zymes, e.g. /3-amylase (Englard et al., 1951) and phosphoglucomutase (Mil-
stein, 1961), that the reaction with o-iodosobenzoate is slow. For this reason,
many of the results given in Table 6-1 are not comparable, since different
times of incubation with the inhibitor were used and usually the times were
not given. Unless preincubation of the enzyme with o-iodosobenzoate for a
reasonably long period (at least 30 min) is done, it is likely that the inhibi-
tions determined are partial and do not accurately represent the true effect
of the oxidant on all of the enzyme present.
Fig. 6-2. Rates of inhibition of ribonuclease by
1 raM o-iodosobenzoate at various pH's. The times
indicate the duration of the incubation of the en-
zyme and inhibitor. (From Ledoux, 1954.)
Effects of pH
It has been emphasized that in titrations of protein SH groups a pH near
neutrality must be maintained if specificity is desired; below a pH of 7 the
oxidizing power of o-iodosobenzoate increases, and oxidation of other pro-
tein groups may occur. Also the solubility of o-iodosobenzoate decreases
rapidly below pH 7. One might expect, therefore, that the inhibition of en-
zymes might decrease as the pH is raised above 7, but just the opposite
has been observed. /^-Amylase is inhibited more strongly and more repro-
ducibly at pH 7.8 than at pH 7 (Englard et al., 1951 ), succinate dehydrogen-
ase is inhibited more at pH 7 than at pH 6 (Stoppani et al., 1953), and
ribonuclease is inhibited more rapidly and completely as the pH is increased
from 7 to 10, no inhibition occurring at pH 5 (Fig. 6-2) (Ledoux, 1954). The
very rapid initial oxidation of ribonuclease at pH's above 8.5 may be due
INHIBITION OF ENZYMES 717
to the ionization of the SH groups. Penicillinase is inhibited 29% by 2 xnM
o-iodosobenzoate at pH 7.4 but not at all at pH 6 (Smith, 1963 b). On the
basis of these results, one might conclude that one should avoid pH's of
7 or below in enzyme work. Unfortunately, there are no data on the speci-
ficity of o-iodosobenzoate at higher pH's.
Protection of Enzymes against Inhibition by o-lodosobenzoate
Protection of an enzyme by addition of some thiol with the o-iodoso-
benzoate does not tell one anything about the mechanism of inhibition,
since the inhibitor is simply depleted by oxidation of the thiol. However,
protection by substances interacting with the enzyme active center pro-
vides some evidence for the site of the inhibition by o-iodosobenzoate. The
normal substrate of an enzyme has been shown frequently to protect against
o-iodosobenzoate, if it is present during the incubation of the enzyme with
the inhibitor. Thus, alcohol dehydrogenase is protected by ethanol (Barron
and Levine, 1952), fumarase by either fumarate or malate (Favelukes and
Stoppani, 1958), D-amino acid oxidase by alanine (Frisell and Hellerman,
1957), choline oxidase by choline (Rothschild et al., 1954), glutamate se-
mialdehyde reductase by glutamate semialdehyde (Smith and Greenberg,
1957), succinate oxidase by succinate (Thorn, 1959), and homogentisicase
by homogentisate (Tokuyama, 1959). In some instances the protection may
be very marked; e.g., fumarase is completely protected against 0.5 vaM
o-iodosobenzoate by 25 mM fumarate, and a 58% inhibition of alcohol dehy-
drogenase is reduced to 6.5% by ethanol. Coenzymes can likewise protect
in certain instances: aldehyde dehydrogenase is protected by NAD and
NADP (Stoppani and Milstein, 1957 a,b), alcohol dehydrogenase is protect-
ed by NAD (Barron and Levine, 1952), and D-amino acid oxidase is pro-
tected by FAD (Frisell and Hellerman, 1957). It is interesting that the dif-
ferent aldehyde dehydrogenases are protected to different degrees by their
coenzymes. The K+-activated yeast enzyme is protected by both NAD and
NADP, as well as by acetaldehyde, whereas the NAD-linked liver enzyme
is protected only by NAD and not by NADP. These observations may be
taken to mean that the inhibition is the result of a reaction of o-iodoso-
benzoate with SH groups at or near the active center, and that when the
active center is covered by substrate or coenzyme the oxidant is unable to
attack these SH groups. Such effects must be taken into account when
o-iodosobenzoate is used in cellular preparations. In one instance the effect
of substrate is abnormal. The lactate dehydrogenase of Propionibacterium
pentosaceum is inhibited more readily in the presence of lactate than in its
absence (see accompanying tabulation) (Molinari and Lara, 1960). This
was explained by assuming that lactate increases the fraction of free SH
groups, suggesting that the SH groups may be involved in the electron
transport.
718 6. O-IODOSOBENZOATE
o-Iodosobenzoate
%
Inhibition
(mM)
No lactate
Lactate 50 mM
1
5
30
2
2
36
5
36
54
Reactivation of o-lodosobenzoate Inhibition
If the only action of o-iodosobenzoate is the oxidation of SH groups to
disulfide bonds, one might expect some reversal of the inhibition by thiols,
and this has been observed with certain enzymes. The inhibition of D-amino
acid oxidase is reversed completely by cysteine (Rocca and Ghiretti, 1958)
but in most cases only partial reactivation is possible, for example, succinate
oxidase by dimercaprol (Thorn, 1959) and by glutathione (Slater, 1949),
amylo-l,6-glucosidase by glutathione (Earner and Schliselfeld, 1956), thre-
onine aldolase by dimercaprol (Karasek and Greenberg, 1957), and alcohol
dehydrogenase by glutathione (Barron and Levine, 1952). It is difficult to
interpret partial reactivation since failure to reverse the inhibition com-
pletely may be due to a variety of factors. No reactivation by thiols has
been reported for a few enzymes: Acid phosphatase cannot be reactivated
by cysteine or thioglycolate (Tsuboi and Hudson, 1955 b), nor xanthine
oxidase by cysteine (Harris and Hellerman, 1956), nor /^-amylase by gluta-
thione or dimercaprol (Englard et al., 1951), nor 5-hydroxytryptophan de-
carboxylase by thiols (Buzard and Nytch, 1957). However, these failures
cannot be immediately attributed to other actions of o-iodosobenzoate and
perhaps the most likely explanation is progressive secondary inactivation
consequent to the protein distortion induced by disulfide bond formation.
Some failures might also be due to attempting reactivation in the presence
of oxygen, which can often oxidize the thiols to disulfides, which in turn
can inhibit the enzyme, as pointed out by Slater (1949).
Variation of the Inhibition with the Substrate Used
The degree of inhibition of lipase by 1 mM o-iodosobenzoate varies with
the substrate (Singer, 1948; Singer and Hofstee, 1948 a). The inhibition by
p-chloromercuribenzoate is also dependent on the substrate used and Singer
postulated that the mercurial is bound near the substrate site so that it
sterically interferes with the binding of the substrates, the interference be-
ing greater the larger the substrate molecule. However, how this explanation
could apply to o-iodosobenzoate is not clear, since the simple formation of
INHIBITION OF ENZYMES 719
a disulfide link near the substrate site would not obviously produce a steric
effect. Inasmuch as the general problem of dependence of inhibition on the
substrate will come up several times with other inhibitors, it will be well to
suggest some of the possible mechanisms by which such effects can arise.
Substrate % Inhibition by o-iodosobenzoate (1 m.M)
Triacetin 50
Tripropionin 67
MonobutjTin 74
Tributyrin 89
(1) It is conceivable that the formation of a disulfide structure can dis-
tort the enzyme structure at or near the substrate site so that inhibition
will result, and this inhibition might not be the same for each substrate,
because of either steric factors or changes in the spatial position of the en-
zyme groups involved in the hydrolysis. (2) The SH reagent might not pri-
marily react with the enzyme but with the substrate, as suggested by Wills
(1960) for the inhibition of pacreatic lipase by p-chloromercuribenzoate.
Since this enzyme is not inhibited by o-iodosobenzoate, a relation with sub-
strate cannot be established. Wills believes that the mercurial is adsorbed
onto the glyceride-water interface and, in order to examine this possibility,
shook tributyrin with 10 nxM p-chloromercuribenzoate, washed it, and then
used this as a substrate; marked inhibition was noted, indicating a rather
strong affinity of the glyceride for the mercurial. However, again this ex-
planation would not seem to hold for o-iodosobenzoate, since it should in-
hibit the pancreatic lipase as well as the wheat germ lipase (with which
Singer worked) if the substrates are altered. Also it is not too surprising
that a molecule like p-chloromercuribenzoate would adsorb at an oil-water
interface, since it possesses polar and nonpolar regions, but o-iodosobenzoate
has polar groups at either end. (3) A group, such as — S — I — (p — COO", is
actually added to the enzyme near the substrate site and interferes steri-
cally as p-chloromercuribenzoate may do.
Another explanation involves the basic kinetics of such inhibitions. It
seems to have been generally assumed that when an irreversible inhibitor
reduces the affinities of each substrate of an enzyme equally, the inhibi-
tions will all be the same, which can readily be shown not to be true. Let
us assume that Kj„ is equal to K^, the true dissociation constant of the ES
complex, so that the uninhibited rate is given by:
F^(S)
(S) + K,
720 6. O-IODOSOBENZOATE
If only the affinities of the substrates are altered by the inhibitor, we may
write for the inhibited rate:
(S) + aK,
where a is a factor indicating the magnitude of the effect of the inhibitor
on the substrate binding (a > 1). The inhibition is then given by:
a - 1
(6-1)
(S') + a
where (S') is the specific concentration of the substrate, (^)IK,. Thus, even
though a is the same for each substrate, the inhibition will vary with (S').
Superficially it might appear that an irreversible inhibitor reducing sub-
strate binding would produce inhibitions independent of the substrate con-
centration, but such is not the case. Some of the confusion arises from as-
sociating this type of inhibition with noncompetitive inhibition, which it
is not in any sense. The inhibitions by o-iodosobenzoate, and probably most
SH reagents, are usually competitive, as shown by the protection afforded
by the substrate when it is present during the incubation with the inhibitor,
and K, is altered rather than k^, the rate constant for the breakdown of the
ES complex into products. It may be noted that even though Kj„ is not Kg,
but the more complex {k_i + li2)jk-^, a similar expression for the inhibition
will be found, and the substrate concentration will play a role in the degree
of inhibition produced. Furthermore, if the inhibition actually is noncom-
petitive and k^ is altered rather than K„ it can easily be shown that the
inhibition is given by:
'"^ - (6-2)
^k.
A:_i[(S') + l]
where /? is the factor by which k^ is changed by the inhibitor (/?<!). Here
the variation of the inhibition with the specific concentration of the sub-
strate is different than in the previous case, in that the inhibition rises as
(S') increases, as long as (S') does not greatly differ from unity. Indeed, at
high substrate concentrations, t = 1 — /5, the usually expected purely non-
competitive inhibition and, likewise, if iii,,, = K^., ^ = 1 — /?.
It is consequently not necessary to assume some complex mechanism in-
volving steric factors when the inhibition is found to vary with the sub-
strate used, unless the specific concentrations of all the substrates are kept
equal. It was stated by Singer (1948) in his study of lipase that the sub-
strate concentration was chosen so as to "just saturate the enzyme and
thereby give optimal activity." However, calculation of the values of (S')
INHIBITION OF METABOLISM 721
from the -flT^'s given by Singer and Hofstee (1948 b) shows that, for the
substrates used with o-iodosobenzoate, (S') varies from 4 to 23 at least and,
furthermore, the variation of the inhibition with (S') is as one would expect
from Eq. 6-1, i.e., it decreases with increasing (S'). Thus the results with
o-iodosobenzoate can be explained quite simply. However, it is not implied
that this will explain all of the results obtained by Singer, and it is quite
possible that with p-chloromercuribenzoate, where a bulky group is added
to the enzyme, steric factors also play a role. The purpose of the foregoing
treatment is to indicate the importance of keeping (S') constant when com-
paring inhibitions with different substrates.
INHIBITION OF METABOLISM
Very little quantitative work has been done on the effects of o-iodoso-
benzoate on glycolysis, respiration, the tricarboxylate cycle, or other me-
tabolic pathways, so that the following is not so informative as indicative
of possibly interesting experiments to be done. From Table 6-1 it is evident
that few glycolytic enzymes have been tested and these few are not particu-
larly sensitive to o-iodosobenzoate. Only one investigation of glycolysis
in vivo has apparently been reported, that of Harting (1947), who found
o-iodosobenzoate at 1 mM to stimulate scallop muscle anaerobic glycolysis,
as does p-chloromercuribenzoate. This may not be due to direct action on
the glycolytic system, but to some effect on the muscle membranes facilitat-
ing glucose entry. Glycolysis in muscle homogenates is definitely inhibited
by 4 milf o-iodosobenzoate (Bailey and Marsh, 1952). The changes in the
pH and phosphate fractions with time are modified, as shown in Table 6-2.
The fall in pH is quite strongly inhibited and the normal decrease in ATP
is accelerated, presumably by inhibiting ATP formation. The effects on
fructose- 1,6-diP are interesting; in the control there is an initial accumula-
tion followed by a fall to low levels — o-iodosobenzoate blocks the accu-
mulation partially, but what does accumulate remains, indicating some in-
hibition of aldolase or phosphoglyceraldehyde dehydrogenase. Part of the
depression of the early accumulation may be due to the low levels of ATP,
but it is likely that some inhibition is exerted on the enzymes forming fruc-
tose-l,6-diP, perhaps hexokinase. The prevention of the fall in creatine-P
is undoubtedly due to the potent inhibition of the transfer of the phosphate
to form ATP. The minor accumulation of phosphoglyceraldehyde produced
by o-iodosobenzoate may also point to some block of the dehydrogenase,
not unlikey at this rather high concentration.
One might expect o-iodosobenzoate to inhibit respiration fairly strongly
since several dehydrogenases and cycle enzymes are quite sensitive. The
respiration of sea urchin spermatozoa is depressed almost completely by
0.3-1 milf o-iodosobenzoate, although lower concentrations around 0.1 txlM
722 6. O-IODOSOBENZOATE
Table 6-2
Effects of o-Iodosobenzoate on Glycolysis in Muscle Homogenates"
Conditions ''^™^ Zl pH A ATP A CrP J FrPP A TrioseP
(mm)
Controls
3
-0.28
-14
-27
+27
+ 3
10
-0.76
-19
-29
+ 10
+ 4
30
-1.03
-23
-29
- 2
+ 3
60
-1.04
-23
-29
- 3
+ 3
o-Iodosobenzoate
3
-0.14
-19
- 2
+ 9
+ 5
(4 mM)
10
-0.26
-20
- 5
+ 13
+ 7
30
-0.35
-20
-10
+ 13
+ 7
60
-0.38
-20
-11
+ 13
+ 7
" The values for the phosphate fractions are changes in the per cents of the total
acid-soluble phosphorus. (From Bailey and Marsh, 1952.)
may stimulate, a phenomenon seen with other SH reagents (HgClg, js-chloro-
mercuribenzoate, arsenite, and iodoacetamide) (Barron et al., 1948). The
respiration of Ehrlich ascites tumor cells is inhibited 50% by 0.35 mM
o-iodosobenzoate, 0.1 mM inhibiting 11% and 1 mM 93% (Shacter, 1957).
Thus the susceptibility of respiration is confirmed but there are no data
for locating the principal sites of action.
The binding of K+ in liver mitochondria is believed by Gamble (1957) to
be related to the sites for oxidative phosphorylation, although it is not
directly dependent on ATP. The evidence comes from the ability of 2,4-
dinitrophenol to lower mitochondrial K+ markedly. o-Iodosobenzoate at
0.03 mM produces effects similar to 2,4-dinitrophenol, which does not nec-
essarily imply an uncoupling action of the o-iodosobenzoate, but indicates
some effect on the electron transport chain. Scott and Gamble (1961) have
found mercurials to stimulate the K+ exchange rate of mitochondria and
simultaneously to reduce the bound K+. These effects are also produced by
o-iodosobenzoate: the exchange rate is doubled by 0.08 mM, the mitochon-
drial K+ is half reduced by 0.15 mM, and oxidative phosphorylation is 50%
inhibited by 0.08 mM. These potent actions of o-iodosobenzoate point to
important effects on mitochondrial oxidative systems that apparently play
a role in the depression of respiration.
EFFECTS ON ANIMAL TISSUE FUNCTIONS 723
EFFECTS ON ANIMAL TISSUE FUNCTIONS
The injection of o-iodosobenzoate into animals or its application to skeletal
muscle preparations does not produce rigor so readily as does iodoacetate.
In the whole animal, indeed, the actions on muscle seem to be of little im-
portance, and the paralysis sometimes seen is more likely explained by a
central effect. Applied directly to isolated frog muscle in reasonably high
concentration, o-iodosobenzoate can lead to a loss of excitability and the
development of contracture, whereas o-iodobenzoate, although it depresses
excitability somewhat, does not induce contracture (Jahn, 1914). The turtle
biceps muscle is also slowly and irreversibly shortened by o-iodosobenzoate
(and iodoacetamide) at concentrations much higher than would be reached
in vivo (Pisanty, 1948). If the mechanism of iodoacetate in contracture is a
block of glycolysis, o-iodosobenzoate does not seem to share this selectivity,
which confirms what little is known from the results on enzymes and me-
tabolism (see page 721). The ability of myosin to associate with actin and
to split ATP depends on SH groups and is inhibited by o-iodosobenzoate as
well as by other SH reagents (Bailey and Perry, 1947), and the binding of
Ca++ by G-actin is depressed parallel to the reduction in polymerization
by o-iodosobenzoate (Barany et al., 1962). In these respects, o-iodosoben-
zoate is more potent and rapidly acting than iodoacetamide, and such ef-
fects may play a role in the contractures observed at high concentration,
although a metabolic site of action is not excluded. The contractile response
to ATP by nonconducting psoas muscle fibers is abolished by 0.5-1 mM
o-iodosobenzoate, and this is reversible if the fibers are incubated for 90-
120 min in 10 mM cysteine (Korey, 1950).
The heart appears to be more sensitive to o-iodosobenzoate than is skele-
tal muscle. In the initial pharmacological study by Loevenhart and Grove
(1911), intravenous injection into rabbits, cats, and dogs was found to pro-
duce a rapid fall in the blood pressure, little change in the cardiac rate, and
a decrease in cardiac output with dilation of the heart. o-Iodoxybenzoate
acts very similarly but o-iodobenzoate is inactive, indicating that the oxidiz-
ing activity is essential. Minimal effects are given in the cat by 13.2 mg
(50 //moles), so that the total concentration is probably around 0.5 mM.
However, inasmuch as Jahn (1914) showed that blood appreciably reduces
the action of o-iodosobenzoate — due to reaction with hemoglobin, other
proteins, and glutathione — the concentration of free o-iodosobenzoate is
undoubtedly much less. Jahn also showed that the perfused frog heart is
depressed by as little as 0.038 mM o-iodosobenzoate and that 0.38 mM
causes a prolonged depression of the amplitude, although not standstill or
contracture. The results of Mendez (1946) and Mendez and Peralta (1947)
on the frog heart differ from those of Jahn, in that concentrations of 0.2-
0.4 mM were found to cause an increase in the contractile amplitude, and
0.83 mM to produce systolic standstill within 20 min. Furthermore, the
724 6. O-IODOSOBENZOATE
rate always increases up to the final failure, whereas Jahn observed only
slowing. No conduction disturbances were noted. Since partial reversal can
be achieved by lengthy perfusion with o-iodosobenzoate-free medium, in
tissues some reduction of disulfide groups may occur. The dog heart-lung
preparation is quite resistant to o-iodosobenzoate, 100 mg producing no ef-
fect, although high doses increase the venous pressure, presumably by ini-
tiating cardiac failure (Mendez and Pisanty, 1949).
The effects of o-iodosobenzoate on smooth muscles are at least superficially
similar to those on skeletal and heart muscle (Alanis, 1948). Isolated rabbit
intestine and the uterus in several species are put into a form of contrac-
ture, although this is eventually followed by relaxation and loss of aU rhyth-
mic activity. These actions are similar to those of iodoacetamide and ar-
senicals.
Although the effects of o-iodosobenzoate in the whole animal indicate
marked effects on the central nervous system, no analysis of this has been
made so that sites and mechanisms are completely unknown. Frog nerve
axons are unaffected by 1 mM o-iodosobenzoate as measured by excitability
and conduction (Jahn, 1914). but the central actions are undoubtedly on
synaptic mechanisms. Neuroblastic damage has been found in developing
mice and rats after injections of o-iodosobenzoate, as with other SH re-
agents, but this probably relates more to growth and differentiation than
function (Hicks, 1953).
EFFECTS IN WHOLE ANIMALS
The earliest study of o-iodosobenzoate by Heinz (1899) is not ve»y illu-
minating since he administered potassium iodide simultaneously to generate
"nascent" iodine. However, he showed it to be irritant to the eye, the gas-
tric mucosa, and the peritoneum, and that this action seems to be due to
something other than its acidic properties. Loevenhart and Grove (1911)
confirmed the inflammatory action in the eye and subcutaneously, and show-
ed that intraperitoneal injections can be fatal as a result of the congestion
produced. Whether this is related in any way to the vesicant activity of
many SH reagents is not known. On the other hand, Bernheim et al. (1932)
found that injection into rabbits of o-iodosobenzoate inhibits conjunctival
edema induced by mustard oil. However, this could well be a nonspecific
action, since o-iodobenzoate and benzoate are somewhat active (as the am-
monium salts), and might well be mediated through the adrenal cortex.
Loevenhart and Grove (1909, 1911) investigated the pharmacological
properties of o-iodosobenzoate and related compounds because they believed
that the oxygen of this substance is physiologically active and can be used
by the tissues; e.g., o-iodosobenzoate alone does not oxidize phenolphthalin
to phenolphthalein, but does if some serum is present, this being interpreted
EFFECTS IN WHOLE ANIMALS 725
as an action mediated by peroxidase, the o-iodosobenzoate acting like hy-
drogen peroxide — furthermore, the taste of o-iodosobenzoate is almost
exactly like hydrogen peroxide. Injection of 10-20 //moles of o-iodosoben-
zoate into animals causes an immediate and marked depression of the res-
piration usually lasting 2-3 min, from which recovery occurs spontaneously.
o-Iodoxybenzoate is somewhat more potent but o-iodobenzoate is inactive.
Higher doses are required to elicit the circulatory depression described above
and the apnea is not secondary to the fall in blood pressure. Antagonism
between o-iodosobenzoate and cyanide on the respiration (the latter stim-
ulates respiration) is also observed and felt to support the concept that
o-iodosobenzoate acts by giving up its active oxygen. Jahn (1914) observed
rather nonspecific toxic effects in frogs, followed by a slowly developing
paralysis and loss of reflexes, death occurring when reflex activity has drop-
ped to zero and cardiac failure is evident. o-Iodobenzoate is less than one
tenth as toxic. The relative inactivity of o-iodobenzoate in all of these stud-
ies makes it very unlikely that any of the actions of o-iodosobenzoate are
due to the former compound, which undoubtedly is formed in the tissues.
Jahn postulated an enzyme that splits the iodine from o-iodobenzoate since
he found both organic and inorganic iodine in the urine after o-iodosoben-
zoate, the product presumably being salicylate.
Very interesting effects on the blood are observed following intravenous
infusion of 0.5 millimole of o-iodosobenzoate into rabbits (Loevenhart and
Grove, 1911). Over a period of 3 days there is a slight depression of the
erythrocytes (around 15%) and negligible effects on coagulation mechanisms
but there appears early a very marked leucocytosis, this being confined
almost entirely to the polymorphonuclears, which increase from 2,160 to
11,362 in 24 hr. It is not known if this stems from a reaction with SH groups
or an action on some metabolic system.
One factor which must be taken into account in considering the effects
of any SH reagent on the whole animal is the possible release of active
substances. Thus o-iodosobenzoate at fairly low concentrations (0.1 nxM)
releases catecholamines from the isolated chromaffine granules of the adrenal
medulla (D'lorio, 1957). This was thought to be an effect on the SH groups
located in the granule membranes, but there is no evidence for any mecha-
nism. On the other hand, the release of histamine from rat peritoneal mast
cells by Compound 48/80 is inhibited by o-iodosobenzoate, and presumably
histamine is not released by o-iodosobenzoate alone (VanArsdel and Bray,
1961).
The intravenous lethal dose in rabbits is 150-200 mg/kg (0.57-0.76 milli-
mole/kg) and such values have generally been found in most animals. Dr.
Loevenhart courageously ingested a total of 1.3 g within 5.5 hr without
the slightest effect. The lethal dose of iodosobenzene is the same as that of
o-iodosobenzoate, indicating that the carboxylate group is not essential for
726 6. O-IODOSOBENZOATE
the toxicity (Luzzato and Satta, 1910). Probably the only useful role for
the carboxylate group is to increase the solubility.
EFFECTS ON SEA URCHIN EGG DEVELOPMENT
The effects of 0.66 mM o-iodosobenzoate in sea water on the development
of Arbacia eggs was studied by Runnstrom and Kriszat (1952). It was found
that fertilization and cleavage proceed quite normally up to the blastula
stage (perhaps with a slight delay), but after 6 hr the controls are hatched
whereas the treated ones are not. After 20 hr the controls are bilateral early
plutei, but 80-90% of the treated larvae are still within their membranes,
the formation of the entoderm being suppressed in these. The animal re-
gion is characterized by a high cylindrical region of epithelium carrying a
ciliary tuft, whereas the cells at the vegetal pole are flattened. The treated
larvae contain no pigment and the pigment initially present has disappear-
ed. This effect of animalization of the larvae can be brought about by other
enzyme inhibitors (iodoacetate, parapyruvate, etc.) and has been confirm-
ed for o-iodosobenzoate by Ranzi (1955). If the larvae after 6 hr exposure
to o-iodosobenzoate are removed to normal sea water, some recovery occurs
and fairly normal plutei may be formed, although the arms are lacking and
the archenteron shows no differentiation. The general conclusion was that
oxidation of certain SH groups suppresses primarily the differentiation of
the entomesoderm.
A more detailed study of the earliest stages of Arbacia egg development
was made by Monroy and Runnstrom (1952). The high concentration of
2.64 mM o-iodosobenzoate does not prevent the fertilization reaction or the
formation of the fertilization membrane, but the membrane is somewhat
thicker and more refractile than normally. At 80 min the controls are in 2-
and 4-ceU stages with the membrane unchanged, whereas the treated eggs
are all in the 2-cell stage with conspicuous membranes. One hour later three
fourths of the treated eggs are cytolyzed with escape of pigment. The mem-
brane thickening and the escape of pigment seem to be correlated. If the
eggs are first centrifuged, thickening of the membrane occurs only at the
pole where the pigment is located. Thus the membrane changes do not ap-
pear to be due to a direct action of the o-iodosobenzoate. Possibly the na-
ture of the membrane and its later changes during development depend on
substances formed in the egg and metabolic inhibitors interfere in the pro-
duction or action of these substances.
The exposure of Paracentrotus lividus eggs to 0.35-0.7 mM o-iodosoben-
zoate does not affect subsequent fertilization or suppress cleavage, although
hatching is prevented (Hagstrom, 1963), confirming the earlier results of
Runnstrom and Kriszat (1952) on Arbacia eggs. There are, nevertheless,
differences in the response. First, cleavage is somewhat accelerated: The
EFFECTS ON BACTERIA AND VIRUSES 727
controls at 135 min after fertilization are 2% in the 2 -cell stage, 50% in the
4-cell stage, and 48% in the 8-cell stage, whereas those treated with o-iodo-
sobenzoate are 8% in the 4-cell stage and 92% in the 8-cell stage. Second,
there is no obvious disturbance in development, e.g., no evidence of animal-
ization, and the ciliated embryos inside their membranes appear to be nor-
mally active. Differentiation in Paracentrotus is thus less susceptible than in
Arbacia to o-iodosobenzoate. Higher concentrations of o-iodosobenzoate may
produce other effects on eggs but whether these actions are mediated through
SH group oxidation is not known. The eggs of Hemicentrotus pulcherrimus
and Urechis unicinctus elevated the fertilization membrane when incubated
for 10 min in 10 raM o-iodosobenzoate at pH 4 and then returned to normal
sea water (Isaka and Aikawa, 1963). It was suggested that the vitelline and
plasma membranes are connected by hydrogen bonds and that o-iodosoben-
zoate and other SH reagents react with SH groups in the plasma membrane,
weakening these bonds and allowing separation of the membranes. Move-
ments during cleavage have been supposed to involve contractile proteins
as in muscle, and threads formed from fibrous proteins obtained from Hemi-
centrotus eggs contract when metal ions (e.g., Mg++, Cu++, Cd++, etc.) are
added (Sakai, 1962). This contraction is blocked by 5 rsxM o-iodosobenzoate
and high concentrations of other SH reagents, indicating that SH groups
are necessary.
EFFECTS ON BACTERIA AND VIRUSES
The early interest in the antibacterial actions of iodine led Arkin (1911),
in connection with the pharmacological studies of Loevenhart and Grove
at Wisconsin, to investigate the effects of o-iodosobenzoate and related
compounds on various bacteria. Eberthella typhosa, E. coli, S. aureus, and B.
pyocyaneus are all killed by exposures of 24 hr to 1 mM at 37°, not surpris-
ingly. o-Iodoxybenzoate is even more potent, but o-iodobenzoate does not
kill even at 10 mM. Jahn (1914) found the growth of E. coli to be inhibited
by 0.38 mM o-iodosobenzoate, but not by 38 mM o-iodobenzoate, indicat-
ing the importance of the oxidative action and confirming the results in
animals. Chinard (1942) considered the possibility of using o-iodosoben-
zoate locally in infected wounds. He observed marked inhibition of the
growth of E. coli at 0.02 mM with eventual death of the bacteria in 72 hr,
and death of hemolytic streptococci at 0.38 mif. If the o-iodosobenzoate is
injected with these streptococci subcutaneously into mice, no infections are
seen, but all the control mice die. The flagellar activity of B. brevis is well
inhibited by 2 mM o-iodosobenzoate at 30 sec and maximally at 5 min
(De Robertis and Peluffo, 1951). Yeast is more resistant, since it requires
3.8 mM to inhibit the growth 50% (Loveless et al., 1954). No analyses at
all have been made of the sites or mechanisms of action. It is likely that
728 6. O-IODOSOBENZOATE
the activity against bacteria will be strongly influenced by the media used
and the other conditions; most of the media for pathogens contain substances
readily reacting with o-iodosobenzoate. Phagocytosis of staphylococci and
streptococci by human leucocytes is stimulated by o-iodosobenzoate, but
this is indirect since it occurs only in the presence of serum, and is perhaps
an activation of serum opsonin (Arkin, 1912).
The psittacosis virus is 30-75% inactivated by exposure to 0.1 roM o-iodo-
sobenzoate for 1 hr at 37°, only p-chloromercuribenzoate of aU the agents
tested being more potent (Burney and Golub, 1948). In addition, it is the
most effective substance in reducing viral growth in chick embryo cultures
without inhibiting culture growth. The selectivity is probably not great
enough to warrant clinical interest.
CHAPTER 7
MERCURIALS
The mercurials occupy a rather special niche in the subject of enzyme
inhibition; they are very useful for demonstrating the presence and impor-
tance of SH groups in enzyme reactions, but apparently lack specificity
toward particular enzymes or classes of enzymes. Since so many enzymes
contain reactive SH groups at or near the active center, the mercurials
would seem to inhibit more enzymes than they leave unaffected. When a
mercurial acts on living cells, one cannot state which enzymes are affected
most readily. In other words, they are reasonably specific with regard to
the molecular group attacked, but quite nonspecific at the enzyme or cel-
lular levels. The mercurials wiU, in addition, react with nonenzymic proteins
and may modify complex systems by mechanisms unrelated to metabolism.
The mercurials are thus at present generally useless as tools to study the
relationship of a particular enzyme to the over-all metabolism, growth, or
function of a cell or organism. Nevertheless, with judicious use, they may
give some insight into the broader metabolic basis of function, as in certain
studies of gastric acid secretion, renal transport, and mitosis. Their primary
use, however, is the detection and titration of SH groups on enzymes.
They are often stated to be the most specific SH reagents; this may be
questioned, but without doubt they are among the most reactive reagents
and seldom does one find SH groups resistant to the mercurials and capable
of reacting with other SH reagents. Like all inhibitors, they are valuable
only when used in the proper system. It is always tempting to use inhib-
itors such as the mercurials which will almost always produce definite
effects, but unfortunately the results usually cannot be interpreted satis-
factorily. We shall emphasize the quantitative side of mercurial action and
the inhibitions of pure enzymes, discussing only briefly effects observed on
complex systems, inasmuch as little useful information can be derived
from this latter work.
The medical use of the mercurials can be traced back for over 3000 years,
although their modern therapeutic applications began with the rediscovery
of the diuretic action of mercurous chloride in 1849 (since then this action
has been rediscovered several times), the demonstration of the antiseptic
729
730 7. MERCURIALS
action of mercuria chloride by Koch in 1881, and the introduction of or-
ganic mercurials for diuresis, antisepsis, and other chemotherapeutic pur-
poses from 1900 to 1920. The marked toxicity of inorganic mercury was
recognized in antiquity and became a more critical problem over 400 years
ago, especially in processes such as fur felting for hats and more recently
in the widespread use of mercurials as plant fungicides for various rots and
rusts. There was a good deal of experimentation and speculation on the
nature of mercurial antisepsis between 1900 and 1940, but little of this is
pertinent to our present purposes. The early work was much concerned
with the examination of the validity of certain vague concepts, such as the
Arndt-Schulz law (which states that drugs stimulate in low concentration
and inhibit in high concentration), oligodynamic action, and the Ostwald
adsorption theory. Despite the fact that the combination of mercurials with
thiols, e.g. cysteine, has been known since 1875 at least, investigations on
the metabolic effects and enzyme inhibition are very sparse before 1930.
Actually the mercurials have been intensively used by biochemists for the
characterization of enzymes for only the past several years. Of the some
1350 publications on the effects of mercurials on isolated enzymes, only 4%
were issued prior to 1950, 16% from 1950 to 1956, and 80% from 1956
through 1964. By the time this volume goes to press, approximately half
of the publications on this aspect of the mercurials will have appeared after
1960. These figures indicate essentially that each newly isolated enzyme is
subjected to one or more mercurials for the purpose of detecting SH groups.
One of the major aims of this chapter is to attempt to determine the va-
lidity and usefulness of such determinations.
CHEMICAL PROPERTIES
The most commonly used inorganic mercury compound in inhibition work
is mercuric chloride (HgCla). Some fundamental properties of the Hg++ ion
and its halides are summarized in Table 7-1. It may be noted that although
the linearity of HgXg molecules is established and the configuration of
certain HgX4 complexes appears to be tetrahedral, the nature of the HgClg"
and HgCl4= ions is not clear and a planar arrangement is possible. The
aqueous solubility of HgClg increases with the concentration of NaCl, KCl,
or other halide present; thus the solubility of HgClg in Krebs-Ringer me-
dium is around 12.5 and in sea water around 27 g/100 ml (Barnes and
Stanbury, 1948). The deficiency in the ionic character of HgClg is indicated
by the high solubility in ethanol (26.3 g/100 ml) and even in ether (4.55
g/100 ml). Indeed, HgClg has been said to be reasonably lipid-soluble, a
fact of some importance in considering the distribution in the tissues. HgBrg
and Hglg are much less soluble than HgClg in water and seem to have no
advantages over HgClg in enzyme studies.
CHEMICAL PROPERTIES
731
Table 7-1
Some Properties of Mercury, the Mercuric Ion, and the Mercuric Halides
Radii
Hg atom
Hg++ ion
Bond lengths
Hg-Cl
Hg-Br
Hg-I
Bond ionic character
Hg-Cl in HgCIs,
Bond types
HgCl,
HgClr
Electronegativity
Hg(II)
Solubility in water (g/100 ml solution]
HgCl,
Solubility product
HgCl,
K,^= (Hg++)(C1-)^
pH of saturated solution
HgCl,
Redox equilibrium
Hg++ + Hg(I) ±^ Hg,++
K = (Hg,++)/(Hg++)
Redox potentials {E\r,o)
Hg2++ ±^ 2 Hg++ + 2e-
2 Hg ±5 Hg2++ + 2e-
Hg ±? Hg++ + 2e-
1.59 A
0.66 A
2.20 A
2.40 A
2.55 A
28%
Linear {sp)
Tetrahedral (sp')
1.9
6.8 (25°)
8.9 (37.5°)
1.06 X 10-" (25°)
2.95 X 10-i» (37.5°)
4.7 (25°)
129.2
- 0.92 V
- 0.79 V
- 0.85 V
732 7. MERCURIALS
Equilibria between Hg++ and Halide Ions
In aqueous solution HgClg does not dissociate simply into Hg++ and CI"
ions, but forms a series of complexes, the relative concentrations of which
depend on the CI" concentration and the pH. The following species are the
most important: Hg++, HgCl+, HgClg, HgClg", and HgCl4=. This applies to
acid solutions where hydrolysis and hydroxyl complexes can be ignored
(see next section). Higher CI complexes with Hg++ can be neglected in
biological work, as can univalent Hgg^ + and its complexes (since no equilib-
rium with metallic mercury occurs). Sillen and his collaborators in Stock-
holm have summarized their extensive investigation of the halide complexes
of mercury (Sillen, 1949) and we shall follow their values for the equilib-
rium constants (it should be noted that their work was done at 25° so that
small corrections should be applied for solutions at other temperatures).
However, we shall differ in two ways from Sillen in the expression of the
constants. In the first place, we shall use dissociation rather than association
constants, in conformity to the usage throughout this book. In the second
place, we shall indicate the individual dissociations by ^'s and the cumu-
lative dissociations by /5's, in conformity with the usual terminology in
metal-ligand complexes and chelates (Bjerrum et al., 1957). The fundamental
dissociations and their constants can be formulated as in Table 7-2. The tight
binding of the first two Cl~ ions is evident, but the next two are bound only
weakly, due perhaps to the change in bond configuration and the increasing
negativity; that the latter is not a major factor is indicated by the similar
behavior of the ammonia complexes. The constants for the Br" and I"
complexes are much less than for CI", i.e., the former ions are more tightly
bound to Hg++, but such equilibria are seldom of importance in biological
systems.
The relative concentrations of the various complexes depend in simple
solutions mainly on the Cl~ concentration. The fractions of the total mer-
cury in particular complexes may be calculated from the following equa-
tions:
(7-1)
where
(Hg++)/(Hg,) =
(HgCl+)/(Hg,) =
(HgCl,)/(Hg,) =
(HgCl3-)/(Hg,) =
(HgClr)/(Hg,) =
1/A
(Cl-)/iS.A
(C1-)V/3.A
(C1")VM
(C1-)V/J4A
(C1-) (Cl-)=
(Cl~)^
A = 1 + ^-— + — - — H — +
(Cb)^
i5i /3, /33 ^4
The CI" concentration varies over a wide range in the media used. In isolated
enzyme work it may be very low (unless KCl or NaCl is added); it is 102 mM
CHEMICAL PROPERTIES
733
o
o
o
Q
^-
II
c
O
>.
W
Q
^
w
It
it
it
it
1
5
o
1
o
1
3
+
+
+
+
+
+
be
1
o
1
a
H c
734
7. MERCURIALS
in serum, 126 mM in Krebs-Ringer bicarbonate medium, 143 mM in Tyrode
solution, 154 mM in physiological saline, and 515 vaM in sea water. The
distribution between species of complexes for three situations (low, moder-
ate, and high Cl~) is shown in Table 7-3, and the distribution over a com-
plete spectrum of Cl~ concentrations is illustrated in Fig. 7-1. It so happens
Fig. 7-1. Curves showing the distribution of the different chloride com-
plexes of Hg++ with Cl^ concentration. (From Sillen, 1949.)
that, in most media used in ceU and tissue preparations, the concentrations
of HgClg, HgClg", and HgCl4= are roughly equal, whereas in sea water the
predominant form is HgCl4=. In the media for isolated enzyme study, in
which Cl~ is often low, the predominant form may be HgClg or even HgCl+.
Table 7-3
Distribution of Mercuric Chloride Complexes as Fractions
OF the Total Mercury in Media of Different CI" Concentration
Fraction
(C1-) = 1 ml/
Krebs-Ringer medium
(C1-) = 126 mM
Sea water
(C1-) = 515 mM
(Hg++)/(Hg()
6.03 X 10-8
1.26 X 10-12
9.8 X 10-is
(HgCl+)/(Hg,)
3.31 X 10-"
8.68 X 10-'
2.8 X 10-«
(HgCl,)/(Hg,)
0.9925
0.331
0.0428
(HgCl3-)/(Hg,)
7.09 X 10-3
0.296
0.156
(HgCIr)/(Hg,)
7.09 X 10-5
0.373
0.801
It has often been assumed in the past that the mercuric ion Hg++ is the
predominant form or the active inhibitor, but it is now realized that this
is not the case. The importance of such complexes in inhibition studies is
2-fold. In the first place, the equilibria for the binding of mercury to SH
CHEMICAL PROPERTIES
735
groups are modified, since one may say that the Cl~ ions are competing
with the SH groups for Hg++; this affects the dissociation constants for
R — S — Hg+ and R — S — Hg — S — R complexes (see page 740). In the sec-
ond place, the penetration of the inhibitor into cells will depend on the rel-
ative concentrations of these complexes. A few investigators have realized
the implications of such complexes and have attempted to take into ac-
count the equilibria under their experimental conditions. Jowett and Brooks
(1928) calculated the relative concentrations of the complexes in a 0.2 milf
solution of HgClg in Locke's medium in a study of the effects of HgClg on
tissue glycolysis and respiration, and concluded that the dominant pene-
trating species is HgClg, although they were uncertain as to the form ef-
fective on the enzymes. Barnes and Stanbury (1948) realized that Hg++ is
extremely low in sea water in their investigation of the toxic actions of
HgClg on a copepod, and assumed that the prevalent species were HgClg"
Hg"
56
.10"
10
1 Bl
10''
zs
,0-'^
70
10""
9.8.10"'^
HgCI
3.1
10'
5
50
10-^
1.4
10-^
12
10"^
28>I0'^
ID
-
,
08
-
~~~^
-sHgClj
^^
0.6
-
\
\
/^
0 4
-
\
>/
0.2
FRACTION
OF
-
HgClj
-^"gcij
^
-^
^
^
^::^
TOTAL
1
______—-
100
1000
mM
Fig. 7-2. Fraction of Hg in various forms in acid medium with varying Cl"
concentration. The figures at the top give the concentrations of Hg++ and
HgCl+ at selected Cl~ concentrations.
and HgCl4=. Green and Neurath (1953) likewise discounted the importance
of Hg++ in the inhibition of trypsin in a medium containing 10 mikf CI".
In order to facilitate estimation of the relative concentrations of the com-
plexes in a narrower range of Cl~ concentration as commonly used in inhi-
bition studies. Fig. 7-2 is presented. However, before considering these com-
plexes further, it will be necessary to discuss their so-called hydrolysis in a
pH range around neutrality.
736 7. MERCURIALS
Equilibria between Hg++ and Hydroxyl Ions
It has generally been assumed that the Hg++ ion is hydrated and that
this ionizes according to the following equations:
Hg(H,0),++ ^ HgOH(H,0)+ + H+
P^a,
= 3.70
HgOH(H20)+ ^ Hg(OH), + H-
VK.,
= 2.60
This is essentially saying that the hydrated ion is a dibasic acid (Hietanen
and Sillen, 1952). It is evident that at pH's near neutrality, Hg(0H)2 will
be the predominant form. For our purpose and comparison of these equilib-
ria with those for Cl~, it might be better to express the reactions as simple
complexing with OH" ions. Thus p^oH ~ 10.3, and p^qh — 11-4:
(Hg++) (OH )
Hg+- + OH- ±. HgOH+ ZoH, = ^jrnl.^ = ^-Z^". = ^ ^ 1^""
* {HgOH + ) '
HgOH- + OH- ^ Hg(OH), ^OH, = ^^^h^qh^^^ = ^'^'^"^ = * "" ^^""
which may be compared to p^C^] = 6.74, and p^^ = 6.48 (Table 7-2).
Now in a Cb-free medium, even at pH 5, the ratio [Hg(0H)2]/(Hg++) will
be 5000, and at pH 7 will be 50,000,000, so that Hg++ will be negligible.
Although the affinity of Hg++ for OH" is greater than for CI", when CI" is
present in appreciable concentration (e.g. 10-150 mM) it will compete ef-
fectively for the Hg++ ion since at neutrality its concentration will be 10^
to 10® times that of OH". Therefore one would predict that, in the usual
media for inhibition studies, the Cl~ complexes will predominate over the
OH" complexes although, particularly as the pH is increased above 7, it
is clear that complexes of the type HgCl(OH), HgCl(0H)2", HgCl2(0H)-,
and HgCl3(0H)= may contribute significantly to the total population. The
data given by Sneed and Brasted (1955) allow one to calculate the constants
for the following equilibria:
HgCl- + OH- ^ HgCl(OH) ^"$^pinwfi"^ = 3.2xl0-«
[HgCl(OH)]
HgCl3- + OH- ^ HgCl3(0H)= ^S?.r)nMfi^ = ^"^ ^ 1^"*
[HgCl3(0H)=]
The affinities of the CI" complexes for OH" are thus of the same order of
magnitude, and less than for the Hg++ ion. At pH 7, (HgCl+)/HgCl(OH) =
0.32 and (HgCl3")/HgCl3(OH)= - 0.5, so it is seen that these OH" com-
plexes are indeed significant. The importance of these OH" and mixed com-
plexes for inhibition studies is, of course, the same as that of the CI" com-
plexes, but the concentration of Hg++ wiD be even less than calculated in
the previous section.
CHEMICAL PROPERTIES 737
Complexes of Hg++ with Various Ligands
Most metal ions, including Hg++, form strongly ionic covalent bonds with
ligand atoms capable of donating electron pairs, both Cl~ and 0H~ being
simple examples of this. We would expect that Hg++ might complex readily
with a variety of substances, many of which occasionally occur in media
used for inhibition studies. It is usually stated that Hg++ reacts with SH
groups selectively and that other groups on proteins seldom contribute to
the binding; it is necessary to look into this matter quantitatively, and ob-
tain some idea of the relative affinities of the various groups for Hg++.
Complexes of Hg++ with ammonia are well known so that binding to amino
groups might be predicted and, since some interaction with carboxylate
groups is likely, it may be anticipated that amino acids would provide ef-
fective ligands. Indirect evidence for such complexes was obtained by Salle
and Ginoza (1943) by showing that several amino acids reduce the bacte-
ricidal activity of HgClg. The minimal lethal concentration of HgClg is in-
creased 6 times by glycine, aspartate, glutamate, arginine, and lysine at
67 inM, and 120 times by cysteine. This indicates appreciable complexing
with amino acids under physiological conditions, although the reaction with
the SH group of cysteine is evidently stronger than with other groups.
Haarmann (1943 a,b) claimed that whereas 1 equivalent of Hg is bound to
certain amino acids at pH 7, as much as 4 to 8 equivalents may be bound
at pH 11, some loosely and some tightly. A definitive investigation was
made by Perkins (1952, 1953) and a number of stability constants were
determined. Two major complexes were assumed, probably with the follow-
ing structures:
OC— O OC— O N
H,
Complex I Complex II
The composite constant, ^2 = K^K^, where K^ and K^^ are defined by the
following equilibria:
Hg++ + AA- ±s HgAA+ K,
HgAA+ + AA- ±^ Hg(AA)2 K,
(Hg++) (AA-)
(HgAA+)
(HgAA+) (AA-)
Hg(AA),
was determined in each case, and these values are given in Table 7-4 along
with the dissociation constants for a number of ligand complexes. The form
of the amino acid necessary for chelation with Hg++ is "OOC — E — NHg
738
7. MERCURIALS
Table 7-4
Dissociation Constants for Various Mercuric Complexes"
Ligand
Pi^i
P^.
VP2
Reference *
Inorganic ions
ci-
6.74
6.48
13.22
(1)
Br-
9.05
8.28
17.33
(1)
I-
12.87
10.95
23.82
(1)
OH-
10.3
11.4
21.7
(2)
CN-
—
—
34.7
(3)
SCN-
—
—
17.4
(3)
Pyrophosphate
—
—
17.45
(4)
Nitrogenous compounds
Methylamine
8.6
9.3
17.9
(5)
Triethylamine
7.8
7.8
15.6
(5)
1,2-Diaininopropane
—
—
23.5
(5)
1 ,2,3-Triaminopropane
19.6
—
—
(5)
Ethylenediamine
—
—
23.4
(5)
Ethanolamine
8.5
8.8
17.3
(5)
Diethanolamine
7.8
7.8
15.6
(5)
Triethanolamine
6.9
6.2
13.1
(5)
2,2'-DiaminodiethyIamine
21.8
—
—
(5)
Triethylenetetramine
25.3
—
—
(6)
Ammonia
8.8
8.7
17.5
(7)
Pyridine
5.1
4.9
10.0
(3)
Piperidine
8.7
8.7
17.4
(5)
Imidazole
—
—
16.7
(8)
Ethylenediaminediacetate
9.75
6.05
15.8
(5)
Ethylenediaminetetraacetate
22.1
—
—
(5)
Hexamethylenediaminetetraacetate
21.4
—
—
(5)
Amino acids
Glycine
10.3
8.9
19.2
18.2
(5)
(9)
Glycylglycine
—
—
12.4
(9)
Alanine
—
—
18.4
(9)
Leucine
—
—
17.5
(9)
Proline
—
—
20.5
(9)
CHEMICAL PROPERTIES 739
Table 7-4 (continued)
Ligand
pK,
P^2
P^2
Reference *
Serine
_^
_
17.5
(10)
Tyrosine
—
—
17.1
(10)
Arginine
—
—
17.4
(10)
Histidine
—
—
21.2
(8)
Methionine
6.52
4.93
11.45
(11)
Ethionine
7.25
5.92
13.17
(11)
Cysteine
14.21
—
—
(11)
<S-Methylcysteine
7.20
5.81
13.01
(11)
Purines and pyridimidines
Adenine
—
—
11.5
(12)
Adenosine
—
—
8.5
(12)
Thymine
—
—
21.2
(12)
Thymidine
—
—
21.2
(12)
Cytosine
—
—
10.9
(12)
Miscellaneous
Acetate
4.0
—
—
(13)
Cyclohexene
4.3
—
—
(5)
Penicillamine
16.15
—
—
(11)
" (^2 is the cumulative dissociation constant for HgLj, and is K^^K^
* References:
(1) Sillen (1949).
(2) Hietanen and Sillen (1952).
(3) Simpson (1961).
(4) Yamane and Davidson (1960).
(5) Bjerrum et al. (1957).
(6) Chaberek and Martell (1959).
(7) Bjerrum (1941).
(8) Brooks and Davidson (1960).
(9) Perkins (1952).
(10) Perkins (1953).
(11) Lenz and Martell (1964).
(12) Ferreira et al. (1961).
(13) Gurd and WUcox (1956).
740 7. MERCURIALS
and, in the calculations of the constants, only that fraction of the amino
acid at the pH used was considered.
The fact that simple amines complex with Hg++ to approximately the
same degree as the amino acids indicates that the amino group is the im-
portant ligand, the carboxylate group perhaps contributing slightly to the
stability. Ring nitrogen atoms are probably not as effective as amino groups.
Hg++ reacts with both the amino and imidazole groups of histidine, but
more tightly with the former, the pj^'s being 10.6 and 7.5, respectively
(Simpson, 1961). The effect of the pH on the stability of these complexes
is well illustrated by the constants for the following equilibria with histidine
(Brooks and Davidson, 1960):
Hg++ + 2 hist- ±5 Hg (hist)2 p/ff^ = 21.2
Hg++ + hist- + H-hist i? Hg (hist) (H-hist)+ Pi92 = 18.4
Hg++ + 2 H-hist 15 Hg (H-hist)2++ pp^ = 15.0
where hist designates the ~00C — R — NHg form and H-hist the OCC —
R — -NH3+ form. These complexes with histidine were assumed to be linear
and it was claimed that chelation must play only a small role in Hg++
complexes due to the tendency of Hg++ to form linear complexes. This
brings up an interesting point of importance in understanding the reactions
of HgClg with proteins and enzymes. Certainly some of the most stable
complexes of Hg++ — as with 1,2,3-triaminopropane, triethylenetetramine,
and EDTA — must be chelates and nonlinear, and it is also well known
that Hg++ reacts with dimercaprol (BAL) to form a ring with the two SH
groups. Whether chelation is or is not important in any case probably de-
pends on several factors, such as the spatial arrangement of the ligand
groups, the intrinsic affinity of the Hg++ for these groups, and the entropy
changes accompanying the formation of the complex. Certainly the third
and fourth ligands generally add to the HgLg complex much less readily
than the first two, as we have seen for CI". This is also true for ammonia,
the successive constants being given by pifj = 8.8, piiTg = 8.7, pK-^ = 1.0,
and TpK^ = 0.78. Thus the formation of HgLg and UgL^ type of complexes
must involve some additional factors increasing the stability.
In any event it is clear that the complexes of Hg++ with amino acids
and many other compounds are stable enough so that, when these sub-
stances are present in the media used for the study of inhibition, a signifi-
cant fraction of the Hg may be in the form of such complexes. For example,
it may be calculated for a solution containing 1 mM glycine and a total
concentration of Hg of 0.1 mM that (Hg++) = 6.3 x lO-i^ M. If the Cl"
concentration is appreciable, this will reduce the binding to these other
ligands. One may visualize the situation somewhat as follows. In most
media there will be several substances — Cl~, OH", buffers, amino acids,
substrates, etc. — capable of complexing with Hg++. The end result will
CHEMICAL PROPERTIES 741
be that the Hg++ concentration will be extremely low, the Hg being parti-
tioned between numerous complexes of different types, each one reducing
to some extent the reaction of Hg with enzymes. It must be clearly under-
stood that when inhibition is stated below to be by HgCla or Hg(N03)2 or
Hg acetate, it refers only to what was added and not to the dominant
form present or the active inhibitor.
Complexes of Hg++ with Nucleotides and Nucleic Acids
Complexes between Hg++ and certain purines and pyrimidines, especially
thymine, are quite stable (Table 7-4) (Katz, 1962), and complexes with
phosphates are probably formed readily; thus, one would expect nucleotides
and nucleic acids also to bind Hg++ rather well. Inagaki (1940) found var-
ious nucleotides, such as AMP, GMP, and IMP, to be precipitated by mer-
curic compounds, but unfortunately no further work has been done on these
complexes. One would like to know the nature and extent of the interactions
of mercurials with ATP, NAD, FAD, and related substances. However,
the studies of the reactions between Hg++ and nucleic acids have recently
been accelerated, and it is obvious that the results could be very important
in understanding the effects of mercurials on cellular growth and prolifera-
tion. Hg++ has been found to complex with nucleic acids from thymus
(Katz, 1952), plants (Trim, 1959), pneumococci (Dove and Yamane, 1960),
and tobacco mosaic virus (Katz and Santilli, 1962 b). The general effects
on the nucleic acids may be summarized briefly as follows: a decrease in
viscosity; an increase in turbidity, sedimentation constant, aggregation,
and the dimer : monomer ratio; an increase in the flexibility of the chains
with the assumption of a more compact configuration; and a change in the
ultraviolet absorption spectra (Katz, 1952; Thomas, 1954; Yamane and
Davidson, 1961). It was originally believed that the complexing is with the
phosphate groups, but the nature of the absorption spectrum changes, the
stoichiometry of the reactions, and the release of H+ indicate that the bases
are the sites of binding, the Hg:base combining ratio being 1 : 2 in most
cases, Hg apparently bridging the double strands of the DNA helix (Katz,
1962). The single-stranded tobacco mosaic virus RNA, however, gives a
combining ratio of 1 : 1, as expected (Katz and Santilli, 1962 a). It may be
noted that the combining ratio is 1 : 2 for guanine oligoribonucleotides, such
as GpCxpG (Lipsett, 1964). These complexes are usually completely reversi-
ble upon adding various Hg++ complexers (Cl~, cyanide, EDTA, or thiols),
and indeed the pneumococcal transforming DNA after demercuration retains
aU of its activity (Dove and Yamane, 1960), and the tobacco mosaic virus
after removal of the Hg++ regains its infectivity' (Singer and Fraenkel-
Conrat, 1962), these observations indicating that the original configurations
of the nucleic acids can be restored despite the apparently marked struc-
tural modifications occurring during reaction with Hg++.
742 7. MERCURIALS
Organic Mercurials
It will be convenient to discuss some of the general properties of the or-
ganic mercurials before coming to the important problem of the reaction of
mercurials with SH groups. Many organic mercurials were developed for
chemotherapy, disinfection, and diuretic activity and, although some of
these have been occasionally used in inhibition work, mercurial inhibitors
are generally simpler structurally. The chemical properties and structure-
action relationships will be taken up later for the antiseptics (page 970)
and diuretics (page 917). The aryl mercurials (such as p-chloromercuri-
benzoate) were introduced as enzyme inhibitors by Hellerman (1937) and
the alkyl mercurials (such as methylmercuric chloride) as protein reactants
by W. L. Hughes (1950). The accompanying formulas are for the ions which
V\ A^Hg H3C-Hg
Phenylmercuric ion Methylmercuric ion
^-Mercuribenzoate ion /'-Mercuriphenylsulfonate ion
eventually complex with SH groups and other ligands. They are used as
chlorides, hydroxides, nitrates, or acetates but, once they have been added
to the medium, the ions as formulated complex with various ligands which
may be present, essentially as Hg-^+ does. Thus it is correct to speak of
phenylmercuric acetate or p-chloromercuribenzoate as the mercurial used,
but this terminology does not give an accurate representation of the forms
present in solution. For example, it makes no difference in the final inhi-
bition whether one uses p-chloromercuribenzoic acid or sodium p-hydro-
xymercuribenzoate, the two forms of the p-mercuribenzoate ion commonly
available. For the sake of compression, we shall use the abbreviations shown
in the following tabulation in the remainder of this chapter.
Ion Abbreviation
Phenylmercuric PM
Methylmercuric MM
p-Mercuribenzoate p-MB
p-Mercuriphenylsulfonate p-MPS
CHEMICAL PROPERTIES 743
It will be useful to summarize briefly some of the important differences
between the organic mercurials and HgClg.
(a) Functionality. HgClg is bifunctional in the sense that it can react
with two ligands to form L-Hg-L complexes, whereas the organic mercu-
rials are monofunctional in that they can react with only one ligand to
give R-Hg-L. Hg++ can also form cyclic mercaptides with two adjacent
SH groups but the organic mercurials cannot. These differences are often
very important in the reactions with thiols and enzymes, and in fact one
of the major reasons for the preference of many investigators for the or-
ganic mercurials, especially in the quantitative titration of SH groups, is
their monofunctional nature.
(b) Aqueous solubility. The organic mercurials are less soluble than HgClg
and occasionally this has created a problem if higher concentrations are
required. However, in most titration or inhibition studies, the concentra-
tion required is seldom over 1 mM, and this can be easily attained in most
cases. Phenylmercuric chloride is soluble to the extent of only 0.16 toM in
distiUed water, but a good deal more soluble in salt solutions, and the in-
troduction of anionic groups increases the solubility. It is well known that
23-chloromercuribenzoic acid, which for many years was the only commonly
used mercurial, does not readily go into solution at neutral pH. It is there-
fore usual, to dissolve it in dilute KOH or NaOH solutions (0.01-0.05 M)
and adjust to the desired pH with HCL* However, the sodium salt is now
commercially available and dissolves readily. The phenylsulfonic acid is
also more soluble than the benzoic acid derivative. The solubility will be
determined, as with HgClj, by the concentrations of various complexing
substances in the medium; thus the solubility is reasonably high in most
physiological media containing over 100 mM Cl~, or various other ions
such as pyrophosphate or sulfate, concentrations around 10 mM of mer-
curial being readily obtained.
(c) Lipid solubility. The unsubstituted alkyl and aryl mercurials are more
soluble than the Hg++ ion and its complexes in lipids. Hughes (1957) has
estimated that the simple alkyl mercurials are around 100 times more sol-
uble in lipids than in water. This property wiU presumably allow the organ-
ic mercurials to penetrate more readily than inorganic mercury into cells
and tissues, and evidence for this is provided by the greater central nervous
system toxicity of the organic mercurials (page 951). In this connection,
* Although I have no definite evidence that strongly alkaline media are detrimental
to p-MB, I would prefer not to use 1 M NaOH solution to dissolve the material, as
has been done by some (e.g., Snodgrass et al., 1960), since, as we shall see, the C — Hg
bond is weak and dissociation is a possibiltiy, and, furthermore, such strongly alkaline
solutions are not necessary. For most work it is satisfactory to dissolve p-chloromer-
curibenzoic acid in 0.02 M KOH or NaOH at approximately 2 mg/ml or 7.4 mM.
744 7. MERCURIALS
if maximal penetration is desired, it is probably best to use the alkyl or
the unsubstituted-phenylmercurials, since the C00~ and SO3" groups will
reduce the permeability.
(d) Configuration. Although HgClg is linear, as shown by Raman spectra
and electron diffraction, the organic mercurials for some reason are ap-
parently not. Dipole moment studies (e.g., for phenylmercuric chloride,
ju = 2.99) suggest that the angle of the C — Hg — X bonds is around 130°
or higher (Sipos et al., 1955). The moment is directed as foUows:
+ -
R - Hg - X
(e) Molecular size. The organic mercurials are, of course, larger than the
simple complexes of Hg++. This may be of importance in the reaction with
the SH groups of proteins and enzymes, since steric factors may impede the
approach of the mercurial to SH groups not exposed on the surface. A re-
duction in volume was one of the reasons for the introduction of the alkyl
mercurials by W. L. Hughes (1950). In addition, penetration into cells will
depend to some extent on the molecular size. Other factors will be discussed
relative to enzyme inhibition.
(f ) Affinities for ligands. The organic mercurial ions in solution tend to
complex with various ligands in the same way as the Hg++ ion, forming
complexes of the type R-Hg-L. The affinities seem to be somewhat less
for the organic mercurial ions than for Hg++, although very few have been
studied. Simpson (1961) gave the dissociation constants for MM complexes
(Table 7-5), and generally the pii's are around 1.3 units less than for the
Hg++ complexes. Nevertheless, the affinities are of sufficient magnitude so
that at pH 7 there is perhaps 500 times as much MM-OH as MM+, and if
much Cl~ is present there may be 100 times as much MM-Cl as MM-OH
(Hughes, 1957). Rowland (1952) determined the equilibrium constant,
ii:=(RHg-OH) (H+) (Cl-)/(RHg-Cl), for a variety of diuretic mercurials,
and found a mean value for ])K of 9.9, so that at pH 7 and (Ch) = 100 mM
the ratio (RHg-Cl)/(RHg-OH) is around 100 as for MM. Ledoux (1953)
reported the interaction of p-MB with nucleic acid and from the spectral
changes assumed a complex to be formed with the carbonyl group of pyri-
midines. Various complexes of PM and amino acids were obtained by Smalt
et al. (1957) but were claimed to dissociate rather readily. The remarkably
tight complex of PM and thyroxine has been investigated by Frieden and
Naile (1954). One must thus assume that in solution in most physiological
media the organic mercurials will exist in a variety of complexes, and that
this will be an important factor in determining the degree of reaction with
proteins and enzyme SH groups.
One characteristic of the organic mercurials is the weakness of the C — Hg
bond, the energy of which is only 15-19 kcal/mole (CottreU, 1954), so that
CHEMICAL PROPERTIES 745
Table 7-5
Dissociation Constants for Complexes with Methylmekcuric Ion "
Ligand pK
CI- 5.45
Br- 6.7
I- 8.7
OH- 9.5
CN- 14.2
SCN- 6.1
Acetate 3 . 6
Phenolate 6.5
HEDTA^ 6.2
Ammonia 8.4
Pyridine .4.8
Imidazole 7 . 3
Histidine (NHj group) 8.8
Histidine (imidazole group) 6.4
" From Simpson (1961).
inorganic Hg may be split off more readily than is usually supposed. The
exchange reaction:
Hg2«3Cl2 + -OOC— 97— Hg+ ±? HgCl^ + -OOC— (p— Hg2«^ +
is fairly fast, the rate constant being 5.4 liters mole^^sec"^ at 25° with an
activation energy of 12 kcal/mole (Cerfontain and van Aken, 1956), and
this indicates the instability of the C — Hg bond. It is possible to produce
Hg203-labeled p-MB by this reaction. This problem has assumed a good
deal of importance in diuretic action and will be discussed more fully in
this connection.
The synthesis of p-MB has been described by Whitmore and Woodward
(1941). It may be purified by repeated solutions in dilute NaOH and preci-
pitations with excess HCl (Boyer, 1954). For the accurate titration of SH
groups it is suggested that the purity of the 2>-MB be checked by iodometric
titration or spectrophotometrically by absorption measurement at 232 m//
at pH 7 (£.v = 1-69 X 10*) or 234 m// at pH 4.6 (f^ = 1-74 x 10*). The
stability of 39-MB solutions has not been determined quantitatively, but
Cunningham et al. (1957) found that heating to 80o-82o for 90 min with
various buffers and at different pH's does not destroy more than 2-4%,
and MacDonnell et al. (1951) stated that solutions are stable for a month
at room temperature. Nevertheless, I would advise making solutions daily
for accurate work.
746 7. MERCURIALS
Reactions of the Mercurials with SH Groups
The various types of mercaptide which can be formed are indicated in
the accompanying tabulation. The complexes formed under particular cir-
Monofunctional
Bifunctional
organic mercurials
inorganic mercurials
R'-Hg
Hg-^
Monothiols
R-S-Hg— R'
R-S— Hg'
R— SH
R— S-Hg-S— R
<;>
Dithiols
S-Hg
"■SH
S-Hg-R'
R
^S— Hg— R'
S-Hg
S-Hg-S^
R R
^S-Hg— S^
(—R— S-Hg-S— ),,
cumstances will depend on the relative concentrations of thiol and mercurial,
the presence of ligands capable of complexing with the mercurials, the spa-
tial arrangement of the SH groups, the pH, and the nature of the R and R'
groups. Monofunctional mercurials, such as p-MB and PM, react with
cysteine, glutathione, and 2-mercaptoethanol in 1 : 1 molar ratio to form
R — S — Hg — R' type of complexes (Benesch and Benesch, 1952; Hoch and
Vallee, 1960). These reactions can be conveniently followed polarographi-
cally. Reactions with dithiols are more complex. The dimercaptide formed
from PM and dimercaprol (BAL) is insoluble, but if hydrophilic groups
occur on the mercurial the product is usually soluble. However, the in-
stability of the C — Hg bond may allow further reaction to form the cyclic
mercaptide, as occurs with mersalyl (Benesch and Benesch, 1952). This
reaction, where R = — CH2CONH — 9? — OCHgCOO", leads to the splitting
CHp— OH CHp— OH
I /OCH3
CH— S-Hg-CHaCH^ _ + H
I ^
I /OCH3
CH2— S— Hg— CH2CH
^R"
Hg— CH2CH(OCH3)R
CH2=:CHR"
CH.OH
CHEMICAL PROPERTIES 747
off of one mersalyl and the formation of inorganic mercury from the other,
essentially the reverse of the reaction whereby the mercurial diuretics are
synthesized (the oxymercuration of alkenes). Such reactions presumably do
not occur with the simpler organic mercurials. It has also been shown that
diphenethjTiyl mercury and glutathione react to form GS — Hg — SG and
phenylacetylene (Tanaka, 1961). Mercurials of the type R2Hg might be
expected to be unreactive with SH groups; inasmuch as they are quite
toxic, Webb et al. (1950) studied their reactions and found that, although
most thiols are not attacked, dithiozne is reacted as follows:
R— Hg— R + R'— SH -> R— Hg— S— R' + RH
This type of cleavage of the C — Hg bond occurs at physiological temperature
and pH.
The primary products of the reactions between HgClg and cysteine, gluta-
thione, thioglycolate, and other monothiols are the dimercaptides of the
type R — S — Hg — S — R, and it is difficult to study the initial formation of
a monomercaptide R — S — Hg+ (Shinohara, 1935; Stricks and Kolthoff,
1953; Stricks et al., 1954). The reactions between HgClg and dithiols are
complex and several types of mercaptide may be formed, as indicated in
the tabulation above. The occurrence of cyclic mercaptides and polymer-
captide linear complexes will depend mainly on the spatial configuration of
the SH groups. The pH apparently plays some role in determining the na-
ture of the complexes, since as the pH increases above 2.5, more of the
forms Hg2(SG)2 and Hg3(SG)2 appear (Kolthoff et al., 1954).
We next turn to the problem of the stability of mercaptides and it is im-
portant to establish the dissociation constants for the fundamental com-
plexes formed in simple reactions, such as the following:
R— S- + Hg++ ±^ R— S— Hg+
R— S- + R'— Hg+ ±5 R— S— Hg— R'
Table 7-6 shows a few of the recently determined constants for Hg++ and
MM. If one assumes that -pK^ and p^g ^I's similar in magnitude, which is
reasonable, it is seen that pJ^j (which applies to the reactions above) lies
between 20 and 22, with a mean value of 21.3. This range may thus be
taken provisionally as indicating the usual affinity between mercurials and
thiols in the absence of competing protons and ligands. Comparison of these
values with those in Tables 7-4 and 7-5 shows that the affinity of mercurials
for SH groups is far greater than for any other single ligands, and that, in a
mixture of thiols and various other complexing ligands, a mercurial will be
predominantly associated with the thiols. The variations of pK with the
temperature and the ionization of auxiliary groups on the thiol are shown,
as summarized from the studies of Stricks and Kolthoff (Table 7-7). Proto-
748
7. MERCURIALS
Table 7-6
Some Dissociation Constants for Simple Mercaptides'
Mercurial
Thiol
pA',
Vth
Reference
Hg++
Cysteine^
20.1
_
Simpson (1961)
20.5
—
Perkins (1953)
—
43.57
Stricks and Kolthoff (1953)
Glutathione-
—
41.58
Stricks and Kolthoff (1953)
Thioglycolate~
—
43.82
Stricks et al. (1954)
Methyl-Hg+
Cysteine"
15.7
—
Simpson (1961)
Human seralbumin
22.0
—
Hughes (1957)
Bovine seralbumin
22.6
—
Hughes (1957)
Bovine HbOj
22.1
—
Hughes (1957)
Bovine HbCO
22.6
—
Hughes (1957)
" The p/^i's for the mercaptides with seralbumin and hemoglobin have been recal-
culated (page 755), assuming pK„ for the SH groups to be 8.7 and the pK for complex-
ing with I to be 8.7. These values should not be considered as accurate because of
the assumptions involved.
Table 7-7
Dissociation Constants for Mercaptides with Hg++ Illustrating
THE Effects of pH and Temperature
Cysteine
Constant"
12°
25°
pA'i
41.82
40.25
pK,
45.27
43.60
P^3
45.40
43.57
pA%
7.39
7.10
VK,
10.72
10.48
Glutathione
12°
42.29
43.54
43.47
7.97
9.28
25°
40.96
41.92
41.58
7.85
9.15
Thioglycolate
12°
45.66
45.85
45.52
25°
44.31
44.33
43.82
" The constants are defined as follows:
K^
(Hg++) (R)^
(HgR2++)
K,=
(H+) (HgR,+ )
(HgR,H++) '
(Hg++) (R) (R-)
(Hgi22+)
(H+) (HgR,)
(HgR,H+)
K. =
(Hg++) (R-)^
(HgR,)
where R = +H3N — X — S~ and R~ = N2H — X — S for cysteine and glutathione, and
R = HOOC— X— S- and R- = "OOC— X— S" for thioglycolate. (From Stricks and
Kolthoff, 1953; Stricks et al, 1954.)
CHEMICAL PROPERTIES 749
nation of an amino group on cysteine or glutathione reduces somewhat the
affinity of the thiol for Hg++, as might be anticipated, while ionization of
the thioglycolate carboxyl groups has little effect. With the constants in
Table 7-7 it is possible to predict the relative concentrations of the various
species present; when complexing ligands are in significant concentration,
appropriate corrections must be made (page 737). The variations with tem-
perature allow the calculation of certain basic thermodynamic parameters.
For the formation of the mercaptides, /iF is — 55 to — 59 kcal/mole and
the entropy changes are positive and usually rather large; for the equilibria
expressed by y>Ki (see legend in Table 7-7), AS is +27 cal/deg for cysteine,
-|- 54 cal/deg for glutathione, and + 68 cal/deg for thioglycolate.
We must now inquire into the effect of pH on mercaptide formation and
particularly consider the reactions with the SH group and the ionized S^
group. Taking the two following equilibria:
R— Hg+ + R'— S- ±^ R— Hg— S— R' K^
R— Hg+ + R'— SH ±^ R— Hg— S— R' + H+ K^„
(R— Hg^)(R--S^)
(R— Hg— S— R')
(R-Hg+) (R--SH)
(R_Hg-S-R')(H+)
and the ionization of SH — i.e., K,, - (H+) (R'— S-)/(R'— SH) — it is
easy to show that:
pATg = pK^^ + pK, (7-2)
The p^^ for SH groups varies from 7 to 10, and in Volume I a mean value
of 8.7 was assumed for protein SH groups. In any event, pKf- and p^sh
will differ quite markedly. This is, of course, essentiallj- a competition be-
tween H+ ions and the mercurial for the S~ group. At physiological pH, SH
will predominate over S~, and the apparent pK for mercaptide dissociation
will be smaller than those given in Tables 7-6 and 7-7.
Just as H+ competes with the mercurial for the S" groups, so various
complexing ligands may compete with the S~ group for the mercurial.
Despite the fact that the p/^'s for thiols are much greater than for most
other ligands, a very significant effect on the equilibrium may be exerted.
Let us write for the usual reaction of mercaptide formation in physiological
media:
R— Hg— X + R'— SH ±i R— Hg— S— R' + X" + H+
where X represents some ligand such as CI" or 0H~. The equilibrium is
given by:
(R— Hg— X) (R'— SH)
K
(R_Hg-S-R') (X-) (H+)
750 7. MERCURIALS
If we designate the equilibrium with X by ^^=(R— Hg+) (X-)/(R— Hg— X),
this taken in conjunction with the expressions for K^, K„, and K leads to:
pA' = pAg - pA„ - vK, (7-3)
Thus in any experimental situation the observable p^ will be less than the
true p/Cg for the reaction of R — Hg+ and S~ by the sum of ^K„ and \>K^,
each of these expressing the competitions involved. Thus in the work of
Hughes (1957) on the reaction of human seralbumin with MM, a p^ of 4.6
was found; if piif^ = 8.7 and ^^K^ — 8.7 (Table 7-5), one may calculate p^g
to be 22.0, X being I" in this case.
The rates of mercaptide formation increase with pH as would be expected
if the reactive form of the thiol is the ionized R' — S~. It is not so easy to
decide on the reactive forms of the mercurial. It seems unlikely that the
Hg++ or R — Hg+ ions are the only reactive species because of their ex-
tremely low concentrations in most physiological media, and it is possible
that the S^ group makes a sideways attack on the Hg atom utilizing a
pair of the six .s electron pairs to displace the X~ ligand.
It is generally considered that mercurials do not react with disulfide
(S— S) bonds, and there is sufficient evidence that this is true for many
proteins and enzymes at physiological conditions. However, Cunningham et
al. (1957) have shown that p-MB catalyzes the splitting of S — S bonds in
cystine, insulin, and ribonuclease at pH 7 if incubation is carried out at 80°.
At this temperature, this may well be a matter of equilibria between S — S
and SH groups, i.e., between native and denatured forms of the proteins,
with p-MB shifting the equilibria by reacting with the SH groups. It is
quite possible that in certain enzymes the S — S groups exist in a state
where reaction with mercurials is significant, and such a reaction should
not be completely ignored.
Certain metabolically important cofactors, such as coenzyme A and li-
poate, are thiols, and it is of some interest to inquire into whether the
mercurials react readily with them. Surprisingly little quantitative work
has been done and most of the evidence is indirect. For example, Galston
et al. (1955) found that p-MB increases the yield of peroxidase, catalase,
and tyrosinase in plant breis when added to the preparation medium;
since coenzyme A inactivates these enzymes, it was assumed that p-MB
protects the enzymes by forming a mercaptide with the coenzyme A. In
coenzyme A-deficient rats, the toxicity of mercurials is increased, and the
mercurials inhibit the coenzyme A-dependent acetylation of sulfanilamide
(Leuschner et al., 1957). Mersalyl and HgClg reduce the coenzyme A level
of yeast 25% at 22.5 mM and 0.3 mM, respectively (Estler et al, 1960).
Sanner and Pihl (1962) followed the reaction between 2?-MB and coenzyme
A by changes in the absorption at 255 m/^ and showed that the thiol could
be titrated by the mercurial. Turning to DL-a-lipoate, one finds that its
REACTIONS WITH PROTEINS 751
administration prevents mercurial poisoning in mice, and that it reduces
the inhibition of pyruvate oxidase by HgClg at high concentrations (Gru-
nert, 1960; Grunert and Rohdenburg, 1960). It is interesting that with
lower concentrations of HgCla (1.3 mM), lipoate increases the inhibition of
pyruvate oxidation in intact cells of Streptococcus faecalis, the mercaptide
possibly entering the cells more readily than the complexes of Hg++. It
would thus appear that mercurials react with coenzyme A and lipoate,
but how rapidly and how tightly are not known.
Finally, we consider the problem of the splitting of thioesters by the
mercurials. Sachs (1921) showed that acetylthioethyl esters are rapidly
split by HgClg to acetate and mercurothioethanol, and in general all acyl
mercaptans seemed to behave in this way. Thus Lynen et al. (1951) in
their early studies on active acetate and coenzyme A investigated acetyl-
CoA and found it to be split by approximately 100 niM Hg acetate, a result
of questionable significance in physiological work. Stern (1956) studied ace-
toacetyl-CoA, a thioester of importance in lipid metabolism and possessing
a strong absorption maximum around 303 m//, and found that HgClg at
concentrations higher than 0.001 mM produces a rapid decrease in this
absorption, 0.1-0.2 mM completely abolishing it, this corresponding to
about a 1 : 1 molar ratio between Hg and thioester. The following reaction
sequence was suggested:
Hg++ + AcAc— S— CoA- -> Hg— AcAc— S— CoA+
Hg— AcAc— S— CoA+ + H2O -> Hg++ + AcAc" + HS— CoA
Hg++ + HS— CoA -> Hg— S— CoA+ + H+
The initial reaction is apparently with the enolate group of the acetoacetyl
radical. Stern believes that this reaction, occurring at such low concentra-
tions of Hg++ and so rapidly, may well be of great importance in the effects
of the mercurials on metabolism. However, Gibson et al. (1958) reported
that p-MB does not react with or split succinyl-CoA at a significant rate,
and Sanner and Pihl (1962) found a variety of thioesters to be resistant to
p-MB (e.g., acetyl-CoA, succinyl-CoA, and benzoyl-CoA). From this limited
work one might conclude that HgClj can split some thioesters but that ^^-MB
cannot. But Vagelos and Earl (1959) found that, in contrast to most thio-
esters, malonyl semialdehyde pantetheine reacts readily with p-MB, and
proposed that /5-carbonyl thioesters may be susceptible. The possible role
of these reactions in metabolic inhibitions is at present unknown.
REACTIONS WITH PROTEINS
It was believed in years past that mercury, in common with other heavy
metals, is adsorbed onto proteins, denaturing and precipitating them, but
recent work has shown that under appropriate conditions stoichiometric
752 7. MERCURIALS
combinations of mercury with proteins occur, and that denaturation and
precipitation are by no means a general phenomenon. These definite com-
plexes in most cases are formed through the SH groups of the proteins, and
methods whereby these groups may be titrated quantitatively with the
mercurials have been devised. Some may feel that a discussion of the com-
plexes of mercury with the proteins is out of place in a book on metabolic
inhibition, but actually much can be learned from the thorough and illu-
minating investigations on mercaptalbumin, hemoglobin, and other pro-
teins reported in the past few years. One must also realize that in any
system containing nonenzymic proteins, particularly cellular preparations,
reaction of mercury with these proteins not only may have definite effects
on the enzyme inhibition, but may be responsible, at least in part, for me-
tabolic or functional changes.*
Protein Groups Reacting with Mercurials
The mercurials react rapidly with certain free and exposed protein SH
groups, more slowly with others, and not at all with some which are pre-
sumably buried within the protein structure or otherwise sterically unavail-
able (page 643). Many SH groups react only after denaturation of the pro-
tein, a process which apparently exposes them for attack by the mercurials.
Since mercurials often initiate denaturation, they may produce a progressive
unloosening of the protein structure and themselves make available SH
groups originally unreactive, the process continuing until it is irreversible,
the reformation of the normal configuration not being possible when the
mercurial is removed. Much work with proteins and enzymes, to be dis-
cussed later, provides evidence that SH groups are the primary site of
mercurial binding, and often the only site under certain conditions. The
problem to be considered here is whether protein groups other than SH
can under any circumstances contribute to the binding.
Examination of the constants for the complexing of mercurials with SH
groups and with groups normally present in proteins leads to the immediate
* There is some inherent and unavoidable difficulty and ambiguity in the terminology
of the mercurials. If one uses the inorganic HgClj, how should the inhibitor be desig-
nated — as HgCU, Hg, Hg++, Hg", or otherwise? We have seen that in most, if not
all, media the mercuric ion will exist in a variety of complexes, probably of different
reactivities, so that it is impossible to designate the situation accurately. Similar
problems arise with other heavy metal ions, e.g., copper and zinc. We shall, therefore,
arbitrarily designate inorganic divalent mercury as Hg++, without implying that
this is either a predominant form or an active form. It is fundamentally and finally
the Hg++ ion which complexes and reacts with the various substances present, so that
when this is so written it must be understood that all of the complexes are implied.
The designation of the organic mercurials as 79-MB, PM, etc., similarly is noncommittal
with respect to the forms present or active.
REACTIONS WITH PROTEINS 753
conclusion that no single group can compete very effectively with the SH
groups for the mercurials. Indeed, these other groups probably have their
orbitals occupied by competing with the various ligands present in the me-
dia. However, there are at least three factors which must be taken into
consideration. (1) Certain fortuitous arrangements of two or more non-SH
groups, perhaps allowing chelation of the mercurial, can increase the affin-
ity markedly, as seen in Table 7-4. It is quite possible that occasionally
such situations occur on protein surfaces, although generally the opportu-
nities for successful chelation must be rare; e.g., the binding to glycylglycine
is less than to glycine, and increase in the length of the polypeptide chain
wiU probably reduce affinity except in very special cases. (2) The most
reactive SH group or groups on a particular protein may not happen to
have a strong affinity for a mercurial, due to steric factors or an unfavorable
electric field, so that non-SH groups can compete more effectively. Although
there is little quantitative evidence, one gets the impression that usually the
SH groups of proteins do not bind most mercurials as tightly as do the SH
groups of simple thiols, such as cysteine or glutathione, especially since
one can often remove a mercurial from a protein quite readily by adding
one of these thiols. (3) As pointed out by W. L. Hughes (1950), mercurials
will complex with non-SH groups when the SH groups become saturated,
or actually before in many cases. Since excess mercurial is often present,
especially in enzyme studies, such secondary complexes must be considered,
even though the SH groups are reacted first.
On the experimental side, it has been observed that some proteins bind
more Hg++ than corresponds to the SH content and that some of this is
relatively weakly bound. Haarmann (1943 a) found that with increase in
the pH progressively more Hg++ is bound to various proteins, although
only a fraction is really tightly attached to the protein, and postulated that
CONH groups might bind Hg"+. More recently, Perkins (1958, 1961) re-
ported the binding of 104-130 g-atoms of Hg/10^ g seralbumin and, follow-
ing treatment with bromoacetate (blocking SH and amino groups), the
binding increased to 190 g-atoms/10^ g protein at pH 5.5. The SH groups
could have accounted for only 1 g-atom/10^ g protein and Perkins felt that
numerically the only possible binding sites are the C00~ groups. This is
a situation in which there is excess Hg++ present for reaction with non-SH
groups and, inasmuch as the dissociation constants are not known, it is
impossible to compare the affinities for the various groups. Nevertheless,
these results conclusively establish the non-SH binding of mercurials under
certain conditions, and it would be well to bear this in mind in enzyme
inhibition work.
Examples of Reactions with Specific Proteins
(A) Ovalbumin. Although not much work has been done with this pro-
tein, the results illustrate some of the problems one encounters. Anson
754
7. MERCURIALS
(1941) found it difficult to determine if p-MB reacts with ovalbumin or
not, since ferricyanide and nitroprusside cannot be used to detect these
SH groups (they do not react with ovalbumin) and iodine oxidizes the
group whether free or combined with p-MB. Using an indirect method in-
volving the determination of the binding of p-MB to cysteine in the pres-
ence of ovalbumin, Anson claimed that either p-MB does not react at all
with the ovalbumin SH groups, or, if so, the binding is much less strong
than with cysteine. If the ovalbumin is denatured, demonstrable binding
of the mercurial occurs. W. L. Hughes (1950) reported that MM reacts
with ovalbumin very slowly, the reaction requiring a day to come to equi-
librium. However, MacDonnell et al. (1951) obtained a crystalline derivative
of ovalbumin treated with 59-MB by adjusting the pH to 4.7 and adding
ammonium sulfate to opalescence. Three of the four SH groups of oval-
bumin react and the fourth reacts only after denaturation. The complex
is very stable and is not dissociated by cysteine or prolonged dialysis, al-
though 6-day dialysis against cysteine at pH 7.9 dissociates about 75% of
EQUIVALENTS
AMOUNT OF MERCURIAL ADDED
Fig. 7-3. Theoretical curves for the reac-
tion of a mercurial with a protein, such as
ovalbumin with three reactive SH groups.
See text for explanation.
the mercurial. In such cases as this, if one plots the amount of mercurial
combined, or the disappearance of SH groups from the protein, against the
amount of mercurial added, one may obtain different curves (Fig. 7-3).
If the reactions with the free SH groups are equivalent, a linear relation-
ship to saturation will be obtained (curves A and B), but if interaction be-
tween the SH groups occurs (i.e., if the binding of a mercurial reduces the
binding of the next) or the SH groups combine with the mercurial with
different affinities, a curve concave downward will be obtained (curve C).
If no further reaction with protein groups occurs after saturation of the
SH groups, the curve will be horizontal (curve A), but if other groups with
REACTIONS WITH PROTEINS 755
less affinity for the mercurial continue to react, a sloping or curved line will
be obtained (curve B). It is important to construct such curves whenever
possible in order to understand the binding characteristics.
(B) Hemoglobin. Crystalline human oxyhemoglobin reacts with 2 moles
of MM per mole of protein but the rate is rather slow at pH 7.5 (W. L.
Hughes, 1950). The hemoglobins of other species may contain either two
or four SH groups that react readily. Green et al. (1954) crystallized a de-
rivative of horse hemoglobin in which two SH groups had been reacted
with p-MB and showed that, although the crystals are isomorphous with
normal hemoglobin, the X-ray diffraction pattern is somewhat different.
Ingram (1955) established that Hg+^, MM, and p-MB all combine readily
with four SH groups of horse hemoglobin in the native state, and with six
in the denatured form; ox and human hemoglobins are similar but the
latter presents eight SH groups when denatured. In native hemoglobin,
Hg++ probably reacts with two SH groups simultaneously; at least 2 equiv-
alents of Hg+^ reduce the free SH groups to zero. However, p-MB like-
wise blocks two SH groups and since reaction of a molecule of p-MB with
two SH groups is impossible, it is likely that a pair of SH groups is so close
Fig. 7-4. The reactions of the SH group
pairs on hemoglobin with Hg++ and p-MB,
according to the concept of Ingram (19.55).
that each group cannot react with the large mercurial. It is difficult to say
in the case of Hg++ if the effect is steric or due to the formation of a bridge
between the two SH groups in a pair, but the latter mechanism is favored.
The situation as represented by Ingram is shown in Fig. 7-4. It is interesting
that Hg++ and p-MB compete for the SH groups, and that the former is
bound more tightly, probably due to the reaction with two of the SH groups.
Since the earliest studies of hemoglobin SH groups, there has been diffi-
culty in establishing the exact number of reactive and unreactive groups,
due to the fact that the nature of the reactions and the stoichiometry were
uncertain. The details are not pertinent to our purpose and have been well
reviewed by Huisman (1959). Further complications have arisen in the re-
cognition that different types of hemoglobin react differently with the mer-
curials and that the pH is an important factor. Murayama (1958) claimed
756 7. MERCURIALS
that adult hemoglobin contains two SH pairs, sickle cell hemoglobin three
SH pairs, and hemoglobin C no pairs, while Huisman obtained quite differ-
ent results indicating more reactive SH groups than were found by most
previous workers. At pH 7, adult hemoglobin reacts with 3 p-MB molecules
and fetal hemoglobin with 2, whereas at pH 4.6 the values are 6 and 4,
respectively, the lowering of the pH presumably altering the configuration
of the protein. Studies of mercaptalbumin show that Hg++ can induce the
formation of dimers of the protein, but Allison and Cecil (1958) believe
that only monomers occur in the case of hemoglobin, and found that Hg++
and PM give the same titer, while the results of Cecil and Snow (1962) are
more in accord with those of Ingram, 2.2 reactive SH groups of a total of
6 in adult hemoglobin being detected, the 3.8 sluggish SH groups reacting
differently with Hg++, PM, and p-MPS. These studies of hemoglobin not
only show the influence of many factors on the interaction of proteins with
mercurials, but point out the difficulties of SH titration of even relatively
simple proteins.
Reaction of hemoglobin with mercurials brings about striking changes in
the characteristics of oxygenation: The affinity of hemoglobin for Og may
be unaffected or increased, but the interactions between the heme groups
are reduced or abolished by all the mercurials (A. F. Riggs, 1952; Wolbach
and Riggs, 1955; Riggs and Wolbach, 1956; A. Riggs, 1959). Mersalyl and
Hg++ at pH 6.8 increase the affinity for 0^ quite markedly, Pjq decreasing
to about one third of normal, but p-MB and MM do not alter the affinity.
AU these mercurials lower the interaction constant n from around 2.9 to 1
(or near 1), the latter corresponding to complete loss of heme-heme interac-
tion. Glutathione completely reverses these effects. It is very interesting
that the maximal effects are produced at ratios close to 2 moles of mer-
curial to 1 mole of hemoglobin; however, as the amount of mersalyl is in-
creased, the changes in heme interaction and Og affinity progressively dis-
appear, so that at a ratio of 15-16 moles of mercurial per mole of hemoglobin
there is no longer an effect. This curious reversal is unexplained. The ob-
servation may have some bearing on the use of mercurials for enzyme inhi-
bition, and Riggs and Wolbach (1956) state, "Our observations suggest
that the attempt to inhibit an enzyme with only a single high concentration
of mercurial may lead to spurious conclusions." I know of no example in
which enzyme inhibition is lost at higher mercurial concentrations, but in
any case it would presumably be a rare phenomenon. Occasionally one finds
stimulation of enzyme activity at low mercurial concentrations and this
reverses to inhibition as the concentration is increased, but it is not known
if this has any relation to the above reversal. Mercuration of oxyhemoglobin
increases the rate constants for the dissociation of Og from three of the
hemes, but decreases the rate constant for the dissociation of the last Og,
this being in fair accord with the effects on the Og dissociation curve ob-
REACTIONS WITH PROTEINS 757
served by Riggs (Gibson and Houghton, 1955). Oxygenation of hemoglobin
facilitates reaction of the SH groups with A^-ethylmaleimide and iodoaceta-
mide but not with p-MB (Benesch and Benesch, 1962). The mechanisms
by which these effects are produced are not clear, but hypotheses have been
offered based on the spatial arrangement of the SH groups and the hemes.
Riggs (1959) considers the hemoglobin molecule to consist of two halves,
each with a pair of reactive SH groups and a pair of hemes, the SH groups
perhaps lying between the hemes. Any mercurial which can form a bridge
between the 2 SH groups of a pair — such as Hg++, or mersalyl if the
C — Hg bond is ruptured and inorganic Hg++ is released — increases the
affinity of the hemes for Og, whereas mercurials reacting only with a single
SH group — such as p-MB and MM — do not have this effect. Any interac-
tion with the SH groups reduces the interaction between the hemes, prob-
ably by bringing about reversible structural changes in the protein config-
uration. On the other hand, Klotz and Klotz (1959) favor a mechanism
involving disturbances in the water structure around and between the
hemes. Whatever the explanation, the bearing on the effects of mercurials
on enzyme active centers by reaction with adjacent SH groups is obvious.
(C) Mercaptalbumin. Mercaptalbumin is one fraction of the serum al-
bumins containing a single reactive SH group whereas the other albumins
contain none. It was isolated as the crystalline mercury salt by Hughes
(1947) and its reactions have been studied in detail, so that it has become
the classic example of protein-mercurial interaction (W. L. Hughes, 1950).
The three major reactions maj^ be represented as follows:
Reaction 1: alb— SH + RgX^ ±5 alb— S— HgX + H+ 4- X"
Reaction 2: alb— S— HgX + alb— SH ±^ alb— S— Hg— S— alb -f- H+ -^ X-
Reaction 3: alb— S— Hg— S— alb + RgX^ ^ 2 alb— S— HgX
where alb indicates mercaptalbumin and X some ligand (e.g. Cl~). The
first reaction is mercaptide formation, the second dimerization, and the
third dissociation of the dimer by excess HgXg. Formation of the dimer
increases the turbidity and, if some ethanol is added, crystals form. These
crystals are colorless diamond-shaped orthorhombic plates, containing chan-
nels or enclosures of fair size, with liquid within them, and permeable to
various salts, sugars, and dyes (Low and Weichel, 1951). Hughes and Dint-
zis (1964) have described procedures for crystallizing the dimers from etha-
nol-water mixtures at low temperatures. Viscosity and sedimentation stud-
ies (Low 1952) led to the representation of the dimer as in A of Fig. 7-5,
while the results of X-ray diffraction study are compatible also with struc-
ture B. The lengths of the dimer would be around 140-150 A, monomer
mercaptalbumin being of molecular weight 66,000. The structure is inde-
pendent of the smaller ions making up the crystal; e.g., the dimer will
758
7. MERCURIALS
form crystals with Hgl^g, the interactions being purely electrostatic and
not involving SH groups (Lewin, 1951).
Reaction 1 is quite rapid, but reaction 2 is slow because it involves two
large molecules of similar charge. The dimerization requires about 25 min
for half-reaction and 2 hr for equilibrium when Hg++ is mixed with mer-
~D
Fig. 7-5. Possible forms of the Hg-mercaptalbumin dimer.
The small solid circle represents Hg++. (From Low, 1952.)
captalbumin in a 0.5 : 1 ratio (Edelhoch et al., 1953). Dimerization is an
endothermic reaction, rise in the temperature favoring formation of the
dimer, AH^ being about 7 kcal/mole. The temperature effect indicates
an activation energy for dimerization of 17-21 kcal/mole. The constant
for the equilibrium (alb— S— HgCl) (alb— SH)/(alb— S— Hg— S— alb) was
found to be 3.2 X 10^^^ at pH 4.5 and 25°. Reaction 1 is reversible by sub-
stances forming stable compounds with Hg++, and reaction 2 is reversible
by ligands forming HgX„ complexes. Dimerization is also reversed by reac-
AMOUNT
OF DIMER
(TURBIDITY)
0 0.5
MOLE Hg/MOLE PROTEIN
Fig. 7-6. The formation and dissociation
of the Hg-mercaptalbumin dimer as the
molar ratio of Hg++ to protein is in-
creased. (From Edelhoch et al., 1953.)
tion 3 (Fig. 7-6), Hg++ competing with mercaptalbumin for the alb — S — Hg
monomer. The dissociation of the dimer by cysteine is very rapid (appar-
ently within a few seconds) and yet the dimer is not split nor the Hg++
dissociated readily by dialysis, so that this would seem to be one of those
interesting situations in which a complexer appears to take the metal from
the protein rather than merely combining with free metal ions (Straessle,
REACTIONS WITH PROTEINS 759
1954). The disulfide dimer, alb — S — S — alb, however, is dissociated by cys-
teine very slowly. The dimerization is accomparied by an increase in levoro-
tation, and this implies that the mercaptalbumin molecule undergoes some
unfolding in the region of the reactive SH groups as a necessary prelude
to dimerization; this may be thought of as a partial denaturation, adding
one more item of evidence for configurational changes induced by mercuri-
als (Kay and Marsh, 1959).
Organic mercurials such as p-MB, PM, and MM react in a 1 : 1 ratio with
mercaptalbumin and, of course, no dimer is formed. The equilibrium con-
stant for the reaction:
alb— SH + CH3— Hgl ±5 alb— S— Hg— CH3 + H+ + Cl-
has been found by \V. L. Hughes (1950) to be 3.5 x lO^^ (p^ = 4 45)^
from which the value for the dissociation constant of the mercaptide in
Table 7-6 was calculated. On the other hand, bifunctional organic mercuri-
als, such as:
CH-O
XHg-CH2— HC CH— CH2— HgX
O— CH,
can link two mercaptalbumin molecules together (Straessle, 1951; Edsall
et al., 1954). The pA" for the equilibrium (dimer )/(monomer+) (alb") is 18.2
at pH 4.75 and 25°, the corresponding p^ for the Hg++ dimer equilibrium
being 13.5.* This difference of some 4.7 p/C units between the two dimers
is undoubtedly due to the fact that the mercaptalbumin molecules must
approach about 10 A closer in the Hg++ dimer than in the bifunctional
mercurial dimer, and the steric and electrostatic factors could easily ac-
count for the some 6.7 kcal/mole difference.
The importance of these results with mercaptalbumin for inhibition stud-
ies with the mercurials is clear. First, the possibility of the dimerization
of certain enzymes by Hg++ leads to the concept that inhibition may oc-
casionally result through steric sequestration of the active sites and not
necessarily through the reaction of SH groups at the active sites. It is also
possible that occlusion of active sites could occur by linking a nonenzyme
protein to the protein through a Hg bridge. It is known that Hg++ often
produces an increase in turbidity of enzyme solutions and even precipita-
tion, although this can be due to other factors as well. Second, this reason-
ably well understood and quantitatively investigated system provides a
model on which the effects of ligands, pH, temperature, and other factors
* Gurd and Wilcox (1956) give values of 17.2 and 12.6 for the p/C's, respectively,
due to different assumptions regarding the ligand constants.
760 7. MERCURIALS
may be better, appreciated. Third, it presents one clear instance in which
mercurials react specifically with protein SH groups, since there is no evi-
dence that other groups are even involved in the mercuration of mercaptal-
bumin. Last, it is significant in the use of mercurials in whole animals that
the serum contains sufiicient mercaptalbumin to bind most, if not all, the
mercurial present, a factor that must be considered in the penetration, dis-
tribution, and actions of the mercurials in animals.
Effect of pH
The competition between H+ and the mercurial for the S" group and the
effect this has on the over-all equilibrium have been discussed. On the basis
of only the ionization of the SH group, one would predict that mercurials
would react more rapidly and more completely at higher pH's (particularly
above p^^'s of the SH groups). However, there are many other factors
which may be important. Actually, it has generally been observed that the
rate of reaction of p-MB with proteins is decreased with a rise in the pH.
Both the extent and rate of reaction of p-MB with ovalbumin are affected
by pH: At pH 4.6, 4 moles of p-MB react rapidly with 1 mole of protein,
whereas at pH 7, only 3.2 moles of p-MB react in 24 hr (Boyer, 1954).
Reduction of the rate with increasing pH has also been reported for /5-
lactoglobulin and 3-PGrDH (Boyer and Segal, 1954), it being much faster
at 4.6 than at 7. The work of Huisman (1959) with hemoglobin illustrates
an important point; the rate and extent of reaction may be influenced dif-
ferently by pH, inasmuch as the rate of mercaptide formation is faster be-
tween 7 and 11.2 than at 4.6, but more SH groups are reactive at 4.6. One
also recalls that p-MB is dissociated more rapidly from ovalbumin at pH
7.9 than in the acid pH range (MacDonnell et al., 1951). Another factor
often overlooked is the effect of pH on the secondary denaturation of the
protein following mercuration. The rate of thermal denaturation of seral-
bumin is increased 13.2-fold at pH 3.6, while at pH 7 there is no denatura-
tion by Hg++ at 1.85 mM, while denaturation of /?-lactoglobulin is increased
89-fold at pH 3.6 and not at all at pH 7, this indicating that an acid pH
favors secondary configurational changes resulting from binding of the Hg++
(Stauff and Uhlein, 1958). The reaction of mercurials with non-SH groups
is also pH-dependent, since the complexing of the azomercurial studied by
Horowitz and Klotz (1956) with glycine is maximal between pH 6.5 and
9.5; at low pH's the +H3N — CH2 — C00~ form of glycine dominates and
is less reactive, while at high pH's there is competition by 0H~.
R-Hg-00C-CH3 + H2N-CH2-COO- ±? R-Hg-OOC— CH2-NH2 + CH3-COO-
The following factors, in addition to the ionization of SH groups, must
thus be considered. ( 1 ) The pH may vary the number of reactive SH groups
REACTIONS WITH PROTEINS 761
or their individual reactivity, by effects on the structure of the protein, or
on the association of subunits. (2) The pH will determine the over-all charge
on the protein; e.g., with increased pH the protein will become more neg-
atively charged and possibly repel negatively charged mercurials, such as
p-MB, p-PMS, or the higher Ch complexes with Hg++. (3) Rise in pH will
increase the OH" concentration and this ion will compete with the SH
groups for the mercurial. (4) The pH may determine the degree of hydrogen
bonding of SH groups and thus their reactivity with mercurials. It is likely
that the dependence on the pH will depend on the mercurial used, but in-
sufficient data are available for comparisons.
A final effect of pH involves dimerization where it occurs. The rate and
degree of dimerization in the presence of Hg++ will depend on the total
protein charge, being maximal at the isoelectric point, all else being equal.
Straessle (1951) reported that the dimerization of mercaptalbumin with a
bifunctional mercurial is slower at pH 6 than at 4.75, and Edelhoch et al.
(1953) found the rate of dimerization with Hg++ to be increased 60 times
when the pH is decreased from 6 to 4.75, and doubled with further de-
crease to 4.25. It was calculated that a charge of 9 charge units would ac-
count for this, and titration data indicated a change of 10 units over this
pH range, so the electrostatic mechanism seems to be correct.
Effects of Mercurials on Protein Structure and Properties
The importance of secondary changes in protein structure upon reaction
with a mercurial cannot be overemphasized in studies of enzyme inhibition
and its reversibility, but unfortunately little exact information is available.
Configurational changes have been postulated to explain certain results,
such as have already been mentioned in regard to mercaptalbumin (page
757) and hemoglobin (page 755), and additional examples wiU be presented
in connection with enzyme inhibition, but in most instances the evidence is
indirect and tenuous. Nevertheless most investigators agree, I believe, in
accepting that such changes occur in certain cases; the problems are the
nature of the changes and the mechanisms by which they are induced.
Higher concentrations of Hg++ and most organic mercurials decrease the
solubility of proteins, and may precipitate or coagulate them. This gave
rise to the early concept of the mercurials as denaturing agents. However,
it would appear that the primary effect is seldom denaturation (in the sense
of disruption of the polypeptide chain structure), and that the altered prop-
erties of the protein are more directly related to modification of side-chain
groups and the introduction of new groups. Prolonged contact of proteins
with mercurials occasionally leads to true denaturation as a secondary reac-
tion, but complete reversibility can usually be achieved by removing the
mercurial; this indicates that if structural changes occur they are probably
localized, and that the normal configuration can be restored. Such direct
762 7. MERCURIALS
structural effects should be distinguished from preferential reaction of mer-
curials with denatured protein in cases in which native and denatured pro-
tein exist in equilibrium under nonphysiological conditions (e.g., at low pH's
or high temperatures) (Habeeb, 1960). They should also be distinguished
from changes brought about in proteins evident after precipitation. The
thermal coagulation of serum proteins is enhanced by p-MB and the coagula
are firmer, more elastic, and more transparent, the water bound to the clot
being around 4 times greater (Jensen, et al., 1950), but this does not pro-
vide evidence that protein configuration before coagulation is altered by
the mercurial. High Hg++ concentrations weaken keratin fibers so that they
break under less tension (Hoare and Speakman, 1963). This would be ex-
pected if interchain disulfide bonds are disrupted. Changes in gross protein
properties seldom provide information on the more important and subtler
localized modifications which are believed to occur.
In view of the significance of configurational changes in enzyme inhibi-
tion, and in the belief that more examples will be postulated and established
in the coming years, we may summarize some of the possible mechanisms
by which such effects can be brought about. (1) In those cases in which
there are equilibria between SH and S — S groups, or where there is a cyclic
oxidation and reduction, and in which the S — S bonds contribute to the
stability of a local configuration, mercaptide formation may loosen the
structure. (2) The SH groups themselves may contribute to the stability,
perhaps by hydrogen bonding or the binding of cofactors, so that mercura-
tion may again enhance dissolution of the native structure. (3) The intro-
duction of a charged group, such as occurs with p-MB or p-MPS, will alter
the local electric field, and this may favor instability. (4) Hg++ and or-
ganic mercurials which dissociate to form Hg++ can bind to two groups
simultaneously and thereby distort protein configuration. (5) Reaction of
mercurials with non-SH groups, especially N- and 0-containing groups,
may reduce hydrogen bonding between polypeptide chains. It is not ne-
cessary that the affinity for these groups be especially high, and there is
some indirect evidence that it is often the excess mercurial, above that
required to saturate the SH groups, which is responsible for denaturation.
There is increasing reason for believing that proteins are not rigid structures
but often exhibit a fair degree of flexibility (page 1-199), so that it is reason-
able that reversible modifications of the structure may be fairly easily in-
duced.
Estimation and Titration of Protein SH Groups with Mercurials
The older methods for the determination of SH groups, using nitroprus-
side, ferricyanide, iodine, or other reagents, are now considered to be gen-
erally unreliable when applied to proteins, due mainly to lack of specifi-
city, and, in addition, these methods are often rather laborious. Ampero-
REACTIONS WITH PROTEINS 763
metric titration with Ag+ is still commonly used and is often useful when
combined with mercurial titration; one should consult Leach (1960) for a
discussion of some of the difficulties of this method. Amperometric titra-
tions at a rotating platinum electrode using Hg++ or MM have been shown
to be accurate and reliable in some cases (Saroff and Mark, 1953; Kolthoff
et al., 1954; Leach, 1960), but these technique have not yet been extensively
applied to enzymes. Leach has listed the requirements for an ideal SH re-
agent for titrations: It should (1) be specific for SH groups, (2) be highly
reactive, (3) have a small molecular size, (4) be preferably monofunctional,
(5) be devoid of charge or other reactive groups so that all protein SH
groups are equally favored despite their different environments, (6) be sol-
uble, stable, and reactive over a range of pH, and (7) show well-defined
reduction steps for amperometric use. MM fulfills most of these criteria and
perhaps it has been neglected in enzyme work. Past work on many pro-
teins has indicated that it is often well to use more than one method in
order to increase the reliability.
The most commonly used method at present is the spectrophotometric
titration with p-MB developed by Boyer (1954), since it is convenient and
appears to be generally accurate. Furthermore, the sensitivity is as high
as with the amperometric methods, namely, around 0.01-0.1 ulM SH.
Reaction of p-MB with SH groups leads to an increase in the absorbancy
at 250 m// at pH 7 (Fig. 7-7); at pH 4.6 the maximal increment occurs at
255 m//, but it is usually preferable to titrate enzymes at the more physi-
ological pH of 7 to determine the number of reactive SH groups, unless
the mercaptide formation occurs too slowly. The increase in absorbancy is
a linear function of the SH groups reacted, for both simple thiols and pro-
teins (Fig. 7-8), but the absorbancy change is somewhat different for dif-
ferent SH groups, a fact of no importance in titrations. The p-MB may be
titrated with protein (as in Fig. 7-8), the end-point being the sharp break
between the two linear segments, or the protein may be titrated with p-MB;
in both cases one determines when all the reactive SH groups are transform-
ed into mercaptides. Although the details of the method may be found in
the original paper of Boyer (1954) and the excellent review of R. Benesch
and R. E. Benesch (1962), it may be useful in interpreting such titrations
applied to enzymes to note briefly certain precautions and difficulties.
(1 ) The only mercurial which can be used is jj-MB, and since it is a rather
large molecule with a negative charge, steric or electrostatic factors may
reduce its reaction with certain SH groups. PM and 7)-MPS exhibit ab-
sorbancy shifts but at lower wavelengths where protein absorbs much more
strongly than at 250-255 m//.
(2) The addition of excess p-MB occasionally results in a further small
increment in the absorbance, so that the flat portion of the curve may not
be exactly horizontal, and this may indicate reaction with non-SH groups
764
7. MERCURIALS
or less readily reacting SH groups. Such behavior seems to be rare and does
not seriously interfere with the determination of the end-point.
(3) The time relations must be considered. It is common practice to in-
cubate the protein and p-MB for 10-15 min to allow reaction, but a decision
as to this depends on one's definition of a reactive SH groups. In some
proteins, additional SH groups are reacted when the incubation is prolonged;
is such cases one is not certain if these groups were initially exposed, or
if they arise during a progressive denaturation of the protein.
Fig. 7-7. The spectral absorption curves for
p-MB and its mercaptide with cysteine, at
pH 7 in 60 raM phosphate buifer. (From
Boyer, 1954.)
(4) Both proteins and p-MB absorb significantly at 250-255 m/^ and the
appropriate controls must be run. For example, when protein is titrated
with p-MB, equivalent increments of the mercurial are added to the blank
cell.
(5) Protein or enzyme solutions should be as pure as possible, since even
small amounts of certain impurities may cause large errors, and the solu-
tions should be clear so that light scattering is reduced.
(6) Special consideration should be given to the pH since it has been
shown that both the rate and extent of reaction are markedly affected, as
in Boyer's experiments with ovalbumin. Although reaction may be more
REACTIONS WITH PROTEINS
765
rapid at pH 4.6 than at 7, it is preferable, as mentioned above, to use as
physiological a pH as possible if the normal state of the protein or enzyme
is to be established.
(7) Salt effects on the reaction of p-MB with proteins are also often of
some magnitude, so that attention must be given to the ionic composition
of the medium and the buffers used. It is preferable in most cases to use as
low concentrations of salt and buffer as possible, if maximal reactivity of
the SH groups is desired, but occasionally it is useful to add some salt,
such as KCl, to reduce the p-MB reactivity in order to eliminate non-SH
group effects.
0.5
0- LACTOGLOBULIN
'0 5
MilO
Fig. 7-8. Titration of cysteine and proteins with p-MB at
pH 4.6 in 330 mM acetate medium. The reaction times
are: cysteine and ovalbumin 15 min, and lactoglobulin
20 hr. (From Boyer, 1954.)
(8) It is worth noting that Boyer found EDTA to interfere, presumably
due to a complex with p-MB, so it is advisable to omit this substance.
(9) Masking of the reactive SH groups, e.g. with alkylating agents, abol-
ishes the absorbancy changes on adding p-MB, this providing evidence
that it is indeed the SH groups which are responsible for the changes. Ti-
tration of proteins or enzymes treated with various agents can thus provide
information on the disappearance of SH groups,
(10) The p-MB must be pure, should be analyzed iodometricaUy or spec-
trophotometrically, should be standardized against glutathione (details are
766
7. MERCURIALS
given by Benesch and Benesch), and should be used in freshly made so-
lutions.
(11) The presence of two or more SH groups close together on the protein
may prevent the reaction of each with jj-MB, as is the case with hemoglobin.
This will lead to low values for the total number of SH groups in proteins
or enzymes.
A typical titration of an enzyme is shown in Fig. 7-9. The titration of
3-phosphoglyceraldehyde dehydrogenase at pH 4.6 presents a clear end-
point indicating a rapid reaction of the SH groups. This yields 10.3 SH
groups per molecule of enzyme (assumed molecular weight of 118,000).
0.3
4 5
/i MOLES/ML iio'
Fig. 7-9. Titration of 5 times recrystal-
lized 3-phosphoglyceraldehyde dehydro-
genase with p-MB at pH 4.6 and 0.03
/<mole/ml. (From Boyer and Segal, 1954.)
The reaction of the SH groups occurs more slowly at pH 7 and a sharp
end-point was not obtained by incubations up to 15 min; however, longer
incubations would probably have given a sharp break in the curve. Here
the end-point yields 8.3 SH groups per enzyme molecule, suggesting that
2 SH groups become much more reactive when the pH is lowered. A value
of 10.7 half-cystines per molecule for this enzyme has been reported (Velick
and Ronzoni, 1948), so it is evident that most of these SH groups are free
and reactive.
Colored Mercurials and Histochemical Determination of Protein SH Groups
Various colored mercurials, usually azobenzene derivatives, have been
known for many years but were not applied to biological material until
Bennett (1948 a) studied the reaction of p-mercuriphenylazo-/5-naphthol
with tissue thiols. Direct visualization of thiol distribution in the tissues is
possible, but the dye has a very low solubility in water and at the usual
pH's so low a molecular extinction coefficient that its use is limited. How-
REACTIONS WITH PROTEINS 767
ever, Flesch and Kun (1950) found that the addition of strong acid intensifies
the color markedly. It has been claimed that this mercurial is as specific as
PM for SH groups, but one wonders if this complex molecule does not
N=N^ y-Hg
/)-Mercuriphenylazo-/:i-naphthol
through other groups occasionally react with various tissue components
(/5-naphthol derivatives being fairly potent enzyme inhibitors), although
previous treatment of the tissue with Hg++ or iodoacetamide is said to
prevent staining. Aqueous solutions of thiols, proteins, or tissue homogenates
are shaken with an amyl acetate solution of the mercurial dye, and a red
precipitate slowly forms in the aqueous phase as the reaction proceeds; the
amount of precipitate is proportional to the number of SH groups and can
be determined colorimetrically after centrifuging and redissolving in acid
solution. Fragments of dehydrated tissues may also be placed in butanol
or propanol solutions of the mercurial for several hours, and the staining
demonstrated histologically (Bennett, 1951). Bennett ran controls with phe-
nylazo-/5-naphthol to determine if this portion of the molecule contributed
to the binding, and generally found little or no staining. This mercurial
has been used to investigate thiol distribution in muscle (Bennett, 1948 b),
skin (Mescon and Flesch, 1952), and a variety of other tissues (Bennett,
1951).
Another colored mercurial, 4-mercuri-4'-dimethylaminoazobenzene, has
been used by Horowitz and Klotz (1956) to determine protein SH groups.
The solubility in water is so low that colorimetric determinations cannot
4-Mercuri-4'-dimethylaminoazobenzene
be made, but it dissoves sufficiently in 100 raM glycine (due to the forma-
tion of a glycinate complex) that reactions with SH groups in aqueous
medium can be carried out. However, it is also possible to determine the
amount of the mercurial removed from heptanol when shaken with an
aqueous solution containing the protein, although equilibrium usually re-
quires several hours. The specificity of reaction appears to be satisfactory,
768 7. MERCURIALS
since the amount bound to bovine seralbumin increases with the dye con-
centration until the molar ratio of dye to protein is 0.66, following which
no more is bound although the dye concentration is increased 50-fold. This
ratio corresponds quite closely to the known SH content of the protein,
lodination of the seralbumin prevents the reaction with the mercurial.
Ovalbumin reacts readily with two of its SH groups, slowly with a third,
and more slowly with the fourth, the dye perhaps differentiating the rel-
ative reactivities more closely than does p-MB. This method has a high
sensitivity and can be used for very low concentrations of protein.
A more recently examined mercurial dye, 4-(p-mercuriphenylazo)-l-naph-
thylamine-7-sulfonate, must also be dissolved in glyine buffer (Nosoh, 1961).
Absorption at 470 m// is determined and the titration of glutathione and
proteins appears to be quite satisfactory.
INHIBITION OF ENZYMES
The early concept of the mercurials as nonspecific denaturing and coagu-
lating agents for enzymes has gradually been abandoned in favor of a pic-
ture in which definite and often isolatable mercurial complexes are formed
under the proper experimental conditions. A selective reaction with SH
groups on enzymes is now generally assumed and the mercurials are exten-
sively used for the detection of these groups. The possibility of reaction with
other than SH groups has been discussed (pages 737 and 753) and should
never be ignored. We shall note instances in which a selective action on SH
groups is well established, and a few examples of inhibition not involving
SH group. We shall also see that mercurial inhibition does not necessarily
imply an SH group within the active center or the participation of an SH
group in the catalysis. In this connection, it is well to bear in mind the dif-
ferent groups which are introduced on the surfaces of enzymes when the
different mercurials are used (Fig. 7-10), inasmuch as the steric and elec-
trostatic effects of these side chains may be critical in producing inhibition.
Crystalline Mercuri-enzymes
The crystallization of the mercuric derivative of mercaptalbumin was
not the first instance of such a procedure. Warburg and Christian (1941,
1942) introduced this technique for the isolation of fermentation enzymes
and obtained the crystalline Hg-enolase complex from yeast, whereas the
normally active Mg-enolase could not be crystallized. Kubowitz and Ott
(1941) in Warburg's laboratory also crystallized the Hg++ complexes of
lactate dehydrogenases from Jensen sarcomata and rat muscle. The Hg++
complexes in all cases are enzymically inactive, but dialysis against cyanide
solution removes the Hg++ and restores the activity. There is no better
evidence for the homogeneous, stoichiometric, and reversible Hg++ deriva-
INHIBITION OF ENZYMES
769
tives of enzymes than such complexes, which is the reason they are discussed
briefly at this point. Warburg and Christian suggested that the isolation of
mercuri-enzymes might be generally useful, but this technique either was
not used or was unsuccessful until Kimmel and Smith (1954) reported the
crystallization of mercuri-papain. Krebs (1930) had shown that papain is
very sensitive to Hg++, 50% inhibition requiring only 0.005 milf , and this
-H,hQ-C
S-H,hQ
S-H,^^S
p-MPS
Fig. 7-10. The side chains introduced
onto proteins by various mercurials.
The S — Hg — R bonds are not actu-
ally linear but are shown in this way
for convenience.
indicated that a tight complex is formed and might be susceptible to crystal-
lization. Twice recrystallized papain (1.5-2%) was reacted with 1 mM Hg++
in 70% ethanol in the cold; within 24 hr a precipitate formed and in 3-4
days 90% of the activity was in crystalline form. These crystals are long
rectangular plates, often large enough to be visible to the eye, and are
soluble in water. The properties of mercuri-papain have been reviewed by
Kimmel and Smith (1957) and we shall discuss only those aspects relevant
to enzyme inhibition.
Mercuri-papain contains 0.49% Hg and has a minimal molecular weight
of 41,400; this corresponds to 1 Hg atom per molecule of mercuri-papain.
Since the molecular weight of reduced papain is around 20,500, mercuri-
papain must be a 1 : 2 complex or dimer to be represented by E — S — Hg —
770 7. MEKCURIALS
S — E. However, the situation is more complex, the pH being an important
factor in determining the type of complex occurring, and it is likely that
the crystalline mercuri-papain is the least soluble form of several possible
derivatives (Smith et al., 1954 b). Sedimentation studies at pH 4 indicate
a monomer or 1 : 1 complex, probably to be designated by HS — E — S — Hg+,
while at pH 8 there is a heavy component corresponding to a hexamer,
possibly cyclic with alternating — S — S — and — S — Hg — S — bonds. It is
interesting that there are two electrophoretic peaks at pH 4, one of unit +
charge greater than the other; since dissociation of the dimer must result
in equal proportions of HS — E — SH and HS — E — S — Hg+, this would tend
to confirm the dimeric structure. Oxidized papain is a mixture of
e(^\ and E— S— S— E
and does not react with Hg++; thus it is very important in studying the
combining ratios to be certain that the papain is fully reduced. Mercuri-
papain is actually purer than papain, as indicated by electrophoretic stud-
ies, has fewer N-terminal residues detected by the fluorodinitrobenzene
technique (Thompson, 1954), and has some 10% greater activity following
removal of the Hg++ with cysteine and EDTA, and it is also more stable.
The proteolytic enzyme, pinguinain, also forms stable complexes with Hg++
which are stable for much longer times than the pure enzyme (Messing,
1961). Other enzymes to be crystallized as the mercury complexes are a
lysozyme from papaya latex (Smith et al., 1955) and 3-phosphoglyceralde-
hyde dehydrogenase from yeast (Velick, 1953), the latter after reaction with
p-MB. There is some evidence that a mercuric dimer of ficin occurs (Liener,
1961) while carboxy peptidase forms very stable Hg++ complexes which still
possess esteratic activity, although they no longer function as peptidases
(Vallee et al., 1961; Coleman and Vallee, 1961). There is thus sufficient
evidence that many enzymes form well-characterized mercurial complexes
and are quite stable in this state; we shall note other examples in the dis-
cussion of SH titrations of enzymes.
These complexes of enzymes with Hg++ offer strong support to the con-
cept that completely selective reaction with SH groups can occur. How-
ever, if Hg++ is added in excess of that required for mercaptide formation,
it is quite possible that other enzyme groups may be attacked. It is likely
that other enzymes under the appropriate conditions can form dimers, or
other polymers, with Hg++, in which case the active centers may be made
inaccessible even though the SH group is not within the confines of the cen-
ter. The appearance of polymers will presumably depend strongly on the pH
since, at pH's progressively removed from the isoelectric point, one might
expect polymerization to be more and more reduced, due to the increasing
charge on the enzymes.
INHIBITION OF ENZYMES 771
Types of Inhibition Observed with the Mercurials
The concentration-inhibition curves for mercurials are generally sigmoid
and rather steep, as would be expected of inhibitors combining tightly with
enzyme groups. Indeed, when such curves are fairly flat, encompassing sev-
eral pi units, one has the right to question if the inhibition is related to
mercaptide formation, although it may well be. It should be emphasized
that adequate kinetic studies can be made only in preparations of pure
enzymes. The presence of impurities may distort the entire picture and the
kinetics of inhibition.
One may classify the inhibitions classically into competitive, noncompet-
itive, uncompetitive, and mixed types, but the proper plotting procedures
have seldom been used so that in the majority of cases we have little or no
information. However, sufficient has been done to show that all these types
of inhibition occur (Table 7-8). Competitive behavior has been observed
in a surprisingly large number of instances. This is surprising at first if one
assumes reaction with SH groups to be the primary mechanism of inhibi-
tion, because the tightness of the binding might be considered to prevent
the exhibition of competition. Actually, most inhibitions by mercurials are
probably competitive — either with substrate, coenzyme, or cofactor — in
the fundamental sense of the word, but it is often difficult to demonstrate
this by the usual analytical techniques which assume equilibrium conditions.
It is easier to show that the presence of the substrate, coenzyme, or cofactor
slows the development of the inhibition, although the equilibrium inhibition
may not be detectably different (page 778). Formally competitive behavior
might be expected to occur in the following circumstances. (1) The inhibitor
acts by a non-SH reaction; the organic mercurials particularly possess group-
ings capable of interacting with active sites independently of the Hg atom,
and such might be involved, for example, in the inhibition of D-amino acid
oxidase by p-MB, the benzoate structure being of primary importance. (2)
The binding of the mercurial to the SH groups may for some reason be
weaker than usual and of a comparable magnitude to the affinity for the
substrate. (3) The mercurial is bound much more tightly than the substrate
but measurements are made before equilibrium is reached, as in the exper-
iments showing protection of the enzyme by the substrate; when the inhi-
bitions are determined soon after adding the mercurial in the presence of
variable concentrations of the substrate, the data may provide formally
competitive plots. One would expect this third explanation in certain exam-
ples given in Table 7-8, e.g., carbonic anhydrase, where KJK,,^ = 3.87 X 10^'
for ??-MB (Chiba et al., 1954 b). In the case of homogentisate oxidase, p-MB
and MM inhibit competitively with respect to Fe++ but noncompetitively
with respect to homogentisate, the mercurials being bound roughly 40-100
times as tightly as the Fe++ (Flamm and Crandall, 1963). Here, and in
other instances where metal ion cof actors are involved, both cofactor and
772
7. MERCURIALS
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CH I— I
INHIBITION OF ENZYMES 773
mercurial are bound to the same SH group. In most cases, increase in the
substrate concentration does not reduce the inhibition once established, but
Robert et al. (1952) claim that acetylcholine is able to displace Hg++ from
horse serum cholinesterase, the affinity of the enzyme for the acetylcholine
actually being greater than for the Hg++; it is not known if mercaptide
formation is involved. One suspects that the inhibition may sometimes ap-
pear to be competitive where actually the mercurial is reacting with the
substrate, either exclusively or in addition to the enzyme (compare curves
in Figs. 1-5-1 and 1-5-14), and such might be the case in the inhibition of
/^-amylase by p-MB and PM (Ghosh, 1958), although the extent of reaction
of mercurials with starch is not known.
Noncompetitive inhibition may be observed when the mercurial reacts
with groups, SH or other, adjacent to the active center, and thus suppresses
the rate of breakdown of the ES complex, and when the affinity for the
enzyme is not so high that mutual depletion kinetics hold. The interference
with ES breakdown may be steric through the side chains introduced or
secondarily by alteration of the protein structure. It must be remembered
that mutual depletion systems usually indicate formally noncompetitive
behavior if the common plotting procedures are used (compare Figs. 1-5-3
and 1-5-24), despite the fact that the inhibition may be fundamentally
competitive. No pure instances of uncompetitive or coupling inhibition have
been reported, but it is not unlikely that preferred reaction with the ES
complex occurs. The inhibition of alkaline phosphatase is actually mixed
(noncompetitive and uncompetitive), but p-MB reacts more readily with
the ES complex {K,' = 0.163 mM) than with E {K, = 2.5 milf) (Lazdunski
and Ouellet, 1962). There are some examples in which such reaction with
the ES complex is possible, e.g., the inhibition of urease by Hg++ (Evert,
1952), of acid phosphatase by p-MB (Newmark and Wenger, 1960), and of
succinate dehydrogenase by p-MB (Warringa and Giuditta, 1958). The in-
hibition of lactate dehydrogenase from Propionibacterium pentosaceum by
p-MB is greater in the presence of lactate than when no substrate is present
during incubation, and this was postulated to be due to the greater number
of free SH groups, presumably arising through reduction by lactate (Moli-
nari and Lara, 1960), and a similar situation may occur with glutathione
reductase and NADPH, the presence of the reduced coenzyme increasing
the inhibition markedly (Mapson and Isherwood, 1963).
Another approach to the classification of mercurial inhibitions, and per-
haps the primary one, is the determination of the component — enzyme,
substrate, coenzyme, or cofactor — with which the mercurial reacts. Reac-
tion with the apoenzyme has generally been assumed above and with respect
to the molecular mechanism might be divided into three types: (1) binding
to an SH group at the active center, preventing complexing of the apo-
enzyme with any of the other components, (2) binding with an SH group
774 7. MERCURIALS
vicinal to the active center and interference with the catalysis sterically or
electrostatically, and (3) secondary altering of the protein structure to dis-
rupt the normal configuration of the active center. In the last case, which
is probably fairly common (see page 787), the inhibition may be formally
competitive (if the substrate stabilizes the enzyme structure), noncompeti-
tive, or quite complex. Reaction imth the substrate must often occur, espe-
cially when the substrate is protein, nucleic acid, nucleotide, or thiol, but
in most cases this possibility seems to have been ignored. It is obvious for
glutathione reductase and this complicates the analysis of the inhibition
(Mapson and Isherwood, 1963), but it may also be an important mechanism
when thioesters are involved, e.g., acetoacetyl-CoA in fatty acid synthesis
(Stern, 1956) or malonyl semialdehyde pantetheine in propionate metabol-
ism (Vagelos and Earl, 1959). The inhibition of NADPH: methemoglobin
oxidoreductase by p-MB occurs when either the enzyme or the methemo-
globin is incubated with the mercurial (Bide and Collier, 1964). Sometimes
one finds indirect evidence for reaction with the substrate, as with 5'-aden-
ylate deaminase (Lee, 1957). Here the inhibition by p-MPS is much greater
when it is preincubated with adenylate and the reaction started by adding
the enzyme than when preincubation is with the enzyme and reaction start-
ed by adding the substrate. Reaction with coenzymes is evident when lipoate
or coenzyme A is involved, but may be more general than is usually sup-
posed. A reaction of -p-MB with NAD was detected spectrophotometrically
by Palmer and Massey (1962) and this was considered to be significant in
titrations of certain dehydrogenases. Hill (1956) had previously established
a 1 : 1 complex of Hg++ with NADH, but had found no complex with p-MB.
Onrust et al. (1954) considered the possibility that at least part of the inhi-
bition of pyruvate oxidase by p-MB might be due to reaction with the sul-
fur of thiamine-diP, but excluded this when they found that thiamine-diP
does not reverse the inhibition. However, Pershin and Shcherbakova (1958)
observed that thiamine is able to reduce the bacteriostatic action of Hg++,
although this could be by a mechanism other than reaction of the thiamine
with Hg++. Kuratomi (1959), on the basis of preincubation experiments
with components of the pyruvate oxidase system, postulated that j'-MB
can react with thiamine-diP. This problem remains to be settled and possibly
is an important one. It would be interesting to know if mercurials can open
the thiazole ring under physiological conditions (which is not likely) or
react with the SH groups after ring opening, in which case the state of the
thiamine-diP in the preparation would be important. Another possibility
is that a complex is formed with groups other than the sulfur since oppor-
tunities for chelation exist.
It may be suggested that in all studies of enzyme inihibition, in which
substrates or coenzymes capable of reacting with mercurials are involved,
the appropriate preincubations with the inhibitor be carried out, as pre-
INHIBITION OF ENZYMES 775
viously described (page 1-569), since this technique will often provide in-
formation on complexes formed with components other than the apoenzyme.
Whatever the mechanism or formal type of inhibition by mercurials, it
is certain that many systems must be represented by mutual depletion
kinetics. This is clearly seen in many of the enzyme titrations (page 804),
inhibition being produced by mercurials at roughly equimolar concentra-
tions relative to the enzymes, but at this point the problem will be treated
in a more general manner. Mutual depletion behavior implies that the inhi-
bition will depend on the concentration of the enzyme. This is seen with
yeast pyruvate decarboxylase in the work of Stoppani et al. (1953) (see
accompanying tabulation), and even more markedly with pig heart suc-
Pyruvate % Inhibition by:
aecarooxyiase
(/'g/ml)
Hg++
p-MB
7.8
85.0
95.0
15.7
—
75.0
30.5
43.0
33.0
01.0
15.9
7.0
cinate oxidase, which is inhibited 89% by 0.01 mM p-MB when the enzyme
concentration is 0.15 mg/ml but only 59% by 0.76 mM p-MB when the
enzyme concentration is 30 mg/ml (Stoppani and Brignone, 1957). Another
example is muscle p>Tuvate oxidase (see accompanying tabulation) (Onrust
Enzyme extract „^ inhibition by p-MB 0.11 mM
(ml)
0.4 82
0.8 56
1.2 34
1.5 33
et al., 1954). These few examples well illustrate the importance of this factor
and very clearly demonstrate the quantitative meaninglessness of most
reported inhibitions if the relative enzyme concentration is not known or
stated. Impurities also may contribute to the depletion of the mercurial.
The crude bacterial enzyme for converting histidinol to histidine is not
inhibited by 0.02 mM p-MB, but the partially purified enzyme is inhibited
50% (Adams, 1954), and it is likely that the pure enzyme would be inhib-
776 7. MERCURIALS
ited even more strongly; such work points out the importance of enzyme
purity for accurate studies of mercurial inhibition. The elevation of the
plgo from 0.0002 mM to 0.014 uiM by serum for the inhibition of 3-phos-
phoglyceraldehyde dehydrogenase by p-MB is a further example (Weitzel
and Schaeg, 1959).
When an enzyme is reported to be inhibited to a specified degree, say
50%, by a certain concentration of mercurial, exactly how is this to be
interpreted? Is 50% of the enzyme combined with the mercurial in a com-
pletely inactive EI complex, or is all the enzyme combined with the mer-
curial and the EI complex possesses 50% of the original activity? If the
ordinary equilibrium formulation is followed and it is assumed that the
fractional activity of the EI complex is r, noncompetitive inhibition will
be given by
.^ (l-r)(I)
(I) + Ki
and
(7-4)
1 Ki
+ -r. -^;vr (7-5)
i (1-r) (l-r)(I)
so that a plot of \ji against 1/(1) will give a straight line intersecting the
1/i axis at 1/(1 — r), or \jimax- If mutual depletion occurs (zone C), a sim-
ilar result is obtained, although the slope will be different. A simple plot
of this type may help to decide between the two possibilities above. If the
plot is not linear near the \ji axis, one might suspect that another type of
inhibition is occurring at higher inhibitor concentrations, or that secondary
inactivation of the enzyme is a factor.
One example of the deviations from classic inhibition kinetics that may
be seen with the mercurials is the inhibition of human plasma cholinesterase
by Hg++ as analyzed by Goldstein and Doherty (1951 ). This slowly develop-
ing, pH- and temperature-dependent inhibition presents some interesting
but often uninterpretable results. The l/v-l/(S) plots exhibit two sorts of
deviation (Fig. 7-11). The results from long incubation with low concentra-
tions of Hg++ fall on reasonably straight lines (A and B), but the slopes are
a good deal greater than expected for pure noncompetitive inhibition, as
for mixed inhibition (Fig. I-5-6A) the interaction constant a being some
finite value > 1. Of course, it may not actually be true mixed inhibition,
the deviation being due to some other factor. The results from short in-
cubations with high concentrations of Hg++ differ so much from any sort
of classic behavior that it is impossible to interpret them (C and D). It was
suggested that low and high concentrations of Hg++ inhibit by different
mechanisms, possibly with different SH groups, the former with groups
outside the active center causing secondary irreversible inactivation and
the latter directly with groups in the active center. This would to some
INHIBITION OF ENZYMES
777
extent explain why the inhibition is more competitive at high Hg++ and
more noncompetitive at low Hg++, but it does not explain the deviations
discussed above. Curves C and D presumably do not represent equilibrium
inhibitions and are more illustrative of protection of the enzyme by the
substrate; it would seem that acetylcholine above 50 mM protects the en-
zyme almost completely against very short exposures to Hg++, which is
not too unreasonable considering the relatively high affinity of the enzyme
for acetylcholine. Although the kinetics of protection and the application
2000-
1500-
1000-
500
.
A
0 0126 mM FOR 4 5 HOURS
/
B
O0Z73mM FOR 4 HOURS
/
C
2 28 mM FOR 20 MIN
^X /
0
4 S4 mM FOR 3 MIN
/ /
/c
^/ / _J— —
CONTROL
20
60
Fig. 7-11. Double reciprocal plots for the inhi-
bition of human plasma cholinesterase by Hg++,
showing deviations from linearity at high Hg++
concentrations. (Modified from Goldstein and
Doherty, 1951.)
to plotting procedures have never been worked out as far as I know — and
it would be difficult to treat the phenomenon rigorously one might predict
that curves with rather steep slopes in the l/v-l/(S) plot would be found, and
that such curves would occasionally intersect the control curve to the right
of the \\v axis, i.e., the longer the incubation, the closer to equilibrium
would the inhibition come, and the less competition or protection would
be exerted by the substrate. One also wonders if the increased tilt of curves
A and B might be due to the fact that these relatively high substrate con-
centrations protect the active center against structural changes brought
about by reaction of the Hg++ at vicinal sites, since there are many examples
778 7. MERCURIALS
in which the substrate can slow down spontaneous or induced enzyme de-
naturation. However, neostigmine, which can protect chohnesterase against
thermal denaturation, does not protect at all against Hg++. There is ac-
tually some doubt as to whether the inhibition is related to SH groups,
since p-MB and MM up to 1 raM do not inhibit even after 2 hr at 37^, or it
might mean that the reacting SH groups are not at the active center and
the inhibition by Hg++ is due to a dimerization or polymerization. If all
enzymes subjected to mercurials were studied in as much detail as in this
work, there would probably be many more interesting examples of devia-
tions from classic theory; as long as one tests an enzyme under standard
conditions with one concentration of a mercurial, as is done in most reports,
interpretation presents no problems.
Protection of Enzymes against Mercurials
Enzymes may be protected against mercurials by (1) substrates, (2) co-
enzymes, (3) metal ion cofactors, (4) reversible inhibitors, and (5) thiols or
other mercurial complexers. Various conclusions have been drawn from
such experiments, mainly regarding the relation of SH groups to the bind-
ing of the protector, but there are many pitfalls; the discussion of protection
with respect to iodoacetate (page 47 and Fig. 1-5 in Volume III) applies
equally well to the mercurials. Protection may occur by two general mech-
anisms: reaction of the protector with the enzyme to block off the mer-
curial, or reaction of the protect&r with the mercurial. The latter mechanism
applies to the thiols such as cysteine or glutathione, which have been widely
used for this purpose, but, as has been pointed out several times, such pro-
tection does not provide much useful information, since in reality all one
does is to reduce the effective mercurial concentration. It also applies to
other complexers and perhaps is involved in the following: the protection
of fumarate hydratase (Mello Ayres and Lara, 1962) and fumarase (Fave-
lukes and Stoppani, 1958) by phosphate, of ascorbate oxidase by amino
acids and RNA (Frieden and Maggiolo, 1957), of acid phosphatase by EDTA
(Macdonald, 1961), of urease by ascorbate (Mapson, 1946), and of thyroxine
delahogenase by FMN (Tata, 1960). However, in these cases it is often
difficult to interpret the mechanism of the protection. We shall not be con-
cerned with this type of protection, but only with those protectors presum-
ably reacting with the enzyme.
Some examples of protection are summarized in Table 7-9 along with
instances in which protection does not occur (or at least is not observed
under the conditions used). The + sign does not indicate that complete
protection can be achieved; indeed, in most cases only partial protection
has been reported, and this is what we would expect. The degree of pro-
tection may depend on the concentrations of mercurial and protector; e.g.,
protection may be complete with low mercurial concentrations, whereas
INHIBITION OF ENZYMES 779
the protector may be relatively ineffective against high concentrations,
as in the effects of arginine on the inhibition of its oxidative decarboxylation
(see accompanying tabulation) (Van Thoai and Olomucki, 1962). In most
% Inhibition
p-MB
(mM)
0.033
57
0.05
66
0.067
89
0.083
98
p-MB alone p-M.B + arginine 10 mM
0
0
43
61
reports it is difficult to decide if the protection is simply due to a slowing
of the rate of inhibition or to a true effect on the final equilibrium inhibi-
tion, since measurements are often made over arbitrary time intervals. It
is evident that it is easier to slow down an inhibition than to modify its
final level; enough substrate, coenzyme, or cofactor to saturate the enzyme
substantially will quite markedly slow the reaction of the enzyme with
the inhibitor, but the final inhibition need not be significantly changed,
particularly with the mercurials which are usually bound tightly, if slowly.
Most investigators have noted that although the protectors in Table 7-9
are effective when present during the development of the inhibition, they
do not reverse the inhibition at all once it has reached a steady level, this
apparently indicating that most of the protection results are fundamentally
due to a slowing of the rate of inhibition.
Occasionally two components of the enzyme reaction, forming a ternary
complex with the enzyme, protect more than each component alone. This
is the situation with malate oxidative decarboxylase, the protections by
malate and Mn++, or malate and NADP, being additive; the protections
by Mn++ and NADP are not (Rutter and Lardy, 1958). It may also be the
case with liver alcohol dehalogenase, ethanol and NAD protecting more
than either one alone (Yonetani and Theorell, 1962). In one situation, aspar-
tate carbamyltransf erase, neither substrate alone protects, but together they
do so quite effectively (Reichard and Hanshoff, 1956). An example of pro-
tection by a reversible inhibitor is the reduction in the inhibition of succinate
dehydrogenase by p-MB or Hg++ in the presence of oxalacetate (Stoppani
and Brignone, 1957). Actually, an effective competitive inhibitor might be
expected to protect better than the substrate.
The information derived from protection experiments is frequently not
as reliable as commonly assumed, for reasons to be discussed in Chapter 1,
Volume III. The fact that the action of a mercurial is reduced by a sub-
780
7. MERCURIALS
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INHIBITION OF ENZYMES
781
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7. MERCURIALS
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INHIBITION OF ENZYMES 783
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784 7. MERCURIALS
strate, for example, does not necessarily imply that the substrate reacts
with an SH group nor that the SH group is involved in the catalysis, al-
though these may well be the case. A positive result is more valuable than
a negative one. The failure to achieve protection may be due to an inade-
quate concentration of the protector, too low a relative affinity of the en-
zyme for the protector, or a long incubation wherein equilibrium is reached,
and yet the substance examined may participate in the reaction and inter-
act with the enzyme in the same way as effective protectors. Definite pro-
tection allows one to make the reasonable assumption that the mercurial
binds somewhere in the region occupied by the protector. Potter and Du-
Bois (1943) postulated an SH group to be located between the two cationic
groups binding succinate to the dehydrogenase; protection against mercuri-
als by succinate simply implies that succinate is able to shield this SH
group from the mercurial, and not that the SH group is involved in the
succinate binding or participates in the oxidation-reduction reaction. For
steric reasons, the smaller the molecular sizes of the protector and the mer-
curial, the more certain can one be that a common SH group is involved
in the binding of both.
Displacement of Coenzymes and Cofactors from Enzymes
Closely allied to protection experiments are those in which a mercurial
is shown to dissociate an enzyme-coenzyme or enzyme-cofactor complex.
It is now believed that several coenzymes and metal ion activators may be
bound to apoenzymes through SH groups in part (Shifrin and Kaplan,
1960), and if this is so one would expect tightly bound SH reagents, such
as the mercurials, to displace the coenzymes or activators. Certain coen-
zymes or cofactors, such as NAD, have been shown to react with thiols
(van Eys and Kaplan, 1957 b), but in most cases the evidence for binding
to enzyme SH groups is circumstantial. Certainly such displacement of
necessary components of the enzyme reaction would be an important
mechanism in the inhibition produced by mercurials, especially in vivo where
the total binary or ternary complexes usually occur. One molecule of crys-
talline horse liver alcohol dehydrogenase binds 2 molecules of NADH at
physiological pH and this is accompanied by a shift in the absorption
spectrum of the NADH. Addition of p-MB was found by Theorell and
Bonnichsen (1951), to reverse the spectral shift, and it was concluded
that the bond between an enzyme SH group and the NADH pyridine
ring is broken by the mercurial. However, some doubts have recently been
cast on this simple interpretation. The liver alcohol dehydrogenase mol-
ecule has 28 SH groups as determined by p-MB titration; the presence
of NADH does not reduce this number, although NADH protects the en-
zyme moderately (Witter, 1960). On a rather tenuous basis, Witter postulat-
ed that the function of the SH groups is to maintain the stable enzyme
INHIBITION OF ENZYMES 785
structure rather than bind the NADH; disintegration of the structure
brought about by p-MB would secondarily lead to release of the coenzyme.
As the NADH is split from the apoenzyme by p-MB, rotatory dispersion
titration indicates changes in optical rotation associated with denaturation,
so that Li et al. (1962) likewise inclined to a theory involving structural
changes as a basis for the displacement, since it is known that denaturation
by heat or other agents releases the NADH. Yonetani and Theorell (1962)
have used the very sensitive spectrofluorometric method for the measure-
ment of NADH binding and dissociation.* They demonstrated that the en-
zyme configuration is stabilized by the NADH and additionally by the iso-
butyramide, although these can be associated directly with only a small
fraction of the total number of SH groups, and suggested that denaturation
may be Initiated by local changes at the active centers and from there
spread throughout the molecule. The SH groups may form a network of
hydrogen bonds contributing to the stability of the tertiary structure, so
that mercurials could create instability either locally or generally. All of
this recent work shows that mercurials probably induce configurational
changes in the enzyme, these being irreversible by the usual means, and
they provide an alternative explanation for NADH release, but do not dis-
prove the original hypothesis that a direct binding between NADH and
SH groups occurs. The Zn++-dependent alcohol dehydrogenase of yeast is
inhibited by mercurials, and this was attributed to displacement of the
Zn++ from SH groups (Wallenfels and Sund, 1957 a) on the basis that res-
toration of activity requires both glutathione and Zn++. However, an inves-
tigation of the time course of the inhibition showed that glutathione alone
is sufficient to reactivate if it is added soon after the mercurial, but the inhi-
bition progressively becomes irreversible, at which time no Zn++ has been
released (Snodgrass and Hoch, 1959). Zn++ is displaced progressively over
a period of several hours, but the inhibition does not appear to be mediat-
ed through this displacement. Such a slow release, without correlation with
the mercurial reaction or the inhibition, is probably due to structural
changes in the enzyme.
The early work on NADH splitting from alcohol dehydrogenase was soon
confirmed for 3-phosphoglyceraldehyde dehydrogenase (3-PGDH) by Velick
(1953). The yeast enzyme requires 2 SH groups for full activity and inhi-
bition by p-MPS increases until 2 equivalents of the mercurial are added.
In contrast to the alcohol dehydrogenase, this inhibition is readily revers-
* Liver alcohol dehydrogenase forms a very stable ternary complex with NADH
and t«obutyramide, and this complex is strongly fluorescent. The enzyme may be
titrated in the presence of 100 mM isobutyramide with NADH, measuring the fluo-
rescence increase at 410 m/<, and then back-titrated with p-MB or p-MPS as the NADH
is released from the apoenzyme. This will probably be a very valuable technique for
the study of coenzyme binding.
786
7. MERCURIALS
ible with cysteine. Muscle 3-PGDH behaves similarly but, after charcoal
treatment, reacts with 3 equivalents of p-MPS. If the enzyme is ultracen-
trifuged it carries down most of the NAD but the inhibited enzyme does
not, most of the NAD being free in the medium. The correlations between
these various events are shown in Fig. 7-12. The change in spectral absorp-
tion at 340 ji/m associated with NAD binding disappears progressively with
added p-MPS, again indicating coenzyme release. Velick was careful not to
assume that these results necessarily point to a binding of the NAD by
the SH groups, but stated that the p-MPS can sterically or electrostatically
1.5
p-MPS BOUND
0.5 ,
"0 I 2
EOUIVS p-MPS ADDED
Fig. 7-12. Dissociation of NAD from 3-phosphogly-
ceraldehyde dehydrogenase by . p-MPS, and the si-
multaneous loss of activity. (From Velick, 1953.)
interfere with the coenzyme binding if the SH groups are close enough to
the active center. Fluorometric titration of the 3-PGDH- (NADH)3 complex
from rabbit muscle with p-MB demonstrates NADH dissociation, but the
kinetics indicate that each p-MB bound weakens the coenzyme binding at
the other sites, so that perhaps structural changes in the protein occur
(Velick, 1958). The same number of equivalents of p-MB is required to
release the NADH from 3-PGDH- (NADH )i, 3-PGDH- (NADH)2, and 3-
PGDH-(NADH)3. If the displacement were due to direct competition with
the NADH for SH groups, one would expect the p-MB to attack the unoc-
cupied SH groups first, with no release of NADH, but this is not the case,
the release beginning immediately, as if the reaction of any SH group al-
tered the enzyme structure throughout. These structural changes, if they
occur, must be readily reversible.
Results with a few other enzymes will be discussed briefly. Reports on
coenzyme displacement from lactate dehydrogenase have not been entirely
consistent. Apparently the state of the enzyme and particularly the source
are important factors. Kaplan and Ciotti (1954) found that p-MB releases
NAD from the liver enzyme, this being associated with a fall in absorption
at 300 m//, but Chance (1954) could detect no release from heart lactate
INHIBITION OF ENZYMES 787
dehydrogenase at 5°, an observation confirmed by Velick (1958). However,
Winer et al. (1959) find a slow dissociation of NADH from heart lactate
dehydrogenase, complete release occurring after 1 hr at 26° and pH 7 with
0.135 vciM 2?-MB. The L(+)-lactate dehydrogenase of yeast (cytochrome
bj) possesses a flavin prosthetic group and this is readily dissociated by
p-MPS (Armstrong et al, 1960, 1963). The binding of NADH and NADPH
to cytochrome 65 aporeductase is blocked by p-MB (Strittmatter, 1961 b)
and there is some evidence that pyridoxal-P may be split from L-threonine
deliydrase by the same mercurial (Nishimura and Greenberg, 1961). The
evidence for the displacement of Fe++ from homogentisate oxidase by p-MB
has already been discussed, and the reactions of inhibition and reactivation
(Crandall, 1955) may be written:
Inhibition: E— S— Fe+ + R— Hg— X -> E— S— Hg— R + Fe++ + X-
E— S— Hg— R + GSH -> E— SH + GS— Hg— R
Reactivation: „ „ „
E— SH + Fe++ -> E— S— Fe+ + H+
The nonheme Fe of succinate dehydrogenase is lost more rapidly by dialysis
after treatment with p-MPS, and this may be related to the marked spectral
changes observed upon reaction with the mercurial (Massey, 1958). The
iron of the photosynthetic pyridine nucleotide reductase is released as Fe+++
by p-MB with proportional loss of activity (Katoh and Takamiya, 1963).
The inhibition of aminopeptidase by EDTA is made irreversible by simul-
taneous treatment with p-MB and it was concluded that the Mn++ is bound
to an SH group (Bryce and Rabin, 1964). A final type of experiment will
be mentioned. Mn++ activates the hydroxylamine reductase of P. aeruginosa
and this activation is prevented by p-MB, "suggesting that SH groups may
be involved in binding the metal to the enzyme" (Walker and Nicholas,
1961). It seems to me that such conclusions are unjustified, inasmuch as
any mechanism of inhibition would presumably abolish activation by Mn++,
whether it affected the binding or not.
The results on coenzyme displacement may be summarized by stating
that the same difficulties are encountered as in protection experiments.
There are three general mechanisms by which a mercurial could dissociate
an enzyme-coenzyme complex: (1) compete with the coenzyme for the SH
group, (2) sterically or electrostatically interfere with coenzyme binding by
reacting at an adjacent site, and (3) alter the enzyme configuration in such
a way as to disrupt secondarily the coenzyme binding. In no case have these
mechanisms been distinguished.
Changes in Enzyme Structure Brought About by Mercurials
Evidence has accumulated during the past several years that mercurials
occasionally initiate configurational changes in enzymes; certain aspects of
788 7. MERCURIALS
this have been discussed in the previous section and we shall now inquire
what further evidence on this important problem has come to light. One
of the more obvious reasons for suspecting denaturation is the progressive
development of irreversibility during contact with the mercurial, such as
has been reported for cholinesterase (Goldstein and Doherty, 1951), pros-
tatic phosphomonoesterase (Tsuboi and Hudson, 1955 a), muscle aldolase
(Swenson and Boyer, 1957). and muscle 3-PGDH (Elodi, 1960) — to men-
tion only a few instances, in most cases reversal being attempted with glu-
tathione or dimercaprol. Of course, one might attribute failure to reverse
to very tight binding to the enzyme, but the progressive increase in the
irreversibility points more to structural changes. The question often remains
as to whether these changes are responsible for the inhibition or are super-
imposed upon it, i.e., inhibition followed by inactivation.
Elodi (1960) investigated the changes in several properties of pig muscle
3-PGDH treated with p-MB, and found significant deviations in the op-
tical rotation and the intrinsic viscosity, the latter increasing linearly with
the equivalents of mercurial added. The following phases were postulated:
(1) an initial reversible binding and inhibition, (2) a progressive disintegra-
tion of the secondary structure of the enzyme as a result of the blocking of
SH groups, this probably involving an unfolding of the polypeptide helices,
and (3) polymerization and precipitation consequent to the freeing of
groups which form intermolecular bridges. The simultaneous changes in the
activity, NADH binding, and rotatory dispersion of yeast alcohol dehydro-
genase treated with p-MB led Wallenfels and Miiller-Hill (1964) to postulate
that modifications of the secondary and tertiary protein structure occur
when the SH groups are blocked. Reaction of 10 SH groups on muscle
aldolase with p-MB does not reduce the activity but the susceptibility to
tryptic digestion is increased (Szabolcsi and Biszku, 1961). Untreated al-
dolase or treated enzyme in the presence of fructose-diP is not digested by
trypsin; thus the substrate apparently protects the active center, and per-
haps the entire molecule, from hydrolysis. It is thought that reaction of the
first 7 free SH groups labilizes the tertiary structure of the enzyme, and
from then on a progressive denaturation occurs. Addition of substrate may
restore to some degree the normal structure. The inhibition that occurs
later or with excess p-MB does not seem to be directly related to mercaptide
formation but dependent on the structural changes when they have pro-
ceeded past a certain point. Another interesting approach was made by
Massey (1958) in showing that the chelation of the nonheme iron of suc-
cinate dehydrogenase by o-phenanthroline is accelerated by treatment with
p-MPS, this being interpreted as a structural change exposing the iron.
The SH groups of yeast hexokinase can be titrated with p-MB in the pre-
sence of glucose without loss of activity, but spontaneous denaturation
quickly follows (Fasella and Hammes, 1963). Glucose-6-P does not prevent
INHIBITION OF ENZYMES 789
loss of activity during the titration. These results indicate that the SH
groups are not directly involved in the catalysis, but function to stabilize
the enzyme in the active configuration.
Another type of structural change is depolymerization of the enzynae
into subunits following mercurialization. Muscle phosphorylase is progres-
sively inhibited by p-MB until around 18 equivalents of the mercurial are
combined, and this is accompanied by the appearance of a new molecular
species in the ultracentrifuge, the sedimentation constant being lower than
that for either phosphorylase a or b (Madsen and Cori, 1955):
Phosphorylase a: S = 13.2
Phosphorylase b: S = 8.2
Inactive enzyme: S = 5.6
Both phosphorylase a and b form this new species with p-MB and it was
suggested that the former is split into 4 subunits, the latter into 2 subunits.
Light scattering studies are consistent with this interpretation (Madsen,
1956). The inhibition develops more rapidly than the depolymerization,
however, so the relationship between them is not clear. Removal of the
p-MB with cysteine restores both activity and the normal dimer or tetra-
mer (Madsen and Cori, 1956). The extent of the conversion of the phos-
phorylase tetramer to the monomer is proportional to the number of SH
groups reacted and an all-or-none dissociation of the units is likely (Madsen
and Gurd, 1956). The sedimentation constant of yeast alcohol dehydroge-
nase is reduced from 7.2 to 3.3 by p-MB, this being secondary to the inhi-
bition of the enzyme, so that here dissociation into subunits apparently
occurs (Snodgrass et al., 1960). Reaction of myosin ATPase with MM also
causes the appearance of a small subunit, but this is not related to the
binding to the SH groups responsible for the activity (Kominz, 1961). In
addition there is some aggregation to a faster sedimenting species and this
is perhaps correlated with reaction of SH groups at the active center. The
inhibition of rabbit muscle enolase by p-MB was considered to be secondary
to denaturation and not directly due to SH group reaction, on the basis of
the variation of activity with the equivalents of mercurial present and the
appearance of turbidity (Malmstrom, 1962). The sedimentation constant of
liver glutamate dehydrogenase is reduced by MM and PM and again the
most likely explanation is a splitting into subunits (Rogers et al., 1962,
1963; Greville and Mildvan, 1962). The relationship of the disaggregation
to the unique changes in enzyme activity is not clear.
There is no doubt that mercurials can induce structural changes in cer-
tain enzymes, and cause aggregation or fractionation into subunits in others,
but the significance for primary inhibition has not been clarified. Is the
inhibition due to the blocking of functional SH groups or secondarily to
the structural changes? Does the denaturation result from general SH group
790 7. MERCURIALS
reaction or can it originate solely by mercaptide formation at the active
center? The results taken all in all tend to signify that inhibition usually
occurs upon the initial reaction of the SH groups at or near the active
center, this being reversible, and that slower structural alterations proceed
as a result of either the mercurial already combined or the continued reac-
tion with more mercurial (perhaps with the less available SH groups), these
changes becoming more and more irreversible, a progressive inactivation
being superimposed on the primary inhibition. Lability may also come
about by a displacement of coenzyme or cofactors, since these undoubtedly
help maintain configurational integrity, especially in the abnormal state in
which isolated enzymes find themselves. This does not imply that all en-
zymes behave in this fashion; it is quite possible that in some the structural
changes may be primary and the sole cause of the inhibition. Certain en-
zymes suffer only the primary inhibition and the stability is not reduced
by the mercurial, and indeed stability may be increased, as we have ob-
served with papain and pinguinain. The requirement to solve the problems
of the relation between inhibition and inactivation, and between both proc-
esses and the types of SH group reacted, is for more detailed studies cor-
relating the time courses of as many of these changes as possible as they
occur after introduction of the mercurial. Another approach might be to
do occasional experiments at low temperatures, where inactivation or de-
naturation would occur very slowly, in this way possibly separating the
primary inhibition from these other changes. Finally, it might be suggested
that every effort to create conditions favoring stability of the enzymes be
made. One gets the impression that often so little attention is paid to the
proper pH, ion concentrations, buffers, and other factors, that the enzyme
as studied is in a relatively unstable state and hypersusceptible to any in-
hibitors subjecting the normal protein configuration to even minor stress.
Effects of pH, Ions, and Buffers on Mercurial Inhibition
The effects of pH on OH"" complexes with mercurials (page 736), on
mercaptide formation (page 749), and on reactions of proteins with mer-
curials (page 760) have been discussed. The results and the factors which
may be involved can be summarized as follows: (1) pH affects the ioniza-
tion of the SH groups or the competition between mercurial and H+ for
the S~ group, (2) pH alters the concentration of 0H~ and hence the amount
of mercurial complexed with this anion, (3) pH influences the protein
charge possibly attracting or repelling charged mercurials, (4) pH deter-
mines the rate of secondary inactivation or denaturation, (5) pH affects
the aggregation state of protein-mercurial complexes (e.g., the degree of
dimerization of mercaptalbumin complexes with Hg++), (6) pH affects both
the rate and the number of SH groups reacted, and (7) generally there is
an increased rate of protein reaction with mercurials as the pH is reduced.
INHIBITION OF ENZYMES
791
The effects of pH on enzyme inhibition by the mercurials are even more
complex and one would not anticipate consistent behavior, a prediction
that is borne out in the following discussion.
Some pH effects on mercurial inhibition are shown in Table 7-10. In 7
cases the inhibition is greatest at low pH and in 6 cases at high pH, and
this certainly indicates that more than one factor must be involved. Indeed,
many of the inhibition-pH curves are complex (Figs. 7-13 and 7-14) and
80
60
20
Fig. 7-13. Effects of pH on the inhibitions of /J-fruc-
tofuranosidase by HgClj and Hg(N03)2. (Data from
Myrback, 1926.)
often biphasic (/^-glucuronidase, /J-fructofuranosidase, ATPase, and ribo-
nuclease). In some cases marked stimulation is found within a certain pH
range, inhibition occurring outside this range. This implies that the activity-
pH curves and the pH^p^ are shifted by the mercurials, usually to lower
pH's, as for yeast proteinase (Lenney, 1956) and ascites cell ribonuclease
(EUem and Colter, 1961; Colter et al., 1961). It is difficult to interpret such
shifts in pH^p^, but if the pH^p^ is related to the ionization of two or more
groups on enzyme and substrate (see page 1-660), a shift implies some mod-
ification of the enzyme groups in or near the active center with a resultant
alteration of the interaction of the substrate with the enzyme. Dixon's
method of plotting K,„ and K^ against pH (page 1-683) was applied to ^-
glucuronidase by Fernley (1962), and the curves are shown in Fig. 7-15.
The pK^ curve is suggestive of two ionizing groups on the enzyme with
pi^L^'s around 4.4 and 6.3 if it could be simply interpreted, or possibly the
792
7. MERCURIALS
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INHIBITION OF ENZYMES 793
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794
7. MERCURIALS
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INHIBITION OF ENZYMES
795
group with p^a near 4.4 could be the benzoate group of p-MB, particularly
as no inflection is shown here in the K„, curve. However, as Fernley clearly
pointed out, one must proceed with caution because of the many complexes
possible with the mercurial, and particularly the buffer effects. It seems
unlikely that the decrease in K, above pH 5 is due to complexing with OR-
ions entirely, but it might be the result of reactions with the buffer; the sim-
ilar inflection for K„^ makes it reasonable that an enzyme group is involved.
The results with PM are quite different than with Hg++ or p-MB (Fig. 7-14);
100
80
60
40
20
-20
Hg=0.05 mM
Ag^O 1 mM
Hg**
Cui= 1 mM
PM<0 4 mM
/ /*'*
K
^ PM
r"
■
y^
Fig. 7-14. Effects of pH on the inhibitions of ^-glu-
curonidase by heavy metal ions. (From Fernley, 1962.)
PM is generally less inhibitory than p-MB, does not increase Kf„ whereas
p-MB does, facilitates the formation of the ESg complex (favoring substrate
inhibition) whereas p-MB does not, and increases the pH^pt whereas Hg++
and p-MB decrease it. The marked effect of PM on substrate inhibition is
shown in Fig. 7-16 but no explanation is available. There is certainly a need
for more accurate comparisons of the different mercurials, not only with
respect to pH effects but generally. The different curves obtained with Hg++
and PM acting on pancreatic amylase (Fig. 7-17) are intriguing and one
feels that an explanation of such phenomena might well aid in our under-
796
7. MERCURIALS
3-
■
■— -~^'
^^^
•-^iljm
'
^~~^^.,^WG V^ N.
Fig. 7-15. Effects of pH on the ^„ and V„ for ^-glucuronidase,
and the K, for the inhibition by p-MB. (From Fernley, 1962.)
standing of the mechanisms of enzyme inhibition. The pH changes may af-
fect the reactivities of different SH groups on an enzyme to different de-
grees. Xanthine oxidase has two types of SH group, one reacting relatively
100
80
40
20
CU (0 63mM|
PM(025mM)
Fig. 7-16. Effects of substrate concentration
on the inhibition of /3-glucuronidase by
p-MB, PM, and Cu++ at pH 5.9. (From
Fernley, 1962.)
INHIBITION OF ENZYMES
797
rapidly and unrelated to the enzyme activity and the other reacting slowly
to produce inactivation; altering the pH from 5.2 to 7.2 modifies the rates
of reaction of these groups with p-MB in quite different ways (Gilbert,
1963).
Increase of any ligand capable of complexing the mercurials should re-
duce the inhibition, but this has been studied very little. Fernley (1962)
noted that raising the CI" concentration suppresses the inhibition of /?-
glucuronidase by Hg++. It is very difficult in such cases to separate a direct
complexing action from an ionic strength effect. Green and Neurath (1953)
-6 -5 -4
LOG CONCENTRATION (mM)
Fig. 7-17. Concentration-inhibition curves for the actions of Hg++
and PM on pancreatic amylase. (From Owens, 1953 b.)
varied the ionic strength with NaCl, SrClg, and (NH4)2S04 and found the
inhibition of trypsin by Hg++ to be suppressed at high ionic strengths
(Fig. 7-18). Since all the salts had essentially the same effect, they assumed
that this is not due to specific ions, but certainly part of the reduction
must be due to increasing formation of the Hg++ complexes with CI", NH4+,
and 864=. The nature of the inhibition of trypsin is not clear since free SH
groups are generally not considered to play a role in the active center.
The inhibition of pancreatic amylase by Hg++ and PM was postulated by
Owens (1953 a) to involve phosphate ion, and the unusual configuration of
the PM inhibition curve was attributed, at least in part, to the formation
of phosphate complexes, although it is strange that Hg++ is not similarly
affected. It is also not clear to me why the curve should assume this shape,
particularly why increase in PM from 0.1 to 1 vaM should bring about a
798
7. MERCURIALS
lessening of the inhibition. The inhibition of NADH dehydrogenase by p-
MPS is weaker in phosphate than in THAM buffer, and this could again be
due to complexes with phosphate (Minakami et al., 1963). Despite the pau-
city of experimental information, the effects of ions and buffers should be
more frequently taken into account in the use of the mercurials. In addition
to the ionic effects just described, certain specific actions have been noted,
100
50
25
X
ACT
rONIC STRENGTH ON
HgClj INHIBITION
(HgClj).0 05mM
'0 0.01
IONIC STRENGTH
0.02
0.03
0.04
005
Fig. 7-18. Inhibition of trypsin by Hg++ and the ef-
fect of ionic strength. (From Green and Neurath, 1953.)
especially with ATPase (Novikoff et al., 1952; Lardy and Wellman, 1953;
Sacktor et al, 1953), where the concentrations of Mg++ and Ca++ determine
to some extent the susceptibility of the mitochondrial enzyme to mercurials,
generally Mg++ lessening the inhibition, although here one may be dealing
with more than one enzyme.
Titration of Enzyme SH Groups
The primary purpose of a titration with a mercurial is to determine the
number of reactive SH groups which an enzyme possesses, either normally
or after being treated in various ways (e.g., denatured with urea, guanidine,
INHIBITION OF ENZYMES 799
or high temperature), and in this respect the problem is no different from
that of protein titration (page 762). We are interested here not so much
in the number of SH groups on an enzyme, but how these SH groups relate
to the catalytic activity and the mechanisms of mercurial inhibition, and
thus in following simultaneously the loss of SH groups and the develop-
ment of the inhibition as more and more mercurial is added. Let us assume
that we have a solution of a pure enzyme and we add to this a solution
of p-MB, or other mercurial, so that the molar ratio of mercurial to enzjnne
is slowly increased, and further assume that we allow time for the reaction
to come to equilibrium. We measure the number of SH groups reacted
(spectrophotometrically, polarographically, argentimetrically, or otherwise)
and the enzyme activity. There are only a few fundamental relationships
between mercaptidization and inhibition that could emerge, and these are
illustrated in Fig. 7-19. These are extreme situations, of course, and inter-
mediate behavior would more often be expected. Let us consider what each
result may mean and how valid certain interpretations may be.
Case A: The inhibition runs parallel to the SH groups reacted.
(1 ) The only SH groups that react are at the active center and mercaptide
formation abolishes the enzyme activity; titration to 100% inhibition will
give the number of SH groups at the active center.
(2) There are n equireactive SH groups on the enzyme, but only a cer-
tain fraction of these is at the active center or involved in the catalysis;
the titration will not provide the actual number at the active center.
(3) The SH group or groups are not at the active center, but reaction
of them leads to inactivation of the enzyme by some means; titration will
not provide useful information.
It is impossible to distinguish between these possibilities by simple ti-
tration nor can one determine accurately the number of SH groups at the
active center. Let us assume that an enzyme has 10 reactive SH groups
totally but that complete inhibition occurs when only 3 are reacted. It has
sometimes been concluded that 3 SH groups are necessary for the enzyme
activity. This is not a valid conclusion. If 1 of the 3 groups were related in
some way to the activity, one would obtain the same data. In other words,
the number of equivalents of mercurial added, or the number of SH groups
reacted, to achieve 100% inhibition does not provide directly the number
of SH groups involved in the catalysis.
Case B: SH groups are reacted but inhibition does not occur.
The conclusion here is obvious: the reactive SH groups are not involved,
directly or indirectly, in the enzyme activity. It is, of course, possible that
there are SH groups at the active center, perhaps even functional, but that
they do not react with the mercurial under the experimental conditions.
800
7. MERCURIALS
Case C: Inhibition develops only after so many SH groups have reacted.
(1) The most reactive SH groups are not related to the activity of the
enzyme, but the less readily available ones are.
(2) None of the SH groups is directly related to the activity, but when
a sufficient number are reacted the enzyme is structurally altered so that
inhibition appears.
These situations may be distinguished sometimes by determining the
reversibility of the inhibition, but occasionally denaturation is reversible.
Case D: Inhibition develops completely before all the SH groups react.
(!) The most reactive SH groups are at the active center and when they
P-MB m^
D
1
/
/ /
/ /
/ /
/ /
/ /
i'
F
/ /
/ /
' /
/
/
/
/
/
/
/
/
Fig. 7-19. Illustrations of the various relation-
ships between the reaction of enzyme SH
groups with mercurials and the inhibition of
enzyme activity. These are all extreme cases
and intermediate situations can also occur.
p-MB has been designated but any mercurial
may be used. The solid lines show the inhi-
bition and the dashed lines the reaction with
SH groups; i is the fractional inhibition and r
is the fraction of the total SH groups reacted.
INHIBITION OF ENZYMES 801
are all combined with the mercurial the enzyme is completely inhibited,
the less readily reacting SH groups being now available.
(2) Only a small number of SH groups are normally reactive, but follow-
ing mercaptide formation the enzyme unfolds and exposes other SH groups,
the groups reacted after total inhibition being secondarily released.
As discussed under Case A, the titration to 100% inhibition does not
necessarily provide the number of SH groups involved in the catalysis.
Reversal experiments run in parallel to the titration may give some infor-
mation as to which mechanism is involved.
Case E: Inhibition parallels the SH group reaction but levels off.
One must assume that complete combination of the enzyme with the
mercurial does not produce complete inhibition, which could easily be the
case if the SH groups reacted are sufficiently far from the active center so
that the introduced side chain would modify only the catalysis. A control
without the enzyme is, of course, mandatory to eliminate nonenzymic com-
ponents to the rate.
Case F: Inhibition develops without the reaction of SH groups.
(1) The mercurial, which was assumed to be specific for SH groups, is
not, and is inhibiting by another mechanism.
(2) Reaction of SH groups leads to structural changes by which more
SH groups appear; lack of reaction might be concluded from titrations in
which residual SH groups are determined, not by direct spectrophotometric
methods.
(3) The method for the estimation of SH group disappearance may be
faulty, i.e., those groups reacted by the mercurial may be resistant to the
titrating agent.
There are some instances in which non-SH enzymes are inhibited by
mercurials, but in general a result of this type should suggest a re-examina-
tion of the methods used.
Let us now examine a simple system in greater detail. An enzyme having
2 SH groups per molecule, one of these groups involved in the catalysis and
one not, is treated with increasing amounts of a mercurial; how will the
relative reactivities of the SH groups affect the results? Representing the
mercurial by M, the two equilibria may be written:
802 7. MERCURIALS
If we assume that the mercurial is so tightly bound that fiee mercurial in
the medium may be neglected, the conservation equations take the form:
(M,) = (MSJ + (MS,) (7-8)
(Sr). = (Si-) + (MSJ (7-9)
(Sr)e = (S,-) + (MS,) (7-10)
The fractions of each S~ group combined with mercurial will be represented
by fy and f^, and it is easy to show that these are related by:
l-/i 1-/2
The ionization of the SH groups has purposefully been assumed to be com-
plete in order to simplify the expressions. If S^" is assumed to be necessary
for enzyme activity, i = fi = (MSi)/Si-)^, while the reaction of Sg" is with-
out effect on the activity. Finally, we shall designate by r the fraction of
the total SH groups reacted with the mercurial, this being determined by
titration. In a specific case where (S^-)^ = (82")^ = 10"^ M, if we vary the
ratio K2IK1 — i.e., the relative affinities of the S~ groups for the mercurial —
the curves in Fig. 7-20 are obtained, r always being linear to complete reac-
tion while i can follow any of the curves between KJE^ = 0 and 00. When
K2IK1 < 1, the situation corresponds to case C in Fig. 7-19, and when
KJKy > 1, it corresponds to case D. One thing we immediately note is
that the affinities of the mercurial for the different S" groups must be quite
different if the i curves are to deviate from the r line appreciably; i.e.,
unless K2IK1 is much greater or much less than 1, it will be difficult to
demonstrate that only one of the S~ groups is necessary for the enzyme
activity. This treatment can be readily extended to enzymes with more
than 2 S~ groups, in which case:
fiKi fiKz f„K„ (7-12)
1-/1 1-A ■■■ l-/n
and to situations in which more than one S" group are involved in the
catalysis. One can also plot i against r to obtain curves characteristic of
the various situations described above.
A few examples of the different types of behavior are summarized here,
as far as it is possible to evaluate the data published.
Type A
Lactate dehydrogenase — beef heart (Millar and Schwert, 1963)
Malate dehydrogenase — pig heart (Wolfe and Neilands, 1956; Pfleiderer et al., 1962)
3-Phoshoglyceraldehyde dehydrogenase — yeast (Velick, 1953)
Pyrophosphatase — pig brain (Seal and Binkley, 1957)
Succinate dehydrogenase — rat liver (Hirade and Hayaishi, 1953)
INHIBITION OF ENZYMES
803
0 8-
0.6
0.2-
FiG. 7-20. Theoretical curves showing the relationship between
reaction of enzyme SH groups and inhibition of the enzyme ac-
tivity for an enzyme containing two SH groups, only one of which
is involved in the catalytic activity. (SHi)^ = (SHa)^ = 0.01 mM
and K2/K1 varied as indicated by the numbers on the curves; r is
the fraction of SH groups reacted and is represented by the
straight line. It is assumed that free mercurial is zero.
Type B
Catalase — beef liver (Schiitte and Niirnberger, 1959)
Enolase — rabbit muscle (Holt and Wold, 1961)
Hexokinase — yeast (Fasella and Hammes, 1963)
Type C
ATPase — moysin (Singer and Barron, 1944; Barany, 1959; Gilmour and Gellert,
1961)
Alcohol dehydrogenase — yeast (Barron and Levine, 1952)
Aldolase — rabbit muscle (Swenson and Boyer, 1957; Szabolcsi and Biszku, 1961)
/?- Amylase — barley (Rowe and Weill, 1962)
Malate dehydrogenase — pig heart (Thome and Kaplan, 1963)
Phosphorylase — rabbit muscle (Madsen and Cori, 1955, 1956)
Rhodanese — beef liver (Sorbo, 1963)
Urease — jack bean (Hellerman et ah, 1943)
Xanthine oxidase — milk (Gilbert, 1963)
Type D
Aldehyde oxidase — rabbit liver (Rajagopalan et al., 1962)
804 7. MERCURIALS
Carbamyl-P synthetase — frog liver (Marshall et ah, 1961)
Cytochrome c reductase — calf liver (P. Strittmatter, 1959)
Malate dehydrogenase — beef heart (mitochondria) (Siegel and Englard, 1962).
Type E
Lactate dehydrogenase — pig muscle (Jecsai and Elodi, 1963)
Phosphoglucomutase — rabbit muscle (Milstein, 1961)
Phosphorylase — potato (Lee, 1960 b).
No clear-cut example of type F has been reported, but presumably the
inhibition of D-amino acid oxidase by p-MB, which is competitive with
respect to the benzoate portion of the inhibitor, would fall into this category.
Certain enzymes have been shown to have a single SH group at the active
center and necessary for activity, namely, papain (Kimmel and Smith, 1957;
Finkle and Smith, 1958; Sanner and Pihl, 1963), ficin (Liener, 1961), and
glycerol-P dehydrogenase (van Eys et al., 1959). They seem generally to
belong to type A.
A few selected titration curves may further illustrate the relations be-
tween SH reaction and inhibition. The titration of 3-phosphoglyceraldehyde
dehydrogenase with p-MPS has been discussed (Fig. 7-12) and is seen to
follow type A behavior (deviating toward type C), although the release of
NAD is not exactly parallel to the disappearance of SH groups. Yeast alcohol
dehydrogenase contains 10-12 SH groups per molecule but some inhibition
occurs when only one is reacted, although the curve (Fig. 7-21) shows the
inhibition at first to lag behind; it is difficult to know if this is type C or D.
ATPase presents a more complex situation (Fig. 7-22) since reaction of the
first 4 SH groups seems to produce only some stimulation of the activity,
reaction of the next 2 SH groups causing complete inhibition. Other prop-
erties of myosin, e.g., the ability to complex with actin and the viscosity
response of actomyosin to ATP, are more directly dependent on the reac-
tion of the first SH groups. ITPase activity conforms more to type A be-
havior. The relationship of ATPase activity to SH reaction depends on the
state of the enzyme (Fig. 7-23), no initial stimulation being observed when
EDTA is the activator instead of Ca++. The titration of muscle phosphor-
ylase a gives partial type C behavior, but over most of the range there is a
linear relationship between SH reaction and inhibition (Fig. 7-24). Since
the inhibition may be completely reversed by cysteine, it is unlikely that
a secondary inactivation is involved. Microsomal cytochrome c reductase
demonstrates typical type D behavior, one SH group being closely related
to the enzyme activity, as shown by the extrapolation of the inhibition
curve to complete inhibition (Fig. 7-25), from which it may be estimated
that K2IK1 is around 25-50.
Accurate, reliable, and directly interpretable titrations of enzymes are
not easy to perform in some cases. Some of the possible difficulties which
may arise will be summarized. (1) There is a failure to reach equilibrium,
INHIBITION OF ENZYMES
805
i.e., reaction of the mercurial is not complete at the time chosen for the
readings. Some enzyme SH groups react almost instantaneously with mer-
curials and others require 30-60 min at least (see page 809). Kinetic studies
should always accompany any titration; to decide arbitrarily that n min-
utes of incubation with the mercurial at each concentration is adequate is
not a satisfactory procedure. (2) The enzyme may be altered structurally
by the mercaptide formation so that new SH groups are progressively ex-
FiG. 7-21. Titration of yeast alcohol clehydrogenase
with 77-]\IB. sliowiny the nonlinearity at low con-
centrations of the mercurial. (From Barron and
Levine, 19.-i2.)
posed, in which case there is no clear end-point and the results do not
correspond to the original native enzyme. Sometimes the stability of the
enzyme can be increased by creating a more physiological environment.
(3) The enzyme SH groups may be oxidized during the titration, reducing
the number of titratable SH groups. Use of oxygen-free solutions and a
nitrogen atmosphere often eliminates this problem. (4) The mercurial may
react with other groups or other components of the enzyme system, e.g.,
in the spectrophotometric titration with p-MB or jj-MPS, causing absorp-
tion changes unrelated to SH groups. (5) The mercurial may do something
that secondarily alters the ultraviolet absorption, e.g., split off a coenzyme,
as demonstrated for NADH: lipoamide oxidoreductase (Palmer and Massey,
1962). (6) The presence of substances, especially buffers, reacting with the
806
7. MERCURIALS
mercurial may alter the rate and extent of reaction with the SH groups.
The titrations of yeast alcohol dehydrogenase by p-MB in phosphate and
in THAM buffers at pH 7.5 are quite different (Hoch and Vallee, 1960).
It is probably advisable to reduce the buffer concentration as far as possible.
(7) The number of reactive SH groups on an enzyme and the titration of
these groups vary with several experimental conditions, such as pH, temper-
ature, and absence or presence of substrate, and the question often arises
100
80
60
40
S 20
-20
SENSITIVrTY OF
ACTOMYOSIN TO
ATP
8 , 6
SH (M/IO G MYOSIN)
Fig. 7-22. Titration of myosin with mersalyl, showing
the effects on ATPase activity, the ability to form
actomyosin, and the sensitivity of the actomyosin
to ATP (measured by viscosity changes). (From
Barany, 1959.)
as to what conditions are optimal. Titrations are often done at unphysi-
ological pH's because reaction is faster or more complete, but it must be
remembered that the results do not necessarily apply to the enzyme under
normal conditions. Boyer and Segal (1954) showed definite difference in
the titration of 3-phosphoglyceraldehyde dehydrogenase spectrophotome-
trically at pH 4.6 and 7, and this is probably a general phenomenon. The
effect of temperature is weU illustrated by the study of yeast hexokinase,
INHIBITION OF ENZYMES
807
the SH groups at the active center becoming unavailable for reaction below
30° (Barnard and Ramel, 1962). The presence of substrate may either fa-
cilitate reaction of SH groups — as with xanthine oxidase (Fridovich and
Handler, 1958) and myosin ATPase (Gilmour and Grellert, 1961) — or
protect certain SH groups. The question as to which pH, temperature, and
medium should be used, or whether substrate or coenzyme should be pres-
ent during the incubation, can only be answered generally by stating that
REACTION WITH
SH GROUPS
0.25
020
0.05
16 20
MMOLESiIO^/MG
Fig. 7-23. Titration of myosin ATPase with p-MB, showing
the different responses of the EDTA-treated and Ca++-acti-
vated activity. (From Kiellye and Bradley, 1956.)
whenever possible one should strive for physiological conditions. It is nec-
essary, of course, to vary these factors in many instances in order to study
the behavior of the SH groups, but the variation should be from a standard
set of conditions designed to provide information relevant to the enzyme
in a normal state.
It is frequently difl&cult to determine with certainty the total number of
free SH groups in a native enzyme under standard conditions and especially
to relate certain SH groups to the catalytic activity. Thorne and Kaplan
(1963) titrated pig heart malate dehydrogenase with p-MB, allowing 1 hr
808
7. MERCURIALS
for reaction at 25°, and could obtain no reliable end-point. As the molar
ratio of I : E is increased there is no marked effect on the activity, except
for a slight stimulation, until after a value of 5 is exceeded, and then there
is a progressive loss of activity as the ratio is elevated, nearly complete inhi-
bition occurring at a value of 21.6. It is impossible to interpret these data in
terms of relating SH groups to activity. Indeed, it is likely that the enzyme
is structurally altered so that SH groups normally not accessible are second-
arily unmasked, since good titrations can be determined with the urea-
100
2 4 6 8 10
MOLES p-MB/MOLES ENZYME
Fig. 7-24. Titration of muscle phosphorylase a with
p-MB. (From Madsen and Cori, 1955.)
denatured enzyme. The aspartate: or-ketoglutarate transaminase from pig
heart contains a total of 7 SH groups; when 2 moles of p-MB per mole of
enzyme are added, this reacting with 1-2 SH groups, the activity is reduced
by 50% (Turano et al., 1963). Such data again are uninterpretable and it is
impossible to conclude that SH groups are related in any way to the cat-
alysis. Di Sabato and Kaplan (1963) titrated the lactate dehydrogenases
from a variety of sources with both Hg++ and p-MB. The total number of
SH groups per mole of enzyme varied from 17 to 27 but generally inactiva-
tion occurred when 4 moles of mercurial were bound for each mole of en-
zyme. It is likely that no major configurational changes occur because no
alterations of fluorescence, sedimentation constant, or rotatory dispersion
and no immunological changes could be detected, and furthermore cysteine
could essentially completely reverse the inhibition. They felt that certain
SH groups are part of the active site rather than being vicinal, since statis-
INHIBITION OF ENZYMES
809
tically it is unlikely that in all the dehydrogenases such a distribution would
occur. On the other hand, Jecsai and Elodi (1963) claimed that pig muscle
lactate dehydrogenase in the native state does not react with p-MB, but
that at pH 10 the blocking of 20 SH groups leads to 50% inactivation. They
concluded that in this particular enzyme the SH groups are not at all in-
volved in the catalysis. These examples only illustrate some of the problems
which arise in enzyme titrations and emphasize that a program for relating
SH groups to enzyme activity cannot be undertaken lightly.
100
^ 20
Fig. 7-25. Titration of microsomal cytochrome c
reductase with p-MB. The sohd curve gives the
development of the inhibition as equivalents of
mercurial are added. The dashed line continuing
from the initial linear portion of the curve shows
that one SH group is required for activity; the
other dashed curve shows the assumed reaction
with SH groups (it was not experimentally de-
termined). It may be estimated that K^IK^ is
near 25-50 if there are two SH groups. (From P.
Strittmatter, 1959.)
Kinetics of Mercaptide Formation and Development of Inhibition
Apparently SH groups range in reactivity all the way from those which
combine with mercurials so rapidly that the rates are difficult to measure,
to those which are completely blocked and do not react at all. It is thus
810
7. MERCUKIALS
not surprising that one finds a great deal of variation in the rates at which
enzyme SH groups react and at which inhibition occurs. In some cases the
inhibition has been said to appear instantaneously, or to reach full magni-
tude within 1-2 min; such is the inhibition of succinate dehydrogenase
(Fig. 1-12-12) (Slater, 1949), bromelain (Murachi and Neurath, 1960), pyru-
vate decarboxylase (Stoppani et al., 1953), and leucine decarboxylase (Sut-
ton and King, 1962). Then there are enzymes which require about 5 min
for maximal inhibition to develop; examples are catalase (Cook et ah, 1946),
transaminases (Grein and Pfleiderer, 1958; Segal et al., 1962) and phospho-
glucomutase (Milstein, 1961), although in the last instance only 2 of the
3 SH groups react so rapidly. It is interesting to note that Nygaard (1955)
has reported marked differences in rates between the mercurials, lactate
dehydrogenase being very rapidly inhibited by Hg++ but only slowly by
p-MB. The next group of enzymes seems to require about 15-20 min for
complete inhibition: 3-phosphoglyceraldehyde dehydrogenase (Boyer and
Segal, 1954) and enolase (Malmstrom, 1962) may be cited. These are, of
course, arbitrary categories and if one knew the rates of reaction for many
100
80
60
40
20
0
.
^______
/35°
^^ "^
/ /
/zo"
(/
ALCOHOL
DEHYDROGENASE
0 20
TIME »
40
60
80
MIN
Fig. 7-26. Rates of inhibition of liver alcohol
dehydrogenase by p-MB, at pH 7.6 and two
different temperatures. ADH = 1.78x IQ-s M,
p-MB = 10-« M, and NAD = 3x10"* M.
(From WaUenfels et al, 1959.)
enzymes, there would be a continuous distribution, and furthermore the
rate in any particular case will depend on a number of factors, so that the
values given above and below must be taken as applying only to the ex-
perimental conditions imposed on each enzyme.
More interesting are those enzymes which react slowly enough with mer-
curials for the kinetics to be investigated. Some typical curves for p-MB are
given in Figs. 7-26 and 7-27, and similar rate curves have been previously
presented for cholinesterase (Fig. 1-12-8) and lactate dehydrogenase (Fig.
1-12-11). The results in Figs. 7-26 and 7-27 have been exponentially plotted
INHIBITION OF ENZYMES
811
in Fig. 7-28 to indicate more clearly the relative rates of inhibition (see
Eq. 1-12-14 and Figs. 1-12-3 and 1-12-9), the slopes being proportional to
the bimolecular rate constants. One notes that most of the curves deviate
from linearity, frequently at high inhibitions; this is probably an expression
of the different relative reactivities of the SH groups on. a single enzyme,
some reacting initially at a rapid rate and others reacting more slowly.
The rate constants have been calculated for some enzymes, e.g., 18.8 liters/
mole/sec for glutamate decarboxylase (Shukuya and Schwert, 1960), 51 li-
ters/mole/sec for muscle phosphorylase (Madsen and Cori, 1956), and 61.4
liters/mole/sec for heart lactate dehydrogenase (Takenaka and Schwert,
1956), in all cases p-MB being the inhibitor. The effects of temperature and
mercurial concentration on the rates of inhibition are well illustrated for
alcohol dehydrogenase (Fig. 7-26) and /?-fructofuranosidase (Fig. 7-27), re-
spectively. Glutamate decarboxylase presents an interesting phenomenon,
in that exposure of the enzyme to low temperatures appears to liberate
additional SH groups (Fig. 7-27).
)9 - FRUCTOFURANOSIDASE
GLUTAMATE DECARBOXYLASE
80 " 0 60 120 180 240 300 360
MIN TIME »- MIN
Fig. 7-27. Rate titrations of various enzymes with p-MB. The reaction
of SH groups was determined by absorption changes at 250 m/<. Phos-
phorylase a from rabbit muscle: p-MB = 0.04 mM, pH 6.7, and 21°
(Madsen and Cori, 1956.) Lactate dehydrogenase from heart: p-MB
= 0.00448 Mm, pH 6.8, and 25° (Takenaka and Schwert, 1956.)
^-Fructofuranosidase from Neurospora: p-MB concentrations given in
the graph, pH 6.8, and 0° (Metzenberg, 1963). Glutamate decarboxylase
from E. coli: preincubation with p-MB for 4 hr at either 0° or 25°,
and reaction run at 25° and pH 6.5 (Shukuya and Schwert, 1960).
812
7. MERCURIALS
Progressive inhibition or inactivation of enzymes by mercurials is very
common and takes a variety of forms. Epididymal a-mannosidase is inhi-
bited 62% by 0.01 mM Hg++ without preincubation with the inhibitor;
the inhibition is 67% at 30 min, 77% at 60 min, and 88% at 120 min
(Conchie and Hay, 1959). This is one of the numerous examples in which
an enzyme is rapidly inhibited to a certain level, further increase in inhibi-
FiG. 7-28. Logarithmic plots of if— i for the en-
zymes in Figs. 7-26 and 7-27, either loss of ac-
tivity or reaction of SH groups being used as a
measure of the reaction with p-MB. These curves
have been estimated from the published curves
and hence are not strictly accurate, but indicate
the relative rates for the more slowly reacting
SH groups. 1, Alcohol dehydrogenase (35°);
2, Alcohol dehydrogenase (20"); 3, phosphorylase;
4, lactate dehydrogenase; 5, glutamate decar-
boxylase (25°); 6, glutamate decarboxylase (0");
7, ^-fructofuranosidase (0.1 mM); 8, ^-fructo-
furanosidase (0.02 raM); 9, j3-fructofuranosidase
(0.01 mM); 10, /3-fructofuranosidase (0.04 mM.)
tion being slow. Such behavior is not surprising when one measures SH
reaction, since one assumes generally the occurrence of SH groups of dif-
ferent reactivities; thus when 3-phosphoglyceraldehyde dehydrogenase is
titrated with p-MB, 11 SH groups react immediately, but 3 more require
at least 40 min (Koeppe et al., 1956), and aldolase behaves very similarly,
7 SH groups being blocked rapidly and 3-4 more groups taking 40 min for
reaction (Szabolcsi and Biszku, 1961). But the interpretation of inhibition
following such a time course is not so clear. If a certain level of inhibition
is reached rapidly and then the rate falls off markedly, one must assume
that the enzyme is not completely mhibited when its rapidly reacting SH
INHIBITION OF ENZYM.ES 813
groups are combined with mercurial (assuming that there is sufficient mer-
curial to react with all these groups). The more slowly developing inhibition
could be due to reaction of less readily available SH groups or to a secondary
inactivation following the initial mercaptide formation. Quite different re-
sults are obtained with amylase, mersalyl at 1 mM not inhibiting at all
during the first hour, but slowly inhibiting until there is 40% depression
after 48 hr (Muus et ah, 1956), or with bromelain, jJ-MB inhibiting only
25% after 4 hr and 80% after 20 hr at 0.1 mM (Ota et al, 1961). Many
different time courses of inhibition are observed and it is likely that the
major factors involved are (1) the relative reactivities of the SH groups,
(2) the relationship between the SH groups and the catalytic activity, and
(3) the tendency for structural changes leading to inactivation to occur.
However, it is quite clear that many enzymes react quite slowly with mer-
curials and require 2-4 hr (and occasionally more) to complete the process.
Such enzymes are difficult to titrate, since one does not know how many
of the SH groups finally reacted were originally present, and, when one is
adding increasing amounts of mercurial to correlate mercaptide formation
and inhibition, it is not easy to decide on the optimal preincubation in-
terval.
In the previous section the correlation between inhibition and SH reac-
tion by mercurials was considered in terms of variable quantities of mer-
curial. Another approach to relate these phenomena is to determine their
changes with time at a particular mercurial concentration. If the SH groups
which are combined initially are necessary for enzyme activity, one would
expect inhibition to parallel blocking of these groups; if the most readily
reacting SH groups are not related to activity, or the enzyme undergoes
progressive inactivation, the inhibition may lag behind mercaptide forma-
tion. Madsen and Cori (1956) observed that inhibition of phosphorylase by
p-MB developed more slowly than the change in absorbance at 250 mjit
(Fig. 7-27), so that when 50% of the reactive SH groups had been blocked
the inhibition was only 14%. If the SH blocking itself is not responsible
directly for the inhibition, but initiates an unfolding of the enzyme, the
rate of inhibition may be more dependent on the rate of configurational
change. In the case of phosphorylase, we have seen that splitting into sub-
units occurs during reaction with p-MB, so the rate at which this occurs
may have something to do with the inhibition rate. Inasmuch as cysteine
reverses the inhibition completely, marked structural alterations would not
be very likely.
Another phenomenon which must be taken into account in kinetic studies
is the spontaneous recovery of enzyme activity in the presence of the mer-
curial, first observed, I believe, by von Euler and Svanberg (1920) in studies
of the inhibition of yeast /5-fructofuranosidase by Hg++. Reisberg (1954)
reported that the inhibition of choline acetylase by p-MB is less at 30 min
814
7. MERCURIALS
than at 10 min. Other more recently observed examples of this include
epididymal /5-galactosidase with low concentrations (0.0002 milf ) of Hg++
(Conchie and Hay, 1959), xanthine oxidase with 0.44 mM p-MB, which
inhibits NADH oxidation 50% initially but less and less as the reaction
proceeds (Westerfeld et at., 1959), and leucine decarboxylase with 0.005 mM
p-MB (Sutton and King, 1962). The most marked spontaneous recovery is
seen with pig heart lactate dehydrogenase, the rate and degree of reactiva-
tion being dependent on the molar ratio of p-MB to enzyme (Fig. 7-29)
50
HOURS
Fig. 7-29. Effects of p-MB on pig heart
lactate dehydrogenase, showing the ini-
tial inhibition and the spontaneous reac-
tivation. The numbers on the curves
are the molar ratios of p-MB to LDH.
(From Gruber et al., 1962.)
(Gruber et al., 1962). The most common explanation for such recovery is
a slow migration of the mercurial from those groups initially attacked to
other groups not involved in the enzyme activity. The rates at which var-
ious SH groups react with a mercurial are not necessarily related to the
affinities of the groups for the mercurial. Groups which bind the mercurial
very tightly may be masked and react very slowly, as fairly conclusively
demonstrated for myosin ATPase by Gilmour and Gellert (1961). Another
factor which may be of importance when the inhibition decreases during
the period when the enzyme activity is measured, as was the case with
leucine decarboxylase, is the displacement of the mercurial by the substrate.
Leucine was shown to protect the enzyme against 2)-MB and it could even-
tually overcome the inhibition somewhat, especially since its concentration
was some 1000 times greater than the mercurial. A substrate might also be
able to restore toward a normal configuration a slightly luxated active cen-
ter, substrates being known to stabilize the active forms of certain enzymes.
INHIBITION OF ENZYMES 815
This phenomenon of spontaneous recovery must be even more common in
celkilar preparations than with pure enzymes, because there is much greater
opportunity for redistribution of the mercurial.
Stimulation of Enzymes by Mercurials
Mercurials is common with other heavy metals and SH reagents frequent-
ly increase enzyme activity, especially at low concentration, the action-
concentration curves being biphasic. Polis and Meyerhof (1947) first ob-
served the stimulation of Ca++-activated myosin ATPase by PM, a 30-40%
elevation of the rate occurring with concentrations between 0.005 and 0.12
mM, and this has been confirmed in several more recent reports, the degree
of stimulation, however, varying greatly with the experimental conditions.
Many different types of enzyme exhibit this phenomenon (Table 7-11) but
the mechanisms involved have only rarely been clarified. Let us briefly
consider some possible mechanisms and what relevant evidence is available.
(A) The mercurial inactivates a naturally occurring inhibitor. If an in-
hibitor is isolated w^ith the enzyme and is suppressing the activity, and if
this inhibitor is an SH protein (as many natural inhibitors seem to be), a
mercurial by reacting preferentially with the inhibitor may release the en-
zyme from its inhibition. J. S. Roth (1953 a, 1956, 1958) has been a pro-
ponent of this theory with respect to the activation of rat liver homogenate
ribonuclease by p-MB or PM, and has found a natural inhibitor with which
the mercurials react at concentrations having no direct effect on ribonu-
clease. The degree of stimulation varies with the tissue from which the ri-
bonuclease is obtained — all the way from 0% with pancreas, 57% with
brain, 104% with muscle, 284% with liver, to 1500% with ascites carci-
noma — and this may be due to the different amounts of inhibitor present
(Ellem and Colter, 1961). Indirect evidence often points to such a mechan-
ism for other enzymes. PhiUips and Langdon (1962) found that p-MB stim-
ulates microsomal NADPHxytochrome c reductase but only inhibits the
purified enzyme. Of course, the activation could also be due to some effect
of the mercurial on the microsomal structure. It has also been noted occa-
sionally that stimulation occurs, and is relatively constant, over a wide
range of mercurial concentration, inhibition appearing rather suddenly when
this range has been exceeded, and this indicates some component with which
the mercurial reacts readily and completely.
(B) The mercurial reacts with the substrate to labilize it. Ledoux (1953)
initially attempted to explain the stimulation of ribonuclease by p-MB as
due to a reaction with RNA, this favoring in some manner the enzymic
hydrolysis, and detected spectral changes upon mixing RNA and p-MB
(see page 741). Although mercurials do complex with nucleic acids, it is
doubtful if this is a major factor in the activation. The proper preincubation
816
7. MERCURIALS
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818 7. MERCURIALS
procedures might easily solve this problem. There is no other instance where
this mechanism has been postulated, but it is a possibility that must be
borne in mind.
(C) The mercurial reacts with a SH group to create a more favorable electric
field at the active center. Since the binding of the substrate and the break-
down of the ES complex are influenced by the electric field arising from
charged groups vicinal to the active center, it is possible that the binding
of a charged mercurial could so alter this field as to facilitate the catalysis.
There is no evidence for this and it would be difficult to obtain.
(D) The mercurial increases the active form of the enzyme. Certain enzymes
within the cell or as extracted may be in an inactive form, possibly with
the active center not in the proper configuration. Reaction with a mercurial
might so alter the protein structure as to release the activity, much as
some enzymes can be activated by heat. This would probably apply only
to true activations, i.e., where the initial state of the enzyme is inactive,
as in the work of Hilz and Klempien (1959) on ascites tumor ribonuclease,
Hg++ at 0.005 and 0.05 uvM increasing the enzyme rate from 0 to 12 and
24 ^moles TCA-soluble phosphate/hr, respectively, but here it may be only
that the enzyme is completely inactivated by a natural inhibitor.
(E) The Hg^+ ion may replace a normal metal ion activator. Carboxypepti-
dase which is normally activated by Zn++ can be activated with respect to
the esterase activity with Hg++ (Coleman and Vallee, 1961). This situation
is probably very uncommon. Rapoport et al. (1955) reported that Hg++
stimulates glycerate-2,3-diphosphatase only in the presence of certain or-
ganic nitrogen substances, and the activation was assumed to be due to
the complex formed. Substances which are necessary for the Hg++ effect
include nonenzyme proteins and certain amino acids, of w^hich histidine is
the most effective (Sauer and Rapoport, 1959). It was concluded that in
addition to an SH group, activity requires the presence of some metal ion
and some complexer bound in a cylic resonance system:
COO
CHo — CH
Resonance
system
NH,
Enzyme
Me '^
^ cf
with the enzyme. In this case, Hg++ would simply function as a metal ion
for linking the resonating system to the enzyme.
(F) The mercurial may disrupt water structure. The structure of water
around the active center may be such as to retard somewhat the access of
INHIBITION OF ENZYMES 819
the substrate. Reaction of a mercurial with a vicinal SH group could by
introducing a new side chain break down this water structure.
(G) The mercurial reduces the binding of an inhibitory 'product. L-Gluta-
mate dehydrogenase is stimulated by PM at pH 8.5 and this is reduced by
substrate, NAD, and glutarate, a competitive inhibitor (Greville and Mild-
van, 1962). Thus PM must combine at or near the active center. The prod-
uct of the reaction, a-ketoglutarate, is inhibitory. PM increases the K^ for
a-ketoglutarate 8-fold and for glutarate more than 20-fold. Part of the stim-
ulation by PM can be due to reduction of the effects of a-ketoglutarate.
It would seem that such behavior would be reflected in the rate curves,
little stimulation being expected initially.
(H) The mercurial dissociates the e^izyme into active subunits. Some of the
active centers may be more accessible when the enzyme is disaggregated,
and it is known that mercurials can sometimes split enzymes into subunits
(page 788). GreviUe and Mildvan (1962) observed that PM dissociates glu-
tamate dehydrogenase, and Rogers et al. (1962) also noted effects on the
sedimentation properties. The possibility of such dissociation playing a role
in the mercurial activation was studied by Rogers et al. (1963), who found
no change in molecular weight upon treatment with MM when the enzyme
is in high concentration. However, when low enzyme concentrations were
used, a disaggregation sensitive to the mercurial was detected, but it is not
certain if this is related directly to the stimulation.
(I) The mercurial reacts primarily with an inhibitory SH groups. The stim-
ulation of ATPase by mercurials has generally been explained since the
report of Kielley and Bradley (1956) in terms of differently located SH
groups around the active center. An SH group, for example, might bind a
group on ATP and interfere with the optimal orientation on the enzyme.
This SH group has been postulated to react with the 6-amino group of ATP;
when mercaptide formation occurs, this discouraging action on ATP is abol-
ished (Gilmour, 1960; Greville and Tapley, 1960). In essence, the mercurial
prevents the excessive occupancy of the active center by disoriented ATP,
allowing ATP to proceed directly to hydrolysis. A somewhat different view
has been voiced by Blum (1960): ATP induces a configurational change in
the active center, this involving the SH groups, and mercurials at low con-
centrations tend to prevent this change. ITP does not so alter the structure
and its hydrolysis is inhibited only by mercurials. Mercurials would thus
maintain the active center in the configuration binding ITP, a state con-
ducive to rapid hydrolysis of ATP.
(J) The mercurial inhibits a second enzyme which suppresses the reaction
rate. A number of possibilities for stimulation were discussed in Chapter
1-7. In a monolinear chain:
E, Es
820 7. MERCURIALS
inhibition of the Eg will increase the steady-state level of B or its rate of
formation. If the formation of B is being measured and an enzyme destroy-
ing it is present, inhibition of this enzyme will appear to stimulate E^.
Likewise in a divergent chain, inhibition of one branch may increase the
rate of the other branch; whether stimulation will be observed will depend
on what is measured. In the incorporation of nucleotides into amino acid
transfer UNA, the presence of any enzyme attacking the nucleotides will
reduce the incorporation, and the inhibition of such an enzyme will ap-
pear to stimulate the incorporation. Stimulation was indeed observed with
p-MB by Starr and Goldthwait (1963), who thought that this might be
due to a contaminating phosphodiesterase, but Anthony et al. (1963) ob-
tained evidence that such an enzyme did not occur at significant levels in
their preparation.
The stimulation of many enzymes by mercurials is strongly pH-depend-
ent. The activating effect of Hg++ on glycerate-2,3-diphosphatase becomes
progressively less as the pH is increased beyond 7 (Rapoport et al., 1955),
and this is true also for ascites cell ribonuclease, although here the activa-
tion disappears as the pH is decreased to 5 (Colter et al., 1961). In the latter
case, a 50% stimulation changes to a 42% inhibition as the pH rises from 8
to 8.5. Liver mitochondrial ATPase stimulation by p-MB is optimal around
a pH of 9 and falls off rapidly on both sides (Myers and Slater, 1957 b),
and the optimal pH for activation of myosin ATPase is 7.8, little stimula-
tion being observed at pH 5.7 or 10 (Tonomura and Furuya, 1960). Such pH
effects may be important in working out the mechanisms of the stimulation
but have not so far been studied in enough detail to contribute evidence.
Temperature can apparently also play a role, since p-MB stimulates myosin
ATPase at 25° but only inhibits at 0^ (Fig. 7-30) (Gilmour and Griffiths,
1957). It is also evident in this figure that the DNP-activated ATPase is
not further stimulated by p-MB. The EDTA-activated ATPase is likewise
not stimulated by p-MB, whereas in the presence of Ca++ the stimulation
is marked (Fig. 7-23). Finally, the effect of mercurial concentration is oc-
casionally very striking, as in the case of myosin ATPase (Fig. 7-23), max-
imal stimulation by p-MB being observed when about half the SH groups
are reacted. Another example of variation of stimulation with mercurial
concentration is shown in Fig. III-l-l for ribonuclease, although here the
form of the curve may depend primarily on the nature and amount of
the natural inhibitor, as well as the susceptibility of the enzyme itself.
The problem of enzyme stimulation by mercurials, or other inhibitors,
is a very interesting one, and probably quite important in understanding
not only the mechanisms whereby the mercurials can affect enzymes but
some of the many instances of stimulation of metabolism, which will be
mentioned later in this chapter.
INHIBITION OF ENZYMES
821
Reversal of Mercurial Inhibition
The general treatment of the reversal of enzyme inhibition by substances
binding the inhibitor (page 1-615) wiU now be extended to those situations
often encountered in studies of the mercurials. The most common reversors
are thiols, e.g., cysteine, glutathione, mercaptoethanol, mercaptoacetate,
and dimercaprol (BAL), and the reduction in the inhibition can be consid-
ered as a transfer of the mercurial from enzyme SH groups to the reversor
(R) SH groups. In many reactivation experiments the concentration of free
mercurial is very small and the reversal reaction can be written as:
EI + R ±5 E
(E,) - X (Rt) - X X
+
RI
X
(7-13)
If the enzyme is initially completely inhibited, (EI) = (EJ, and the final
equilibrium concentrations are as written under the equation. The equilib-
rium is characterized by the following constants:
(EI)(R) [(E,) -x] [(Rt)-x]
(E) (RI)
(E) (I)
(EI)
K,
(R) (I)
(RI)
K
= Kr
(7-14)
Fig. 7-30. Effects of p-MB on myosin ATPase
at 0° and 25°, and in the presence of 2,4-di-
nitrophenol. The ATPase activity is given
as /^moles P,/mg/min. (From Gilmour and
Griffiths, 1957.)
822 7. MERCURIALS
The value of x, the concentration of active enzyme regenerated, can be
calculated from the quadratic equation:
x^{\ - K)-x [(R,) + (E,)] + (E,) (R,) = 0 (7-15)
if the total concentrations of enzyme and reactor are known. The value of
the final inhibition is given simply by:
If both enzyme and reversor SH groups ionize similarly, we need not in-
clude the effect of (H+), and since the mercurial is assumed to be bound
to either the enzyme or the reversor, we need not consider (X~), the con-
centration of mercurial-complexing ligands in the medium. The situation
described here is that in which an enzyme is titrated to complete inhibition
and the reversor then added. If any irreversible inactivation has occurred,
the experimentally measured inhibition will be greater than in Eq. 7-16.
In most reactivation experiments, (R^) is much greater than (E,), and
thus the reversor concentration is not reduced significantly by the binding
of the mercurial. The equation for the determination of x is simplified to:
a;=A' + a;(R,) - (E,) (R,) = 0 (7-17)
It is interesting to specify the conditions required to reverse the inhibition
to a determined value ij. Substitution of (E^) (1 — ij) for x in the solution
to the quadratic equation leads to
(7-18)
(7-19)
(E,) V K,
Equation 7-18 gives the ratio of the dissociation constants of the two mer-
captides so that the inhibition will be reduced to ij when the enzyme and
reversor concentrations are given, while Eq. 7-19 shows how much reversor
must be present relative to the enzyme to achieve a reactivation to if when
the dissociation constants are known. In many experiments the reversor is
added at a concentration of 1-10 vaM and thus (R^)/(EJ is often 10^ and
10^, since enzyme concentrations commonly run from 10"^ to 10^^ raM.
If we assume that K^ = K^, which is a reasonable approximation in many
cases, the values of (R^)/(E^) shown in the following tabulation must be
used to reduce the inhibition to the levels indicated. Thus one would expect
that the concentrations of reversor often used would completely abolish
the inhibition, so that if the enzyme activity is experimentally not restored
Kr
*/
(Rt)
K,
(1 — ifT-
(E,)
(Re)
(1 - if)'
Kr
INHIBITION OF ENZYMES 823
to normal values it is likely that (1) some inactivation has occurred, (2) in-
sufficient time has been provided for the reversal, (3) the binding of the
mercurial to the enzyme is much tighter than to the reversor (K^"^ K^),
Final % inhibition
(Ri)/(E,)
50
0.5
20
3.2
10
8.1
5
18
1
98
or (4) some secondary factor has complicated the situation, e.g., failure to
add a displaced cofactor (page 1-625) or the oxidation of enzyme SH groups
by the oxidized thiol reversor (page 1-625). If the complete reversibility
of the inhibition has been established, and if (E^) and K^ are known, it is
possible to calculate the value of K^ from experiments in which low con-
centrations of reversor are added and the ij determined.
It is relatively easy to treat equilibrium conditions quantitatively, but
the problems encountered in considering the rates of reversal are at the
present mainly insoluble. Investigators have been chiefly interested in whe-
ther reversal occurs or not and since extremely few rate studies have been
made, there are not adequate data upon which to base accurate analyses.
If the reversal occurs in two steps — the dissociation of EI and the combi-
nation of I with R — the individual reactions are not as simple as with most
inhibitions. For example, at physiological pH and with most media used,
the dissociation of EI must be written as:
EI + H+ + X- -► EH + IX (7-20)
where X~ represents any complexing anion in the medium. Likewise, the
second step must usually be described by:
IX + RH -> RI + X- + H+ (7-21)
Now it is not known if the reactions occur in this manner, or whether I
can be directly transferred from E to R without forming the IX complex,
either on the surface of the enzyme or in solution. A study of the effects
of (X^) on the reversal rate would help to solve this problem. It is also
not known if the reactive form of the reversor is RH or R~, but from the
effects of pH on the rates of reaction of mercurials with simple thiols it
appears that at least the form R" reacts much more rapidly than RH, as
would be anticipated. In any event, the pH must be an important factor
824 7. MERCURIALS
in the rates of reversal, although it may not markedly influence the equi-
librium conditions.
Despite the lack of data on reversal rates, one obtains the impression
that reversal is often a good deal faster than the development of inhibition.
Unfortunately those workers who measured the rates of inhibition the most
carefully usually did not examine the reversibility or, if they did, remarked
only that it occurs, without giving data on the time required; and in other
cases no reversal was observed. We are thus in the surprising position of
having essentially no information in any one case of mercurial inhibition
as to the relative rates of inhibition and reversal. Certainly in some instances
the reversal is very rapid, as with 3-phosphoglyceraldehyde dehydrogenase
(Velick, 1953) and hexokinase (Sols and Crane, 1954) treated with cysteine
after inhibition by p-MB, as previously discussed (page 1-623 and Fig. I-
13-8). However, in these cases the rates of inhibition are not known, al-
though they are probably quite fast; in both, the enzyme and p-MB were
preincubated for an arbitrary time (for hexokinase 15 min at 0°). It would
be difficult to compare the rates of reversal by dialysis and reversor — it
seems never to have been done experimentally — but dialysis is not a very
efficient method due to the fact that only a small fraction of the dissociated
inhibitor at any time is able to pass out through the membrane, most re-
combining with enzyme, while the presence of a high concentration of re-
versor throughout the medium ensures that free inhibitor is rapidly bound.
Therefore, one does not know in reversal which step is limiting, or if the
reversor can facilitate the dissociation of the mercurial from the enzyme.
It is impossible to predict the relative rates of inhibition and reversal theo-
retically because we do not know the exact reactions involved, particularly
the influence of H+ and complexing ions. There is only one way to solve
these problems: to do a few critical and accurate experiments.
The information to be derived from simple reactivation experiments,
especially those in which a high concentration of reversor is used, is not
as much or as reliable as many seem to have believed (see page 1-624). If
complete reversibility is obtained, this shows that no serious inactivation
of the enzyme has occurred; it does not imply that structural changes have
not been induced by the mercurial. This information is usually important
and must be obtained in any quantitative kinetic studies, but this is all
that this type of reversal experiment will provide. Failure to recover the
activity can be explained in a variety of ways, previously enumerated (page
651). More careful reversal studies, especially determination of rates, the
effects of pH and ligand concentration, and the degree of reactivation with
low reversor concentrations, would undoubtedly be more informative.
Several hundred studies on mercurial inhibition have included statements
relative to reversibility with thiols. It would serve very little useful pur-
pose to list these results. Summarizing all of them one finds that complete
INHIBITION OF ENZYMES 825
reversal was obtained in 59%, partial reversal in 30%, and no reversal in
11%. This shows at least that in most cases the enzyme is not seriously
altered or denatured by severe mercurial inhibition, providing the contact
with the inhibitor is not prolonged. On the other hand, there are the en-
zymes such as papain which are more stable when complexed with Hg++.
Some of the instances where partial or no reversal was found are undoubted-
ly due to inactivation, but some to the other factors mentioned above. In a
few studies, some interesting sidelights were noted and we shall content
ourselves with discussing some of these.
Two more instances of very rapid, almost instantaneous reversal of mer-
curial inhibition may be noted. Yeast alcohol dehydrogenase is immediately
inhibited by p-MB and likewise rapidly reactivated by glutathione (Snod-
grass et al., 1960). The enzyme at 4.5 X 10"^ M is completely inhibited by
2.5 X 10"'' M Hg++; glutathione reverses this inhibition rapidly, but par-
tially — after 30-sec exposure to Hg++ 54% and after 10-min exposure
only 17%. This failure to reverse is not due to displaced Zn++, since Zn++
was found not to be released from the enzyme so rapidly and the addition
of Zn++ does not improve the reversal. Heart lactate dehydrogenase is very
rapidly reactivated from p-MB inhibition by cysteine, and in this case the
rate of inhibition is relatively slow, about 15 min being required for maxi-
mal inhibition (Neilands, 1954). This is one of the few instances in which
reversal definitely seems to occur faster than inhibition. The effectiveness
of reversors in comparison with other methods for reactivation is seen with
succinate dehydrogenase. If the enzyme inhibited by p-MB is dialyzed for
3.5 hr there is no reactivation, but addition of glutathione reverses complete-
ly (Singer et al., 1956b). Also no reversal was observed following dilution of
the inhibited enzyme, whereas thiols reactivate partially (Slater, 1949). The
rate of reactivation of erythrocyte pyruvate kinase by 100 mM glutathione
after 1-min contact of the enzyme with 0.1 mM p-MB is fairly slow (see
accompanying tabulation); however, if contact with the inhibitor is 10 min,
Time
(min)
% Reversal
10 18
15 47
25 59
only 28% reversal is seen after 25-min exposure to glutathione (Solvonuk
and CoUier, 1955). There is no obvious explanation why the reversal rates
with some enzymes are so fast and in others so slow, especially as this is
not correlated at all with the aifinities of the enzymes for the mercurials.
826 7. MERCURIALS
Low concentrations of reversor in a definite molar ratio to enzyme or
mercurial have seldom been used, but in the examples we have the reversal
is only partial. Succinate oxidase preparation from pigeon muscle is inhi-
bited 88% after 15-min incubation with 0.03 mM Hg++; reversal by di-
mercaprol for 30 min depends on the amount of the dithiol added (see ac-
companying tabulation) (Barron and Kalnitsky, 1947). Myocardial succin-
Dimercaprol : Hg++ % Reversal
3.3 0
5 33
10 63
ate oxidase inhibited completely by p-MB can be 31% reactivated by glu-
tathione at a molar ratio to the mercurial of 1 : 1 and 65% at a ratio of 5 : 1
(Slater, 1949). Treatment of 3-phosphoglyceraldehyde dehydrogenase with
33-MB leads to changes in the optical rotation; if the exposure to the mer-
curial is only 1 min, cysteine at a molar ratio of 6 : 1 reverses these changes
around 75%, but the structural changes become progressively more irre-
versible (Elodi, 1960). To reverse the inhibition of glutamate dehydrogen-
ase by p-MB maximally (80%) it requires around 60 times as much glu-
tathione as mercurial* (Olson and Anfinsen, 1953).
* These experiments were done by varying the p-MB concentration from 0.003
to 3.3 mM and keeping the reversor, glutathione, at 10 mM, so that the more normal
conclusion is simply that the reversibility is less, the higher the mercurial concentration,
a phenomenon commonly observed with other enzymes. The plotting is ambiguous;
the final enzyme activity is plotted as % of the uninhibited enzyme, but the results
with inhibitor alone are not given, although they can be estimated from another figure,
so the degree of reversal is not immediately apparent. For example, when p-MB is
0.11 mM the inhibition is given elsewhere as 50%; after 10 mM glutathione (molar
ratio 90 : 1) the enzyme activity is given as approaching 80%, which I would call
60% reversal on one basis or 30% on another. However, this is stated to be 70%
reversal in the text. No incubation times were mentioned so possibly the failure to
achieve more reversal is due to an insufficient time with the reversor. Other than the
work on reversal, this investigation is an excellent, detailed, and quantitative study
on an enzyme, and thus illustrates a rather common phenomenon — the lackadaisical
and disoriented approach to inhibition reversal. In 90% of the reports in which re-
versal is determined, apparently an arbitrary (but high) concentration of reversor is
added, and the enzyme activity is measured after an arbitrary interval, so the conclu-
sions should usually be taken as arbitrary. If these remarks, and others scattered
throughout the book, can stimulate the performance of more accurate and interpretable
reversal experiments, my aim will be achieved.
INHIBITION OF ENZYMES 827
Unusual results on the effects of thiols on the inhibition of /?-fructofu-
ranosidase by Hg++ were observed by Gemmill and Bowman (1960), in
that dimercaprol reverses the inhibition to varying degrees, whereas both
cysteine and glutathione increase the inhibition (Table 7-12). The thiols
themselves, even at the highest concentrations used, do not significantly
affect the enzyme activity, so the additional inhibition cannot be attribut-
ed to an excess of the thiol. Several explanations are possible: (1) The cys-
teine and glutathione reduce enzyme disulfide groups to SH groups and
increase the binding of the Hg++ to the enzyme; (2) the Hg++ reacts with
both enzyme SH groups and these thiols to form E — S — Hg — S — R com-
plexes which are less active than the simple E — S — Hg+ complexes; or (3)
the R — S — HgX or R — S — Hg — S — R mercaptides formed are inhibitory
by a mechanism unrelated to enzyme SH groups. Whatever the explanation
applicable here, one would expect that these situations would occasionally
be important in the reversal of other enzyme inhibitions, particularly when
the bifunctional Hg++ is used, and possibly failure to achieve reversal in
some cases may be due to these reactions.
Complexities arise in some reversal studies and a few will be mentioned
briefly. When the aspartate: or-ketoglutarate transaminase from pig heart
is treated with p-MB there is inactivation but the pyridoxal-P remains
bound to the apoenzyme (Turano et al., 1964). Incubation for 10 hr at pH
6.4 and 4° with 0.35 mM p-MB, however, releases the coenzyme. The ad-
dition of pyridoxal-P restores the activity in the sense that addition of
glutamate is followed by transamination to form pyridoxamine-P, but the
further transamination to oxalacetate is lost. In this instance, reversal thus
occurs for only part of the normal reaction. Carboxy peptidase A is activat-
ed by Zn++ and apparently the Zn++ is bound to the SH groups of a single
cysteine residue and the a-amino group of a terminal asparagine (Coombs
et al., 1964). Zn++ protects the enzyme from inhibition by p-MB, but if the
apocarboxypeptidase is inactivated by p-MB, addition of Zn++ does not
restore the activity.
Spontaneous reversal of enzyme inhibition by p-MB occurs with 3-phos-
phoglyceraldehyde dehydrogenase of muscle (Szabolcsi et al., 1960), and
has occasionally been observed in other systems, particularly homogenates.
This has usually been attributed to redistribution of the mercurial from the
rapidly reacting SH groups to others which bind the mercurials more tight-
ly, but in this case it was thought that intermolecular rearrangements,
whereby eventually some of the enzyme is inactive and some completely
active, are responsible. This should not be too uncommon a phenomenon
if the proper mercurial concentrations are used.
828
7. MERCURIALS
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INHIBITION OF ENZYMES 829
Survey of Enzyme Inhibitions
Some enzyme inhibitions produced by the mercurials are given in Table
7-13. These represent only about a fifth of the enzymes studied. It is a
difficult matter to decide which results should be in the table. I have tried to
include principally two sorts of enzyme, those that are "important" and
those that are inhibited "potently." But which enzymes are important?
Every enzyme is important for a particular pathway, or a certain organism,
or the investigator who studies it. The following groups of enzymes have
generally been chosen: those in the glycolytic Embden-Meyerhof or pen-
tose-? pathways, the tricarboxylate cycle, electron transport systems, phos-
phate transfer, and central amino acid metabolism; those often involved
in cell function (such as choline acetylase, cholinesterase, carbonic anhy-
drase, etc.); and certain classic SH enzymes (such as urease). We now must
consider how the word "potently" should be defined. I have arbitrarily
chosen enzymes inhibited significantly (usually 50% or more) by concen-
trations of mercurial of 0.01 mM or less, since such enzymes could usually
be considered as having reactive SH groups at or near the active center.
It is evident that a certain enzyme differs often very markedly in suscepti-
bility depending on the tissue or species from which it is obtained, so that
one cannot speak of the sensitivity of NADH oxidase, for example, in quan-
titative terms without specifying which NADH oxidase is meant. At least
most will agree I think that the enzymes included in Table 7-13 are impor-
tant in common metabolic pathways and/or are inhibited quite potently.*
Another problem in presenting the inhibitions in Table 7-13 is that the
degree of inhibition by mercurials depends on a number of factors to a
greater extent than with most other inhibitors. Particularly important are
the pH (mainly because of competition of H+ with the mercurial for the
S~ group), the temperature (because of both the high temperature coeffi-
cients of mercurial reactions and the possibility of secondary thermal inac-
tivation), the composition of the medium (principally because of competi-
tion of anions with the S~ group for the mercurials), the period of the in-
cubation with the inhibitor (since in many cases the inhibition does not
attain a constant level), and the presence of impurities (most of which can
complex with the mercurials and reduce their effectiveness). There are too
many of these variables to include in the table. The purposes of the table
are to provide very roughly some information on the relative sensitivities
of the more important enzymes, and to present an experimental basis for
the appreciation of the lack of specificity of the mercurials when used in
complex cellular preparations.
In addition to these problems, it is likely that many of the most notable
* If a reader wishes to obtain information on the inhibition of an enzyme not given
in the table, I shall be happy to try to supply this upon receiving a written request.
830
7. MERCURIALS
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INHIBITION OF ENZYMES 831
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INHIBITION OF ENZYMES 833
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inhibitions constitute mutual depletion systems and are in zones B or C
(see page 1-66). The inhibition is independent of K^ in zone C — i.e., i =
(I;)/(E^) — and so the degree of inhibition indicates, not the affinity of the
enzyme for the mercurial, but only the concentration of enzyme (or of other
substances binding the mercurials). When we see in the table that 0.001 raM
p-MB inhibits an enzyme 50%, does this mean that K^ is approximately
0.001 vciM, or that the enzyme concentration is 0.002 mM (assuming pure
enzyme)? It is actually more likely to be the latter, and this is one reason
why the values in the table should not be taken too seriously. These ques-
tions will be considered in the following section.
It is rather difficult to find many enzymes which are insensitive to the
mercurials. The following enzymes might be considered as relatively resis-
tant ( < 10% inhibition at 1 mM or above): adenylate kinase, most alkaline
phosphatases, a-amylase, potato apyrase, cellobiase, coagulases, copropor-
phyrinogen oxidase, dinitrophenol reductase, elastase, /5-glycerophospha-
tase, glycolate oxidase, kynurenine formamidase, certain lipases, maltase,
neuraminidase, nitrite reductase, oxalate decarboxylase, pepsin, peroxid-
ases, phospholipases, many proteases and peptidases (especially bacterial,
fungal, and venom), some pyrophosphatases, most RNases and DNases,
certain sulfatases, thiamine diphosphatase, and uricase. These certainly do
not constitute as a whole a very extensive or particularly important group
of enzymes. Actually, the majority of enzymes are inhibited in an inter-
mediate fashion between these and the examples in Table 7-13, i.e., 50-
100% by concentrations in the range 0.05-1 mM, although some of these
would undoubtedly exhibit a much greater sensitivity if examined under
appropriate conditions (in a pure form, at physiological temperatures and
pH, and in the absence of high concentrations of ligands).*
Comparison of Mercurials
Since the introduction of p-MB some 30 years ago, it and the related p-
MPS have been used for enzyme inhibition more and more frequently at
the expense of Hg++. It is interesting, therefore, to look into the results
which have been obtained with the inorganic and organic mercurials. Of
the total of 160 reports on enzyme inhibition using both Hg++ and p-MB,
25% do not allow an accurate comparison, due mainly to different concen-
* A word should perhaps be said against the common practice of reporting only
an inhibition of 100% with a single mercurial concentration, particularly if this is
relatively high, since such results are not very meaningful and the enzymes cannot
be accurately classified as to sensitivity. For example, to state that 1 mM p-MB
inhibits 100% is bad on two counts: One does not know the true sensitivity of the
enzyme, since 0.01 mil/ might also inhibit 100%, and such a result does not provide
much evidence for the importance of SH groups at the active center, although it
is often so interpreted.
INHIBITION OF ENZYMES 861
trations being used, or statements that both at certain concentrations in-
hibit 100%. Of the remaining reports, Hg++ is more potent in 65%, p-MB
in 29%, and in 6% they are of equal potency. One must admit that Hg++
is generally more effective. In some cases it is of much greater inhibitory
potency than p-MB or the other organic mercurials. One might expect Hg++
to be more potent than p-MB because (1) it is smaller and might be able
to penetrate and react with SH groups inaccessible to the larger molecule,
and (2) it could possibly in some instances induce dimerization of the en-
zyme (or even polymerization), since it is bifunctional. On the other hand,
p-MB might be considered to shield off, sterically or electrostatically, a
greater area on the enzyme surface once it has combined with the SH groups,
due to its greater size and the charged COO" group. The result in any case
is probably a balance of these and other factors. The fact that Hg++ is
often more inhibitory than p-MB does not immediately imply that it is a
better or more reliable inhibitor to use for the purpose of detecting SH
groups on enzymes, but it does, I think, suggest that Hg++ has been un-
necessarily neglected by many workers. It might be proposed that both
mercurials be used, since not only will the detection of SH groups be made
more certain, but occasionally interesting information on the nature of the
inhibition may be obtained.
The organic mercurials themselves have not often been used in the same
investigation, but in 19 reports using both p-MB and PM, I have found PM
to be more potent in 79%. The differences between them are seldom very
marked, however. One would not expect much difference between p-MB
and 2?-MPS, and examination of the eight reports using both bears this out,
in two p-MB being the more potent, in two p-MPS being the more potent,
and in four the potencies being the same. There is some reason for believ-
ing that the smaller uncharged alkyl mercurials, such as MM, might be
better for enzyme study than any of the other mercurials, but there has
been so little comparison that nothing can be said definitely about their
relative effectiveness at this time.
Meaning of K,- and Methods of Expressing Inhibition by the Mercurials
The values for Kj have occasionally been reported for mercurial inhibi-
tion; e.g., for the inhibition of phosphoribosyl-ATP pyrophosphorylase by
p-MB, pZ, = 5.15 (Martin, 1963), and for the inhibition of heart lactate
dehydrogenase by p-MB, pK^ = 4.10 (Gruber et al., 1962). In the latter case
the binding of the mercurial to the noncatalytic SH groups, causing the
spontaneous reversal of the inhibition, is characterized by a p^j of 5.40.
Most of the values given for p^, are in the range 4-6. However, these values
were obtained by simply taking the concentration to produce 50% inhibi-
tion, which would be valid if (1) the inhibition is classically noncompetitive
(which in most cases probably it is not), and (2) the system is in zone A
862 7. MERCURIALS
(which it seldom is). If the system is in zone B or C, K^ may be a good
deal smaller than PI50, and we have seen that in zone C there is no way
kinetically of determining K^. The only valid calculation of a true dissocia-
tion constant for a mercurial complex with an enzyme, of which I am aware,
is that of Madsen and Gurd (1956) for muscle phosphorylase and p-MB.
They used an ultracentrifugal method to measure the concentration of free
p-MB after equilibration and determined K^ from a plot of 1/r against
r/(p-MBy), where r is the molar ratio of p-MB bound to protein and p-MBy
is the free mercurial. A value of p^, = 6 was found. It is likely that in
most cases in which an enzyme is potently inhibited by a mercurial, a p^,
of 6 or less would be found, and in this range it is very difficult to deter-
mine the constant by the usual plotting procedures. It will be recalled that
W. L. Hughes (1950) found a p^ of 4.46 for the complex of mercaptalbumin
and MM (page 759). We may now inquire into what values of K^ would be
predicted under ordinary circumstances. Equation 7-3 gives the relation
between the experimental constant and the dissociation constant for a mer-
curial complex, and if we alter it to correspond to inhibition of enzymes
we have
p^/ = vKi — pKa - p-fi'x ,
where ipK/ is the experimental or apparent dissociation constant. If p^, is
taken as 21, \)K„ as 8.7, and ipK^ (for Cl~) as 6.5, all of these being approxi-
mations, Y>K/ turns out to be around 5.5. Since pK^ certainly varies from
20 to 22, pK^ from 7.5 to 9.5, and p^^, (depending on the ligand) from 6 to
9, it is clear that pK/ may vary over a wide range, but at least the values
experimentally determined are of the correct order of magnitude. The ex-
perimental i)K/ is thus a good deal less than the true p^, because of the
competitive effects of H+ and the ligand X~.
The question of how best to report mercurial inhibitions is a difficult
one. First, the concentration-inhibition curves are often very steep (Fig.
7-31) so that giving the results of a single concentration may be quite mis-
leading. Therefore one can suggest that in all studies a range of mercurial
concentration be used, such as to provide different degrees of inhibition,
preferably from 0 to 100%. Second, values of K^, which are so useful in
other inhibitions, are difficult if not impossible to determine by the usual
procedures, especially when the systems are in zones B or C, in which case
plgo may vary greatly depending on the enzyme concentration. It is evident
that for work with pure enzymes it is best to state the amount of inhibitor
present in terms of //moles per milligram of enzyme, or if the molecular
weight of the enzyme is known to express this as a molar ratio. However,
when impurities are present, and especially when preparations such as ho-
mogenates are used, this method is not as useful and even a designation
such as //moles of mercurial per milligram of total protein is not very mean-
ingful. Third, as we have discussed previously, mercurial inhibition is
INHIBITION OF ENZYMES
863
strongly dependent on several factors, such as pH, temperature, and me-
dium composition, so that these conditions should be stated accurately
and completely, and it should always be realized that the results reported
apply only to these particular conditions.
Fig. 7-31. Inhibitions of glutamate dehydro-
genase, showing the relative potencies of the
various inhibitors. Glutamate =11 mM, NAD
= 0.17 mM, and pH 7.6. (From Olson and
Anfinsen, 1953.)
Inhibition of ATPase
The results of the actions of the mercurials on ATPase were not included
in Table 7-13 because they are complex and warrant more detailed treat-
ment, particularly as the effects of mercurials on mitochondrial and myosin
ATPase are of importance in the work on oxidative phosphorylation and
muscle contraction, respectively. Some of the reported inhibitions of ATPase
are shown in Table 7-14; the stimulation of ATPase under certain conditions
has been omitted since it was presented in Table 7-11. One immediately
notes a very great variation in results. This is due partly to the different
sources of the enzyme, but also to the different conditions under which
the experiments were run. The response to mercurials depends on the state
of activation of the enzyme, whether Ca++ or Mg++ is present, the pH, and
the temperature, as well as the obvious factors of buffers and nonenzymic
protein. The pH actually determines whether stimulation or inhibition will
864
7. MERCURIALS
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INHIBITION OF ENZYMES 865
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INHIBITION OF ENZYMES
867
be exhibited over a wide range of mercurial concentration (Fig. 7-32). The
fairly sjonmetrical curves for myosin ATPase, the maximal stimulation
being observed at a pH around 7.5, are most likely the result of changes in
ionizable groups at or near the active center, whereas the more complex
curve for mitochondrial ATPase perhaps arises from additional factors relat-
ed to mitochondrial structure or the effects of intramitochondrial compo-
nents on ATPase. It may also be mentioned that Tonomura and Furuya
(1960) found essentially the same behavior for myosin B ATPase, stimul-
ation being maximal at pH 7.8 and absent at 5.7 and 10.
+ 250-
•200
+ 100
+ 50
-100
MITOCHONDRIA
8
Fig. 7-32. Effects of pH on the actions of p-MB on
ATPase. Liver mitochondrial ATPase treated with p-MB
at 0.1 mM (Myers and Slater, 1957 b). Myosin ATPase
curve # 1 treated with p-MB at 0.04 /<mole/mg (Stracher
and Chan, 1961), and curve #2 treated with p-MB at
0.0872 /<mole/mg (Blum, 1962 a).
When ATP is added to a preparation of myosin ATPase, there is an initial
burst of phosphate release, followed by a steady level of hydrolysis. The ef-
fects of p-MB on these two phases of activity have been shown to be quite
different by Tonomura and Kitagawa (1957, 1960). There is a progressive
depression of the magnitude of the initial burst as the SH groups are ti-
trated, but the steady rate is accelerated until around 80% of the groups
have been combined (Fig. 7-33). The rate of the initial burst may be stim-
ulated but the amount of ATP split during this period is reduced. How
these observations correlate with the various theories of how mercurials
activate ATPase (page 819) is not known; for example, does the initial
868
7. MERCURIALS
burst have anything to do with the postulated structural rearrangements
of the enzyme following addition of ATP ? In any event, there is evidence
that mercurials can alter not only actomyosin stability itself (Gergely et al.,
1959) but the structure (Kominz, 1961) or flexibility (Levy et al, 1962) of
the enzyme near the active center.
The configurational changes are apparently complex inasmuch as meas-
urements of the optical rotatory dispersion indicate that p-MB up to 4
moles/ 10^ g myosin A increases the helical content several per cent, but
addition of 8 moles/10^ g decreases the hehcal content (Tonomura et al.,
1963 a). It was suggested that these small changes in the helical structure
200
150-
100
- 50
UJ <
tr >
^
\ STEADY
/
/
\ RATE
\
\
^/titration
\
\ ^
^ OF SH GROUPS
N. INITIAL
\^^
\burst
y^
/>
\V
0 0 02 0.04
p-MB (/iMOLES/MG)
006
0.08
0.10
Fig. 7-33. Titration of myosin B ATPase with p-MB and
the effects on the initial and steady-state rates at pH 6.7
and 20°. The SH reaction measured by absorption at 255
m/<. (From Tonomura and Kitagawa, 1960.)
induce modifications at the active site. If the inactivated enzyme is treated
with /5-mercaptoethanol the mercurial is removed, the activity is restored,
and the rotatory dispersion returns to normal (Tonomura et al., 1963 b).
One might then assume that the effects of the p-MB are completely rever-
sible; however, it was found that the substrate inhibition at high concentra-
tions of ATP no longer occurs, and that EDTA now does not inhibit. It
may be that the treatment with ^j-MB removes divalent cations, possibly
the tightly bound Ca++. It was indeed shown that the progressive depression
of the ATPase activity is accompanied by a loss of the binding of Ca++ and
Mg++ (Martonosi and Meyer, 1964).
INHIBITION OF ENZYMES 869
The interesting treatment of the kinetics of myosin ATPase inhibition
with p-MB recently reported by Walter (1963) should be consulted for the
theoretical development of the equations and appropriate plotting proce-
dures, but in connection with our present subject it is worth noting that
the kinetics are complicated by two factors which should be kept in mind
in such work on all enzymes. First, the reaction of the first SH groups
does not lead to inactivation immediately. Second, this initial reaction re-
duces the mercurial concentration so that the more slowly reacting groups
are exposed to a much lower concentration than was originally added. The
calculated bimolecular rate constant for these catalytically important SH
groups is thus quite different than that obtained from the initial rate.
The relationship between the effects of mercurials and 2,4-dinitrophenol
is very interesting and the results indicate that SH groups are involved in
the stimulation of ATPase activity by the latter substance. Lardy and
Wellman (1953) noted that 0.04 mM p-MB almost completely abolishes
the activation of mitochondrial ATP splitting by DNP, and others not
only have confirmed this in general but have shown that the DNP-activated
enzyme is only inhibited by mercurials (Greville and Needham, 1955; Gil-
mour and Griffiths, 1957; Pullman et al., 1960). This is shown for myosin
ATPase in Fig. 7-30. However, many factors can influence these interac-
tions. Myers and Slater (1957 b) found that p-MB inhibits the DNP-acti-
vated mitochondrial ATPase from pH 6 to 8, but stimulates further at pH 9,
and Cooper (1958 b) has emphasized the importance of Mg++, addition of
this ion decreasing the inhibition by p-MB of DNP-activated enzyme, as
originally shown by Lardy and Wellman. Not all ATPases may behave in
this fashion, and the activity of a particulate preparation from Rhodospi-
rillum rubrum can still be stimulated by DNP in the presence of p-MB and
Mg++ (Cooper, 1958 a). Some investigators have postulated that DNP and
the mercurials activate ATPase by similar mechanisms, but it seems doubt-
ful if the evidence is sufficient to draw this conclusion. It has recently been
found that COg stimulates mitochondrial ATPase markedly and this occurs
in the presence of 0.005 mM Hg ++,which itself has broght about activation;
indeed, Hg++ activates about the same in the absence or presence of COg
(Fanestil et al., 1963). These stimulations thus appear to be approximately
additive. The K+-Na+-activated membrane ATPase contains SH groups
which seem to be specially involved in this activation and react readily
with mercurials (Skou, 1963).
The interesting effects of the mercurials on the P,-ATP and ADP-ATP
exchange reactions occurring in mitochondria and the relations to ATPase
will be discussed later under oxidative phosphorylation (page 872).
870 7. MERCURIALS
ELECTRON TRANSPORT
AND OXIDATIVE PHOSPHORYLATION
The majority of dehydrogenases are quite sensitive to the mercurials,
and inspection of Table 7-13 shows that 50% inhibition is commonly pro-
duced by concentrations of 0.001-0.05 mM. If we define NADH dehydro-
genase as the enzyme component responsible for the transfer of electrons
to a variety of acceptors, it must be placed in the same category with
respect to sensitivity, and, indeed, NADH-cytochrome c reductase is usual-
ly even more susceptible, often being completely inhibited by concentra-
tions of 0.001-0.01 mM. One is thus tempted to attribute the inhibition
of various oxidations by the mercurials to an action early in the electron
transport chain, at least pre-cytochrome. Furthermore, Barron and Singer
(1945) had reported that the oxidation of reduced cytochrome c by a cyto-
chrome oxidase preparation is not affected by p-MB, and this has more
recently been observed with Arum (Simon, 1957) and Penicillium (Sih et
al., 1958) cytochrome oxidases. Finally, some have found that certain oxi-
dases and the corresponding dehydrogenases are inhibited equally by mer-
curials, although the different conditions of testing in such cases make ac-
curate comparison difficult.
This simple picture of inhibition in the electron transport sequence has,
however, been questioned by workers at the Institutum Divi Thomae, who
from 1946 to 1957 obtained increasing evidence that the cytochrome system
may not be as immune to mercurials as generally imagined. Their results
may be summarized in the four following categories. (1 ) Cytochrome oxidase
activity, as determined by the oxidation of ascorbate, is inhibited rather
potently, 50% depression being observed with 0.006-0.012 mM PM and
0.032 mM p-MB (Cook et al, 1946; Kreke et al, 1950). (2) Succinate oxi-
dase is much more sensitive to mercurials than is succinate dehydrogenase.
It requires around 10 times the concentration to inhibit rat heart succinate
dehydrogenase compared to the oxidase (Cook et al, 1946; Kreke et al,
1949; Smalt et al, 1957). (3) The inhibitions of cytochrome oxidase and
succinate oxidase are not reversed by thiols (Cook and Perisutti, 1947;
Kreke et al, 1949, 1950). This led them to suppose that the inhibition might
not involve SH groups, but this conclusion, as we have seen, is not valid.
(4) No evidence for reaction of the mercurials with ascorbate or cytochrome
c could be obtained by spectroscopic or preincubation techniques (Cook et
al, 1946; Kreke et al, 1950). It may also be mentioned that Boeri and Tosi
(1954) found no reaction of p-MB with cytochrome c, and that Strittmatter
and Velick (1956) likewise found no change in microsomal cytochrome spec-
tral absorption after incubation with 1 mM p-MB. All of these data have
been interpreted as indicating that the mercurials may exert a major part
of their effect on cytochrome oxidase.
ELECTKON TRANSPORT 871
Slater (1949) had also observed that succinate oxidase is inhibited more
strongly than the dehydrogenase by p-MB (and also by o-iodosobenzoate
and oxidized glutathione), although the difference was not as great as re-
ported by Cook, Kreke, and their co-workers, and attributed this in the
particulate preparations used to an effect on some link between the dehy-
drogenase and the oxidase, presumably occurring before cytochrome c in
the chain. This effect might be a structural disorganization of the complex
to interrupt the flow of electrons. Nevertheless, Slater observed some inhibi-
tion of cytochrome oxidase. Seibert et al. (1950) made a solubilized deoxy-
cholate preparation of cytochrome oxidase and found by both manometric
and spectrophotometric tests that it is inhibited to the same degree as the
crude preparation: they also demonstrated shifts in the spectral bands of the
oxidase following treatment with the mercurials. The final conclusion of the
Institutum Divi Thomae group is that the actions of the mercurials on heme
enzymes may be nonspecific, may involve denaturation (which could ac-
count for the spectral shifts), and do not involve SH groups, but I doubt
if there is sufficient evidence for any of these statements. However, their
data, which are definite and consistent, must be explained on some basis.
It is important to realize that the inhibitions reported for "cytochrome
oxidase" were all obtained with ascorbate (and occasionally hydroquinone)
as the substrate. Now neither ascorbate nor hydroquinone is oxidized di-
rectly by cytochrome oxidase and the electron transfer occurs through a
series of components. It has usually been assumed that ascorbate reduces
cytochrome c^ or c, in which case the action of the mercurials could be on
some component or link in the cytochrome sequence, rather than on cyto-
chrome oxidase itself. It will be remembered that the work quoted at the
beginning of this section showed that, when cytochrome c is used as sub-
strate, the mercurials do not inhibit. Is it possible that there is a com-
ponent which might be designated as ascorbate dehydrogenase, which is
sensitive to the mercurials? Seibert et al. (1950) actually observed relative-
ly little inhibition of the purified system when determined spectrophoto-
metrically with cytochrome c as the substrate.
There are several ways of explaining the differential inhibitions of suc-
cinate dehydrogenase and oxidase. Since the activities of these two sys-
tems are determined very differently — the dehydrogenase usually by
methylene blue reduction and the oxidase manometrically — one must
question if this could be responsible for the different sensitivities observed.
The dehydrogenase activity associated with methylene blue reduction might
not be exactly the same as in the normal transfer of electrons to the cyto-
chromes; i.e., a region of the enzyme surface, or another component in the
chain, might be involved in the normal transfer but not in the dye reduc-
tion, and this part of the system could be sensitive to the mercurials. If
we look into the details of the procedures (Kreke et al., 1949), we find that
872 7. MERCURIALS
in the dehydrogenase test the pH was 7.2 and the succinate concentration
0.33 rciM, whereas in the oxidase test the pH was 7.4 and the succinate
88 mM; the former was done in strong phosphate buffer, whereas the latter
medium contained 0.7 mM Ca++ and A1+++; in addition, the times for
equilibration and incubation were different. When the conditions are so
diverse, it is impossible to compare these two systems quantitatively. The
structural interference theory of Slater must also be considered and has as
much evidence as the other explanations (i.e., none). It would be important
to know just how much effect mercurials can exert on the cytochrome sys-
tem, inasmuch as it has obvious bearing in considerations of the actions
on various oxidations, mitochondrial systems, and respiration.
A comparable situation with NADH dehydrogenase, NADH: cytochrome
c reductase, and NADH oxidase has been noted by Minakami et al. (1963).
The total oxidase and the cytochrome c reductase are very sensitive to p-MB
whereas the dehydrogenase, as determined by ferricyanide reduction, is not
as sensitive. It was postulated that two types of SH group are involved in
NADH oxidation, one readily accessible to mercurials and functioning be-
tween the dehydrogenase active site and the distal respiratory chain (this
SH group is not required for ferricyanide reduction), and a second con-
cealed in the dehydrogenase complex as isolated, and exposed on degrada-
tion to the cytochrome c reductase. Such an explanation could apply to the
succinate oxidase as well, as was suggested above relative to methylene
blue as an acceptor for the determination of dehydrogenase activity.
Oxidative Phosphorylation
The results summarized in Table 7-15 show that mercurials are not par-
ticularly specific or effective uncouplers of oxidative phosphorylation in
mitochondria, but that a fair degree of uncoupling can occur under certain
circumstances. It is especially interesting that high toxic doses of the mer-
curial diuretics and HgClg can often reduce the P : 0 ratio in the mitochon-
dria of excised kidneys several hours after the administration, without sim-
ultaneously affecting oxidative phosphorylation in the liver, but this is
undoubtedly due to the higher concentration of mercurial in the kidney.
The P : 0 ratio is, however, not altered significantly by the ordinary diuretic
doses, so that it is questionable if this action is related to diuresis. I know
of no instance in which the mercurials augment O2 uptake while simul-
taneously reducing the P, incorporation, so that they are not true uncou-
plers in the same sense as the nitrophenols.
The Pj^^-ATP exchange is quite potently inhibited by mercurials in the
mitochondria obtained from mosquitoes (Avi-Dor and Gonda, 1959), pig
liver (Chiga and Plant, 1959), and rat liver (Plaut, 1957; Cooper and Leh-
ninger, 1957; Lehninger et al., 1958; Low et al., 1958). For rat liver mito-
chondria the exchange is sometimes inhibited 50% by concentrations
ELECTRON TRANSPORT
873
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874 7. MERCURIALS
around 0.002 mM, and completely by 0.01 mM. The ADP^s-ATP and
ADP-C^*-ATP exchanges are also inhibited, but perhaps not as strongly
(Wadkins and Lehninger, 1958; Chiga and Plant, 1959; Kahn and Jagen-
dorf, 1961). These exchange reactions are related intimately with oxi-
dative phosphorylation. Indeed, Wadkins and Lehninger (1958) postulated
that the Pj-ATP exchange is a measure of the two terminal reactions in
oxidative phosphorylation:
Carrier — ^ X + Pi ^ carrier + P ^ — ^ X
P — X 4- ADP ±5 ATP + X
where X is perhaps the enzyme protein, while the ADP- ATP exchange
measures only the last reaction. They further suggest that the mercurials
inhibit this last reaction principally, whereas 2,4-dinitrophenol acts on the
penultimate step. If the mercurials act solely on the transfer of phosphate
to ATP they would be good uncouplers, but actions elsewhere in the elec-
tron transport chain limit their efficiency. It is interesting that Griffiths
and Chaplain (1962) have found evidence for a new phosphorylated deriva-
tive of NAD following incubation of heart mitochondria with succinate
and P(^2. ATP can be formed from the intermediate and this reaction is
completely blocked by p-MB at 0.01 mM. This observation is compatible
with the scheme of Wadkins and Lehninger.
FERMENTATION AND GLYCOLYSIS
The first impression from surveying the studies of mercurial action on
fermentation and glycolysis is that these pathways are often surprisingly
insensitive to this group of inhibitors. In many cases it requires concentra-
tions greater than 1 mM to depress glycolysis significantly in cellular sys-
tems. Reference to Table 7-13 shows that several enzymes in the Embden-
Meyerhof pathway are quite readily inhibited by mercurials, e.g., hexo-
kinase, aldolase, 3-phosphoglyceraldehyde dehydrogenase, enolase, and lac-
tate dehydrogenase, concentrations of 0.001-0.05 mM usually inhibiting
50% or more in muscle, although little is known about the sensitivities of
the yeast enzymes. Since several enzymes in the pathway are susceptible,
one might anticipate that the sequential inhibition by mercurials at con-
centrations above 0.05 mM would produce a very strong over-all depression
of anaerobic COg or lactate formation. Three explanations for the failure to
do so are immediately apparent: (1) The mercurials do not penetrate into
the cells readily; (2) the glycolytic enzymes are protected in the cell (e.g.,
by substrates or coenzymes); and (3) the large amount of nonenzymic ma-
terial in cellular preparations binds much of the mercurial. Cleland (1949)
found that 1 mM PM inhibits glycolysis in oyster eggs only 17% at 0-45
min and 48% at 45-90 min, whereas glycolysis in egg homogenate (with
FERMENTATION AND GLYCOLYSIS 875
glycogen, ATP, and NAD added) is completely blocked. Similar results
were obtained with sea urchin eggs by Cleland and Rothschild (1952 a),
1 mM PM inhibiting lactate formation 17% in whole eggs and 97% in
extracts. These results were interpreted to indicate poor penetration by the
mercurial, but the other two explanations given above are probably as
likely, and undoubtedly all contribute to some extent. It may be noted that
Cleland found endogenous respiration to be inhibited more potently in
whole eggs than in homogenates, which is more difficult to explain. There
are instances of cellular glycolysis quite sensitive to the mercurials; in asci-
tes carcinoma cells there is 50% inhibition by 0.0032 mM Hg++ (Schom
et at., 1961). The glycolysis in spleen slices is also fairly sensitive, although
it requires 4-5 hr to reach maximal inhibition (Fig. 1-12-24) (Jowett and
Brooks, 1928). The interesting questions of the penetration of mercurials
and the effects exerted on cell membranes will be considered later (page 892).
Yeast Fermentation
Since the early work of Schulz (1888), who reported an initial stimulation
of fermentation by low concentrations of Hg++ (0.005-0.008 mM) and inhi-
bition by higher concentrations (> 0.02 mM), there have been many stud-
ies of yeast fermentation with variable results. The stimulation observed
by Schulz has seldom been confirmed. Joachimoglu (1922) could never dem-
onstrate acceleration of CO2 formation by Hg++, concentrations of 0.0031-
0.037 mM exerting no effect and 0.074 mM inhibiting around 70%. Meier
(1926) found even more potent inhibition of aerobic fermentation, 0.009 mM
Hg++ depressing 72%, while Kostytschew and Berg (1930) never observed
stimulation, inhibition beginning at 0.0185 mM Hg++ and reaching 42%
at 0.2 mM. More recently, some have found potent inhibition by Hg++
(e.g., Hurwitz and Chaffee, 1954), but others have not (e.g., Weitzel and
Buddecke, 1959); in the latter work, 1 mM Hg++ inhibited only 70% in
fermenting yeast. Organic mercurials have not been often used, but Spiegel-
man et al. (1948) reported 69% inhibition by 0.01 mM PM and 32% inhi-
bition by 0.05 mM p-MB, indicating these mercurials to be fairly effective.
Certainly much of the variation in the results is due to the different densi-
ties of yeast suspension used, the media employed, and the state of the
yeast (by which is meant its fermentative activity and past history). One
would expect mercurials to attack surface hexokinase and the initial phos-
phorylation of glucose, as occurs in muscle, so one can explain the exam-
ples of weak inhibition only on the basis of relatively dense yeast suspen-
sions.
Muscle Glycolysis
The results obtained on muscle glycolysis with the mercurials have been
quite inconsistent and even more difficult to explain than those with yeast.
876 7. MERCURIALS
Gemmill and Hellerman (1937) found that Hg++, p-MB, and PM all block
glycolysis in extracts of frog muscle, but the concentrations used were too
high and usually unspecified. Separated fibers of cockroach muscle treated
with 1 mikf p-MB show no change of COg formation and a rather marked
increase in lactate formation if only glucose is added, but in the presence
of glucose + ATP, COg production is inhibited 67% and there is no effect
on lactate (Barron and Tahmisian, 1948). This behavior is quite different
from that of iodoacetate, which inhibits only in the absence of added ATP.
The authors felt that the failure to depress lactate formation in any case
is perhaps a characteristic of invertebrate muscle, since Harting (1947) had
observed 1 mM p-MB to produce only stimulation of glycolysis in strips of
scallop and thyone muscle. However, Krueger (1950) has shown that 2 min
perfusion of frog muscle with 37 mM Hg++ essentially doubles the lactate
formation. The only serious study of muscle glycolysis was done by Bailey
and Marsh (1952) on rabbit psoas homogenates. Here p-MB produces def-
inite inhibition (see accompanying tabulation), but the concentration is so
Control
p-MB 4 mM
zIpH
- 0.28
- 0.08
A Fructose-diP
+27
+ 6
A Triose-P
+ 3
- 1
zl ATP
-14
- 5
ATP resynthesis
45
13
ATPase inhibition
77%
high that it is remarkable that the inhibition is not much greater. It was
believed that aldolase inhibition is responsible for the results but from the
data it is not possible to localize the site of action so closely. The authors
pointed out that 3-phosphoglyceraldehyde dehydrogenase is not so readily
inhibited by p-MB as by iodoacetate. The transfer of phosphate from crea-
tine-P to ADP is immediately and completely blocked by p-MB, so that
creatine-P remains at its initial level, and this must also be a factor in the
inhibition, since it would prevent regeneration of ATP. It is thus impossible
in this study to determine what effects p-MB might have directly on the
Embden-Meyerhof pathway. All of the results on intact muscle tissue seem
to be incompatible with the demonstration by Demis and Rothstein (1955)
that glucose uptake by diaphragm is very sensitive to Hg++, being almost
completely inhibited by 0.2 mM. However, respiration and anaerobic lac-
tate formation, being dependent on endogenous substrate, are much less
sensitive and are only slowly inhibited. This will be considered in greater
detail when the effects of mercurials on respiration are discussed (page 884).
TRICARBOXYLATE CYCLE 877
Stimulation of glycolysis by the mercurials is not confined to yeast and
muscle. Hg++ below 0.11 mM stimulates glycolysis in guinea pig blood and
inhibits in higher concentration (Fuentes and Rubino, 1923), while in hu-
man blood Hg++ stimulates anaerobic glycolysis from 0.0185 to 1.85 vaM
although at 18.5 mM there is almost complete inhibition (Rubino and
Varela, 1923). Glucose utilization, COg release, and lactate formation in
human erythrocytes are all stimulated by p-MB up to 5 //moles/ml of
erythrocytes (Jacob and Jandl, 1962). No explanation for these results
is immediately evident.
TRICARBOXYLATE CYCLE
Despite the fact that no analysis of the effects of mercurials on the cycle
or on the operation of mitochondria has been made, one would predict
quite potent inhibition on the basis of the sensitivities of the individual
enz\Tnes (Table 7-13). Mercurial concentrations in the neighborhood of
0.01 mM should depress several enzjTnes very significantly (e.g., pjTuvate
oxidase, isocitrate dehydrogenase, or-ketoglutarate oxidase, succinate de-
hydrogenase, malate dehydrogenase, and some ancillary enzymes, such as
acetate kinase), and concentrations of the order of 0.1 mM should block
completely. However, since we have already noted that glycolysis is often
not inhibited as much as one would expect from studies of the individual
enzymes, we must be very careful in considering inhibitions of the cycle
in cellular preparations. The utilization of pyruvate and acetate by a va-
riety of cellular and subcellular preparations has been shown to be readily
inhibited by mercurials (Table 7-16), but in no case was the operation of
the entire cycle tested, so that the entire inhibition, as far as one knows,
might be on the initial enzyme reaction (p>Tuvate oxidase or acetate kinase).
If the cycle is operating by regenerating oxalacetate, much stronger inhi-
bition would undoubtedly be observed. In work with mitochondria, homo-
genates, or cell suspensions, however, one must always remember the role
of nonenzyme protein in reducing the mercurial available for inhibition,
so that concentrations such as those in Table 7-16 are not of much quanti-
tative significance, but show definite interference with cycle activity.
In order to answer some of these questions relative to the action of the
mercurials on the cycle, Dr. Yang kindly consented to examine the effects
of Hg++ on the Oj uptake of rabbit heart mitochondria by the same tech-
niques used in a previous study of iodoacetate (Yang, 1957). The changes
over a 60 min period obtained from results on three preparations are shown
in the accompanying tabulation. These data show clearly that several steps
in the cycle are inhibited rather strongly as the concentration is raised from
0.003 mM to 0.01 mM, and that at 0.1 mM the cycle activity is essentially
completely blocked. The stimulation observed with a-ketoglutarate as the
878
7. MEKCURIALS
A
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S §
RESPIRATION 879
°o Changes from Hg++ at:
0.001 mM
0.003 mM
0.01 ml/
0.1
mM
Pyruvate + malate
-10
-79
99
a-Ketoglutarate
+24
+5
-74
-
100
Succinate
0
+5
-41
—
100
Malate
- 6
—
-79
—
99
substrate seems to be real since it was consistently obtained. The inhibition
with pyruvate + malate as substrates and Hg++ at 0.01 vcvM is about 50%
at 10 min and then increases more slowly until it is 100% at 60 min. The
figures in the tabulation are mean inhibitions over the 60 min period and
even at 0.01 milf the activity was almost all lost in aU cases by 60 min.
RESPIRATION
The effects of the mercurials on the 0., uptake of tissues vary considerably
and depend on the mercurial used, the substrate, the pH, the species, and
many other factors (Table 7-17). One factor about which little is known,
but which could be very important, is the thickness of the tissue when the
preparation is a strip, section, or slice, inasmuch as the mercurial possibly
does not penetrate equally throughout but acts primarily on the outer
layers of cells. Cascarano and Zweifach (1962) examined rat diaphragm
after exposure to Hg+"^ by determining the ability of the tissue to reduce
a tetrazolium dye, and found that only a well-defined band of surface fibers
had lost the ability, the central portions retaining activity. Measurements
of respiratory inhibition in such cases do not provide true values (see page
1-479); in the extreme case the inhibition may relate only to the fraction
of the tissue affected, and progressively developing inhibition may exhibit
time relations dependent only on the rate of penetration through the tissue.
This would apply not only to respiration, of course, but to all measurements,
metabolic or functional, made on all intact tissues. Failure to reach all of
the cells equally must be one reason for the low degree of inhibition often
observed, lower than would be predicted from the effects on glycolysis,
the cycle, and the enzymes involved.
One notes several examples wherein respiration is stimulated by the mer-
curials, more often at low concentration but in one instance, yeast respir-
ing endogenously (Shacter, 1953), the stimulation appears only at high
concentrations of 1-2 vaM. There are other reports of stimulation not in-
cluded in the table. For example, Gremels (1929) found that when mersalyl
induces diuresis in a heart-lung-kidney preparation, the kidney respiration
880
7. MERCURIALS
P5
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RESPIRATION 881
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RESPIRATION 883
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884 7. MERCUEIALS
is increased. There has been no study of the mechanism whereby such stim-
ulation is produced. One might postulate that the mercurials can increase
membrane permeability so that substrates can enter cells more rapidly, but
although this may be a factor it is clear that endogenous respiration can be
stimulated, as in the work of Shacter. It is also known that subcellular
preparations, and indeed certain enzymes themselves, are stimulated (page
815), so that one cannot expect to provide a comprehensive theory based
only on cell and tissue responses. Shacter felt that the mercurials might
react with certain thiols which regulate metabolism, but despite all the
discussion of such regulators in the past, there seems to be little evidence
at present for their importance. Of the several mechanisms suggested pre-
viously (page 1-453), one is at a loss to select any that would apply partic-
ularly to the mercurials. Since mercurials have been shown to reduce the
P : 0 ratio in various isolated systems, it is possible that in the cell an un-
coupling action might increase Og uptake in a certain limited range of con-
centration, and it is also possible that the mercurials by a membrane effect
might alter ion movements and concentrations, thus secondarily bringing
about metabolic stimulation, but there is no direct evidence for either of
these mechanisms. Intracellular changes are undoubtedly so complex that
metabolic effects usually defy analysis. Consider the situation described by
Estler et al. (1960) in yeast treated with Hg++ (Fig. 7-34), the levels of all
the adenine nucleotides increasing at 0.2 mM, although Og uptake is scarcely
affected, while at higher concentrations the nucleotides change in a com-
plicated fashion and respiration is severely depressed. Unfortunately stim-
ulation was not recorded here, but it is only by thorough studies of this
type that one can hope to penetrate into the mysteries of inhibitor stim-
ulation.
Although Meier (1926) reported that aerobic fermentation in yeast is
more potently inhibited than respiration by Hg++ — at 0.009 mM the
former was inhibited 72% and the latter not at all — subsequent work on
a variety of cells has indicated no general relationship between the sensi-
tivities of glycolysis and respiration, and even in yeast Weitzel and Bud-
decke (1959) found both to be inhibited similarly, at least at high concentra-
tion (1 mM) of Hg++. The respiration of oyster eggs is inhibited more than
glycolysis by PM (Cleland, 1949), whereas in oyster spermatozoa the res-
piration is inhibited by PM when glycolysis as measured by lactate for-
mation is increased (Humphrey, 1950). The utilization of pyruvate in these
cells may be more sensitive to mercurials than the glycolytic pathway.
Certainly the inhibition of respiration does not imply a reduction in glu-
cose uptake: In diaphragm, 1 mM p-MB depresses Og uptake 15%, lowers
the glycogen content, and yet increases glucose utilization somewhat (Haft
and Mirsky, 1952). In most cases (e.g., yeast and Chlorella), glucose respira-
tion is more sensitive than endogenous respiration to mercurials, but this
RESPIRATION
885
has not been investigated sufficiently to draw valid conclusions. Of course,
in tissues such as most muscle and heart, in which endogenous substrates
are responsible for the bulk or all of the respiration for some after time ex-
cision, glucose would not be expected to have much effect on the inhibition
by mercurials. Little is known about the effects of mercurials on the pen-
tose-? pathway or other routes of glucose degradation. In crude extracts
of Pseudomonas converting gluconate-6-P to 3-phosphoglyceraldehyde and
12-
09
1000
- 800
0.6-1
^ 0.3 -
- 600
-400
0 i-o
Fig. 7-34. Effects of Hg++ on the respiration and the
levels of substances in yeast. Concentrations given as
/*moles/g dry weight. Run at pH 6.8 for 60 min. (From
Estler et al., 1960.)
pyruvate, p-MB at 1 mM inhibits completely (Kovachevich and Wood,
1955). The site of the inhibition is not clear, but it is presumably not on
the gluconate-6-P dehydrase, which was purified and found to be only
moderately sensitive to p-MB. The pentose-P pathway is operative in ex-
tracts of tobacco leaves, the oxidations being NADP specific, and p-MB
at 0.1 mM almost completely blocks the reduction of NADP by both glu-
cose-6-P and fructose- 1,6-diP (Clayton, 1959). The utilization of pentose-P
by extracts of Lactobacillus hrevis is inhibited only 20% by 0.1 milf p-MB
886 7. MERCURIALS
(Eltz and Vandemark, 1960). There are no data for comparing the relative
sensitivities of the Embden-Meyerhof and pentose-P pathways.
As appears to have been the case with most of the workers who have
examined the effects of the mercurials on respiration, I find little to say
that seems worthwhile. The most interesting aspects of respiratory inhibi-
tion probably pertain to the metabolic basis of certain cellidar functions,
e.g., gastric secretion (page 914) or renal transport (page 917). The site of
action to inhibit respiration is unknown and multiple sites are likely. We
know little of the penetration of mercurials into cells and the intracellular
concentrations attained, and the information is lacking to evaluate the im-
portance of nonenzymic effects. These gaps in our knowledge apply not
only to respiration but essentially to all cellular activities. One can at least
state with fair certainty that the mercurials do not act like other SH re-
agents, such as iodoacetate or the arsenicals, i.e., their pattern of inhibition
is quite different.
VARIOUS METABOLIC PATHWAYS
In this section we shall consider briefly some of the important types of
metabolism which are readily inhibited by the mercurials. Only the more
interesting aspects and interpretable investigations will be mentioned. The
effects of the mercurials on metabolism are complex and vague in all cases,
so it is essential to emphasize those studies in which clear-cut results have
been obtained, even though the work is limited to only a certain phase of
the over-all pathway and the exact site or mechanism of action is unknown.
The few systems discussed will at least point out clearly the manifold potent
inhibitions which can be exerted by the mercurials and will serve to establish
the fact that specific effects on metabolism can seldom, if ever, be achieved
in cellular systems. Perhaps with the increasing knowledge of the detailed
actions of the mercurials, there will arise situations in which selective blocks
can be produced under controlled conditions, but at the present time there
is not much reason for optimism.
Lipid Synthesis
The long sequence of reactions in the biosynthesis of sterols seems to be
strongly inhibited by mercurials at different sites. The total incorporation
of mevalonate-C^* by Lactobacillus casei over 4 hr is inhibited 59% by 0.1
mif p-MB and 96% by 1 mM (Thorne and Kodicek, 1962). The conversion
of farnesyl-PP and mevalonate to squalene by various fractions of rat liver
is depressed 50% by p-MB, p-MPS, and Hg++ at concentrations near 0.05
mM, and essentially completely by concentrations much above 0.1 mM
(Popjak et al., 1958; Anderson et al., 1960; Goodman and Popjak, 1969).
The further conversion of squalene to sterols is 97% blocked by 0.33 mM
VARIOUS METABOLIC PATHWAYS 887
p-MB (Goodman, 1961). The sensitive enzymes are probably aU located in
the microsomes. One of the enzymes on the pathway from mevalonate to
farnesyl-PP, the isopentenyl-PP isomerase, is inhibited completely by 0.1
vnM p-MB (Agranoff et al., 1960), so this could account for part of the block
in squalene formation, but there are undoubtedly other sensitive steps.
Fatty acid biosynthesis from acetate in mammary gland homogenates is
inhibited 95% by 0.1 mM Hg++ (Popjak and Tietz, 1955), and from acetyl-
CoA and malonyl-CoA in purified fractions from pigeon liver 95% by 0.075
mM p-MB (Bressler and Wakil, 1962). The inhibition is probably early in
the sequence, since various acyl-CoA's protect, but it is not on the NADPH:
acetoacetyl-CoA oxidoreductase. The incorporation of acetate- 1-C^^ into
lipid by chloroplast suspensions is also strongly depressed: 22% by 0.001 mM
p-MB, 50% by 0.01 mM, and 88% by 0.1 mM (Mudd and McManus, 1964).
Fatty acid oxidation is potently inhibited by the mercurials, and one likely
site is the initial activation by ATP (with or without Co A), catalyzed by
fatty acid thiokinase, since this is completely inhibited by 0.1 mM p-MB
(Jencks and Lipmann, 1957). The incorporation of P,^^ into mycobacterial
phospholipids is not depressed so readily, 1 mM p-MB inhibiting only 24%
(Tanaka, 1960), although the synthesis of phospholipid in rat liver mito-
chondria from of-glycerophosphate is completely blocked at this concentra-
tion (Wojtczak et al., 1963).
The direct actions of the mercurials on lipid biosynthesis combined with
other actions which would secondarily inhibit these pathways, e.g., the re-
actions with coenzyme A or the depletion of available ATP, must lead to
serious interference in the formation of fatty acids and sterols in proliferat-
ing microorganisms and contribute to the suppression of growth, and it is
interesting to speculate whether they play a role in chronic mercurial poi-
soning in animals.
Protein Synthesis
In the preparations which have been examined it appears that protein
synthesis is not particularly sensitive to the mercurials. The incorporation
of leucine-C^* into chloroplast protein is inhibited only 30% by 5 mM mer-
salyl (Stephenson et al., 1956) and into reticulocyte protein only 9% by
0.1 mM p-MB, although 1 mM inhibits almost completely (Borsook et al.,
1957), while the incorporation of phenylalanine-C^* into rat liver soluble
proteins is inhibited 85% by 1 mM p-MB (Haining et al., 1960), of amines
into guinea pig liver soluble proteins 100% by 1 mM p-MB (Clarke et al.,
1959), and of amino acids into the acid-soluble proteins of frog egg super-
natant fractions 100% by 0.77 mM p-MB (Burr and Finamore, 1963).
Although these results do not conclusively indicate the exact sensitivity of
protein synthesis to the mercurials, one is somewhat surprised to find that
such high concentrations apparently must be used to inhibit effectively.
The only instance of potent inhibition of which I am aware is that found in
888 7. MERCURIALS
Pseudomonas aeruginosa by DeTurk and Bernheim (1960), the induction
of enzymes for the oxidation of putrescine, benzoate, and }^-aminobutyrate
being reduced 50% by 0.0028 mM p-MB. The enzymes themselves are not
inhibited at this concentration. Partial protection by Fe++ when it is added
with the mercurial or shortly after was observed. It is now known that
enzyme induction is not a valid system for estimating the effects of in-
hibitors on protein synthesis in general, because there are many other fac-
tors involved. In the inhibition cited, it was in fact postulated that some
transport process in the membrane requires Fe++ and that this is the site
of attack by the mercurial.
Porphyrin Synthesis
The formation of porphyrins from glycine and a-ketoglutarate by Rhodo-
pseudomonas spheroides is completely blocked by 0.04 mM p-MB, and from
aminolevulinate by 0.1 mM (possibly by lower concentrations since they
were not tested) (Lascelles, 1956). The formation of aminolevulinate from
glycine, phosphoenolpyruvate, and succinyl-CoA is completely prevented
by 0.44 mM p-MB (Gibson et al., 1958). It would thus appear that steps
both pre- and post-aminolevulinate are vulnerable. The report of Granick
(1958) that 1 mM p-MB does not interfere with protoporphyrin synthesis
from glycine and or-ketoglutarate in chicken erythrocytes is surprising, but
may be attributed to the high density of the cell suspension (around 45%
by volume) and the consequent binding of the mercurial to nonenzyme
proteins. The condensation of porphobilinogen to uroporphyrinogen is al-
most totally blocked by 0.02 mM Hg++ and 0.1 mM p-MB (Lockwood
and Benson, 1960), and the subsequent conversion of uroporphyrinogen to
coproporphyrinogen is again essentially blocked by 0.012 mM Hg++ and
0.7 mM p-MB (Mauzerall and Granick, 1958), if the results on the isolated
enzymes catalyzing these reactions can be applied to cellular preparations.
The incorporation of Fe++ into protoporphyrin to form heme is not so sen-
sitive, in chicken erythrocyte hemolyzate being inhibited 64% and 58%
by 1 mM Hg++ and p-MB, respectively (Kagawa et al., 1959). The purified
chelating enzyme from rat liver is inhibited 75% by 0.1 mM Hg++ (Labbe
and Hubbard, 1961), the greater effect probably being due to the relative
purity of the preparation. Again one can speculate that a depression of
porphyrin synthesis may be of some significance in growth studies or
chronic poisoning.
Biolumlnescence
One of the very few thorough, quantitative, and interesting investiga-
tions on mercurial inhibition was made by Houck (1942), who studied the
effects of Hg++ on the luminescence of Achromobacter fischeri. The standard
conditions were as follows: pH 7.3, temperature 25°, 25 mM glucose as
VARIOUS METABOLIC PATHWAYS 889
substrate, and a suspension density of 4 X 10^ cells/ml. Both respiration
and luminescence of these cells are inhibited potently by Hg++, the latter
being somewhat more sensitive (Fig. 7-35). The rate of inhibition is much
more rapid than with most cellular activities, half maximal inhibition being
reached in about 30-40 sec (Fig. 7-36). It is not immediately evident why
the inhibition is more potent in the rate experiments than in the studies
on the effect of concentration. The effects of cell density on the inhibitions
are very striking (Fig. 7-37). The initial suspension here contained 6 X 10^
cells/ml and this was diluted as indicated in the graph. At 0.001 niM no
lOOi
LUMINESCENCE
0 015
0.02
0 025
0.03
Fig. 7-35. Effects of Hg++ on the respiration and luminescence of
Achromobacter. (From Houck, 1942.)
inhibition is observed until sufficient dilution is made and at low cell den-
sities the inhibition is complete. These curves illustrate very well what es-
sentially must occur in all cell or tissue preparations, whatever activity is
measured. The meaninglessness of statements that such and such a con-
centration of mercurial produces a certain degree of inhibition of some cel-
lular process is all too clear; in this case with 0.001 mM Hg++, one might
observe any inhibition from 0 to 100% depending on the cell density chosen.
Inhibition was determined with 0.001 mM Hg++ at three values of the pH,
and was greatest at 5.3 and 8.4, and least at 7.3 (one can estimate the mean
per cent inhibitions at 1 min to be 91%, 60%, and 86% at pH 5.3, 7.3,
and 8.4, respectively). The light intensity is much greater at pH 7.3 and
this may possibly be related to the rate of glucose uptake. The effects of
temperature have already been illustrated (Fig. 1-15-9) and discussed (page
890
7. MERCURIALS
250
SEC
Fig. 7-36. Effects of Hg++ at different concentrations on lumines-
cence of Achromohacter . (From Houck, 1942.)
LUMINESCENCE
RESPIRATION
1/4 1/9 1/16 1/32 1/64
DILUTION OF CELL SUSPENSION
Fig. 7-37. Effects of dilution of the Achromobacter sus-
pension on the inhibitions of respiration and lumines-
cence by Hg++. (From Houck, 1942.)
VARIOUS METABOLIC PATHWAYS 891
1-786). The increase of the inhibition with rise of temperature was inter-
preted by Houck in terms of an equilibrium between active and denatured
forms of the attacked enzyme, especially luciferase. Actually from this
work one cannot locate the site of the inhibition, or even be certain it is
on the bioluminescent reactions themselves, since interference with glucose
uptake or oxidation, or the supply of ATP, could be responsible. However,
it has been found that Achromohacter luciferase is markedly inhibited in the
range of Hg++ concentrations found to inhibit luminescence (Table 7-13),
so it may well be that luciferase is the major site of action. This is somewhat
substantiated by the fact that luminescence in extracts of Renilla reniforniis,
the sea pansy, is strongly inhibited by p-MB (Cormier, 1960).
Photosynthesis and Photophosphorylation
The marked inhibition of certain phases of photosynthesis by iodoacetate
and iodoacetamide (III-1-156) indicates the necessity of SH groups, so that
one would expect the mercurials to be effective inhibitors, and this is borne
out. The photoreduction of various dyes in isolated chloroplasts or grana
(Hill reaction) is very sensitive. In spinach chloroplasts it is inhibited 90%
by 0.005 mi/ Hg++ (Macdowall, 1949). The dye reduction may be me-
diated through NADPH, which is the initial acceptor. The photosjmthetic
NADP reductase from spinach is inhibited 50% by 0.012 mM and 90%by
0.016 mM p-MB (San Pietro and Lang, 1958) and the photoreduction of
NADP in chloroplasts is similarly inhibited, although slightly less potently
(J. S. C. Wessels, 1959). The photoreduction of cytochrome c and NADP
by the chloroplast enzyme is 50% reduced by 0.004 mM p-MPS and the
enzjTne is bleached by the mercurial (Keister and San Pietro, 1963). In
Chromatium, illumination causes a blue fluorescence presumably due to
bound NADH, indicating that here there is a photoreduction of NAD.
This fluorescence change is completely abolished by 0.02 mM PM (Olson
et al, 1959). Finally, a NADPH diaphorase from chloroplasts, possibly in-
volved in the reduction of the Hill dyes by NADPH, is inhibited 53% by
0.023 mM Hg++ and 50% by 0.13 mM p-MB (Avron and Jagendorf, 1956).
The initial photoreductive changes upon illumination are thus quite po-
tently inhibited by the mercurials, and this must certainly be one site of
action on over-aU photosynthesis. Other evidence for a primary interference
with the photolysis of water was obtained by Damaschke and Liibke (1958),
who showed that Chlorella under anaerobic conditions produces a sudden
burst of Hg upon illumination and that this is completely inhibited by 0.2
mM p-MB (lower concentrations not tested), and by Whittingham (1956),
who found that althoagh 0.12 mM p-MB does not inhibit the initial evo-
lution of Oo by illuminated Chlorella, the steady-state formation of Og is
strongly depressed. It may be mentioned that even high concentrations of
Hg++ do not react with chlorophyll (Macdowall, 1949).
892 7. MERCURIALS
Photophosphorylation to form ATP is not necessarily coupled with NADP
reduction (J. S. C. Wessels, 1959), but nevertheless one might predict that
it would be reduced by mercurials. It has been found that the incorporation
of Pj into ATP in illuminated chlorophasts is inhibited around 50% by
p-MB at 0.05-0.1 mM (Arnon et al, 1956; J. S. C. Wessels, 1958, 1959;
Jagendorf and Avron, 1959), and similar effects were reported for Rhodo-
spirillum rubrum (Smith and Baltscheffsky, 1959). Photophosphorylation
is not inhibited as potently as photoreduction.
The photochemical fixation of C^^Og by chloroplasts is inhibited 14% and
88% by 0.01 and 0.05 mM p-MB, respectively (Gibbs and Calo, 1959 b),
but a reconstructed system (extract + chloroplast fragments) is more sen-
sitive, 61% and 94% inhibition being exerted by these concentrations of
p-MB (Gibbs and Calo, 1960 b). It is not known if this implies some barrier
to penetration in the intact chloroplast. Both the initial and steady-state
rates of fixation of CO2 in illuminated dahlia leaves are only slightly re-
duced (15-25%) by 0.5 mM p-MB, even though plenty of time is provided
for penetration (Massini, 1957), and in Scenedesmus obliquus photosynthesis
is inhibited only 50% by 1 mM p-MB after 260 min exposure (Horwitz,
1957). The failure to inhibit more potently in these cases can at present
be explained only on the basis of inadequate penetration into the cells or a
certain structural integrity of the photosynthetic apparatus which makes
it difficult for a mercurial to exert such inhibition as is observed with isolat-
ed chloroplasts. No detailed study of the effects of mercurials on the rapidly
labeled C compounds has been made, but Miyachi (1960) has found that
p-MB decreases the level of what he calls the primary photogenic agent
(measured by 3-sec C^^Og fixation) in ChloreUa, although it does not inter-
fere with the participation of this substance in the subsequent photosyn-
thetic pathway. Nonphotosynthetic C^^Og fixation is usually inhibited
strongly by mercurials, e.g., the autotrophic fixation by Hydrogenomonas
facilis (McFadden and Atkinson, 1957) or the fixation associated with sul-
fide oxidation in Thiobacillus thiooxidans (Iwatsuka et al., 1962), both being
inhibited around 50% by 0.01 mM p-MB — which is not surprising con-
sidering the sensitivity of the various enzymes usually involved in COg
fixation. This dark fixation is possibly related in some manner to photo-
synthesis, and it has occasionally been pointed out that the same inhibitors
are effective in both.
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION
In the following section we shall discuss the effects of the mercurials on
permeability and membrane transport systems, as a background for under-
standing the responses of tissues to these inhibitors, but it may serve to
clarify the problem if we take up the theory of the role of the cell membrane
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION 893
in heavy metal ion inhibitions as an introduction. Although many workers
have considered the effects of heavy metal ions on membranes, the concepts
presented here will be mainly those of Rothstein and his group at Rochester,
since they have been actively engaged for over 10 years in studying this
problem. Although much of the evidence is based on work with copper,
molybdate, and uranyl ions, and the theory is meant to apply to heavy
metal ion action in general, Hg++ has been used frequently and it is im-
possible to discuss the effects of the mercurials on cells and tissue without
considering this aspect of their actions. The basic concepts of Rothstein
(1959) may be summarized as follows. (1) The cell membrane is exposed
directly to the heavy metal ions in the medium and is that part of the cell
which reacts initially when the heavy metal ions are added. Ligand groups
at the surface or within the membrane will combine with the heavy metal
ions as they diffuse, and hence the membrane will experience the first ef-
fects, and certain changes in cellular metabolism or function may at this
time relate to a selective membrane action. (2) The cell membrane usually
presents a barrier to the penetration of the heavy metal ion into the cell
and thus protects the cytoplasmic enzymes. (3) Nonenzymic or nonfuc-
tional ligand groups in the membrane or within the cell combine with the
heavy metal ions and thereby protect the active sites by reducing the
amount of heavy metal ion which is free to react. (4) As a result of the
second and third postulates, enzymes within cells are less readily attacked
by metal ions than when they are isolated from the cells. (5) As a result
of the first statement and the fourth deduction, it would be likely that the
major site of heavy metal ion action on cells and tissues is often the cell
membrane, rather than the enzyme and metabolic systems within the cell.
(6) The most important active sites in the membrane are enzymes or other
components involved in the transport of substances across the membrane.
Much of the toxicity would therefore be due to interference with the move-
ments of substrates or ions into or out of the cell.
Glucose Uptake and Respiration of Diaphragm Muscle
The uptake of glucose by rat diaphragm is almost completely abolished
within 20-30 min by 0.2 milf Hg++, but the respiration is not affected be-
fore 30 min and is inhibited only 30% maximally after 2 hr (Fig. 1-12-31)
(Demis and Rothstein, 1955). It requires 2 mM Hg++ to inhibit the res-
piration 90% and this occurs after 1.5 hr. Thus glucose uptake is depressed
much more rapidly and is more sensitive than respiration by at least a
factor of 10. These results might imply that Hg++ acts initially on the mem-
brane to block glucose transport, and later on intracellular respiratory sys-
tems; depression of glucose uptake does not in itself inhibit respiration since
the latter is dependent on endogenous substrate. Of some confirmatory evi-
dence is the fact that cysteine will reverse the inhibition of glucose uptake
894
7. MERCURIALS
but will not restore the respiration once it is inhibited; i.e., the surface-
bound Hg++ is available to the cysteine, but penetration of the amino acid
into the cells is inadequate to remove the Hg++ responsible for reducing
the respiration.
Uptake of Hg++ by Diaphragm Muscle
Logarithmic plots of Hg++ uptake with different initial concentrations
in the medium are shown in Fig. 7-38. There appears to be two components,
a fast phase with a half-time of 12 min and a slow phase with a half-time
of around 60 min. The uptake essentially ceases after 20-30 min at low
Fig. 7-38. The uptake of Hg++ by rat diaphragm,
at pH 7.4 and 38°, with time, as determined by the
Hg++ remaining in the medium. (From Demis and
Rothstein, 1955.)
initial concentrations. It was assumed that the fast phase corresponds to
the diffusion of Hg++ into the extracellular space and binding to the plasma
membranes, the slow phase to the penetration into the cells. The time re-
lations point to a correlation between the membrane binding and the inhi-
bition of glucose uptake, and between penetration and respiratory kihi-
bition.
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION 895
There are certain aspects of these results which are puzzling and seem
to me to be difficult to reconcile with the simple theory presented. Why
does the Hg++ uptake cease after the fast phase for the lower initial con-
centrations (0.2 mM or below) (Fig. 7-38)? Let us estimate how much
Hg++ is taken up by the diaphragm in the fast phase (see accompanying
tabulation). If this represents Hg++ bound to membranes and as much as
Initial Hg+ +
Quantity of Hg++
concentration
% Uptake
taken up
(mif)
(/^moles/g tissue)
0.9
63
9.4
0.6
77
7.7
0.2
85
2.8
0.05
84
0.70
0.025
78
0.33
9.4 //moles/g of tissue can be taken up, why does uptake stop when so
little is bound at the lower concentrations? One would expect all the Hg++
to disappear from the medium when the initial amount of Hg++ is less
than that required to saturate the membranes. The maximal Hg++ bound
finally at the highest concentration was stated to be about 15 ^/moles/g
of tissue. Inasmuch as the plasma membranes cannot contribute more
than 1% of the tissue mass, how is it that they can bind over half this
amount ?
The amount of Hg++ diffusing into the extracellular space and existing
there unbound must be negligible, since the diaphragms weighed 0.6 g and
the total medium volume was 10 ml, so that the extracellular space would
be approximately 1 % of the total volume. We have mentioned that Casca-
rano and Zweifach (1962) found diaphragm exposed to Hg++ to show evi-
dence of dehydrogenase inhibition only in the outer few layers of cells
(page 879). Thus Hg++ does not appear to penetrate readily throughout
the tissue. One must ask if the fast phase of uptake may be correlated with
binding only to the membranes of the outermost layer of cells. A rough
estimate of the protein contained in the outermost membranes on both
sides of diaphragms, assuming a generous membrane thickness of 200 A,
gives 1.5 X 10"* //mole/g of tissue. If all the Hg++ taken up w-ere bound by
these membranes, there would be 62,000 Hg++ ions bound per protein
molecule (of assumed molecular weight 100,000), and since this value is
impossibly large, one must conclude that most of the Hg++ must be bound
deeper in the tissue. Demis and Rothstein (1955) assumed that the Hg++
is not bound entirely to the outermost ceUs, but to the plasma membranes
throughout the diaphragm. It is difficult to estimate the amount of protein
896 7. MEKCURIALS
in the total membrane, but it seems very unlikely that it could accommo-
date all the Hg++ taken up at the higher concentrations, especially if pene-
tration deep into the tissue does not occur.
The uptake data by themselves could be explained in a variety of ways.
Binding to proteins often shows different phases due to the different reac-
tivities of the various types of SH group, and in cellular systems one must
perhaps also consider ligands other than SH groups. But how can one inter-
pret the results on glucose utilization and respiration, especially as they
seem to be correlated in time with the Hg++ uptake phases? Particularly,
why is there such a long lag period before respiration is depressed? It may
be noted that a lag period is not always observed in other tissues or cell
suspensions. One possibility which cannot be ignored is that the Hg++ en-
ters the cells early but is initiaUy and preferentially bound to SH groups
not involved with respiration. In muscle cells this might be more evident
than in other tissues because of the large amounts of actin and myosin,
each of which possesses numerous SH groups; only when these groups be-
come saturated with Hg++ would effects on the oxidation enzymes be ob-
served. It is unfortunate that the effects on muscle contraction were not
determined, since if this explanation is valid, contractile activity should
be reduced during the fast phase of uptake. In this case the fast phase
would refer to the binding to membrane and actomyosin (and any other
reactive ligands), the membrane contributing only slightly. The kinetics
of the effects of mercurials on diaphragm contraction have apparently not
been studied, but one notes that the diaphragms exposed to 1 mM p-MB
for 30 min by Kono and Colowick (1961) were stated to be in contracture.
On the other hand, the results obtained with rat atria exposed to 0.05 mM
p-MB indicate that no effect on the contractile amplitude occurs during the
initial 22 min, although effects on the membrane are evident (decrease in
magnitude and duration of the action potential), and that depression of the
contraction proceeds subsequently (Webb and Hollander, 1959). These re-
sults on atria thus would fit into the theory of Rothstein. However, it must
be remembered that in obtaining transmembrane potentials one examines
only the outermost cells, and that contractile amplitude involves the entire
tissue; for this reason one would expect a delay in contractile response. A
decision cannot be made until direct experiments on respiratory and con-
tractile response are made in diaphragms. It must be emphasized that any
modification of the concepts of Rothstein suggested here are not necessarily
applicable to other heavy metal ions or other cells (especially yeast), but
relate to mercurials only.
Another factor which must be considered in tissue uptake studies with
the mercurials is the possibility of damage to the external layers, mani-
fested by increased permeability and exposure of reactive SH groups, espe-
cially with the higher concentrations often used. The high degree of bind-
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION 897
ing observed by Demis and Rothstein (1955) with 0.6-0.9 mM Hg++ might
be due in part to this, the outer layers of cells picking up the Hg++ not
only in the membranes but within the cells. One might try to estimate
roughly the amount of Hg++ required for saturation of membrane sites by
determining the initial concentration so that all the Hg++ is removed from
the medium. If one plots as accurately as possible the amount of Hg++
remaining in the medium after the fast phase against the initial concentra-
tion, one finds that the nearly linear curve passes almost exactly through
the origin. All this shows is that it must require very little Hg++ to saturate
the ligands involved in the fast phase uptake.
It is of some interest to attempt an estimate of the concentration of
membrane SH groups in certain cellular suspensions in order to obtain some
idea of the order of magnitude. In the experiments of Houck (1942) the
suspensions contained 4 x 10^ cells/ml oi Achromobacter fischeri under stand-
ard conditions. Achromohacter is a rod with dimensions 0.9 X 1.8 ;/ and
thus the surface area of a single cell is around 3.8 X 10~^ cm^. If one as-
sumes that the membrane is 200 A thick (which is probablj^ too high), that
the membrane is 50% water (since it is perhaps more condensed than the
cytoplasm), that the membrane solids include 65% protein (values of this
magnitude have been obtained for the membranes of other bacteria), that
protein specific gravity is 1.4., that the mean molecular weight of the mem-
brane proteins is 100,000, and that there are approximately 10 reactive SH
groups on a protein of this molecular weight, one can calculate that the
concentration of membrane SH groups is close to 10"^ rcvM. The lowest
Hg++ concentration to produce reduction of luminescence was 10~^ m.M,
so that even at this lowest concentration the Hg++ was around 100 fold in
excess of the membrane SH groups. Of course, ligands other than SH groups
may occur in the membrane. This suspension of Achromohacter is fairly
dilute relative to most suspensions used, since calculation of total cell vol-
ume indicates that the cells occupy 0.046% of the total volume. In more
concentrated suspensions, such as are often used, the situation can be quite
different. A 10% suspension of human erythrocytes (1.16 X 10^ cells/ml,
cell surface area = 1.4 X 10~® cm^, membrane thickness = 82 A, and 10 SH
groups/protein molecule of molecular weight 100,000) would be 0.059 mM
with respect to membrane SH groups, so that an appreciable amount of
Hg++ might be bound by the membranes in this case. In any study relat-
ing to a theory of membrane binding of heavy metal ions, it would be well
to make some reasonable estimates of the concentration of membrane lig-
and groups. Although such calculations cannot be very accurate, the experi-
mental results may be of an entirely different order of magnitude, which
should impel the investigator to question the validity of the theory.
898 ' 7. MERCURIALS
Comparison of Effects of Hg++ on Intact Diaphragm and Homogenates
The endogenous respiration of diaphragm homogenates fortified with ATP
was claimed by Demis and Rothstein (1955) to be inhibited faster and less
potently than the respiration of intact diaphragm by Hg++. Actually, from
the data given, it is not evident that the rate of inhibition in homogenates
is much faster; at 10 min after adding Hg++, for example, there is no sig-
nificant difference in the rates judged from the points presented, although
from then on the rate in intact diaphragm falls off, so that the inhibitions
are not equivalent again until 50 min. It was stated that it requires 10
times the concentration of Hg++ to inhibit homogenate respiration compar-
ed to intact tissue [in a later review Rothstein (1959) stated 200 times],
but no data on this point are given (the only experiment reported is with
the extremely high concentration of 9 m.M), and in any case it depends
on what time is chosen to compare the inhibitions (e.g., up to 50 min,
homogenate respiration is inhibited more strongly by 9 mM Hg++). It is,
furthermore, very difficult to interpret differences in inhibitions of intact
cells and extracts, since the substrates utilized, the pathways taken, and
the states of the enzymes are probably very different. Mercurial inhibition
has usually been found to be more potent in cell extracts than intact cells,
e.g., Nakayama (1959) reported that while 0.077 mM p-MB inhibits ethanol
oxidation 9% in Acetobacter, it requires only 0.0077 mM to inhibit 14% in
extracts. How much role the membrane plays in any of these observations
is impossible to determine.
Binding of Hg++ to Yeast Cells and Loss of K+
The efflux of K+ from yeast is accelerated by Hg++ as it is from most
cells. Although the effects of the mercurials on permeability and active
transport will be taken up later, the work done by Rothstein and his co-
workers will be treated here since it has bearing on the concept of differen-
tial membrane binding. Rothstein and Bruce (1958) studied the efflux of
K+ into a K+-free medium flowing through a yeast cell column; since the
pH of the medium was 3.5, and lowering the pH enhances the efflux rate,
it was assumed that the process is mainly a K+-H+ exchange. The loss of
K+ from the cells is very sensitive to Hg++, 0.001 mM producing a slight
effect after a long lag period, and 0.003 mM producing at least a tripling
of the rate; at the highest concentration used, 0.1 mM, 80% of the cell K+
is lost in 1 hr.* Passow and Rothstein (1960) used a different technique in
that the rate of K+ loss into a medium (distilled water adjusted to pH 3
with HCl) from a suspension of yeast cells was measured. The minimal ef-
fective concentration of Hg++ to accelerate the efflux was found to be 0.2
* Dr. Rothstein informed me that Fig. 6 of the paper by Rothstein and Bruce
(1958) presents the cumulative K+ loss rather than the rate of K+ loss as stated.
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION 899
ToM, and 1.6 mM produces essentially a complete loss of the cell K+ in
2 hr.* It is possible from the results with the yeast columns that Hg++ at
low concentrations has a specific effect on K+ permeability without depres-
sing active transport, and this is borne out in the work with erythrocytes.
It was stated that the curve obtained by plotting log(IIg++) against max-
imal K+ loss is sigmoid, which fits a "normal distribution" (presumably of
susceptibility of different yeast cells to Hg++), and that the loss of K+ is
probably an all-or-none phenomenon, this being confirmed by determina-
tions of staining by certain dyes in Hg++-treated cells. Although yeast cells
undoubtedly show a variation in the sensitivity to Hg++, it is doubtful if
the evidence is sufficient to categorize the K+ loss as all-or-none, especially
since sigmoid curves of this type (they are not given so one cannot directly
evaluate them) are also compatible with graded effects and, in fact, are the
commonest relations observed in the actions of most inhibitors on cell me-
tabolism or function. There is an increase in general membrane permeability
produced by Hg++, as proved by the loss of a variety of substances from the
cells and a greater penetration of dyes, and this could be a graded phenom-
enon occurring simultaneously with the alterations in K+ efflux, without
the need for assuming cytolysis as the necessary concomitant of K+ loss.
Hg++ is bound relatively rapidly to yeast cells, the half-time being 2-4
min and maximal binding occurring in 15-20 min. Passow and Rothstein
(1960) determined the uptake of both Hg++ and Cl~, and found that ini-
tially only Hg++ is bound, the Cl~ entering when the concentration of Hg++
is sufficiently high. The binding at low concentrations was thus claimed to
represent "binding of Hg++ rather than HgClg." Since the concentration of
the Hg++ ion is actually extremely small, it seems more likely that HgClg
or other chloride complexes react with the yeast cell wall and membrane,
releasing the CI which diffuses into the medium. When the concentration
of the mercurial becomes great enough to lead to a significant increase in
permeability, Cl~ then enters, either alone or with Hg++. The general con-
clusion is that the membrane effect of Hg++ is not specific for K+ but is a
more or less nonspecific breakdown of the membrane, caused by the "mol-
* The approximately 1000-fold difference in sensitivity observed in these two types
of experiment deserves some comment and Dr. Rothstein has kindly provided me with
the reasons. In the suspension experiments the yeast density was 60 mg/ml and at
0.4 mM Hg++ the maximum binding of the metal would be about 7 millimoles/kg
of cells. In the column experiments with 600 mg of cells and a flow rate of 5 ml/min,
the maximum binding in 30 min at 0.05 mM Hg++ would be only 0.015 millimole/kg.
Thus the yeast in the column would be much more readily affected since less of the
Hg++ is removed. Second, the suspension experiments measure the steady-state flux
and the net loss of K+, whereas the column experiments measure the rate of efflux
into K+-free medium. It is therefore difficult to compare the results by the two tech-
niques on a quantitative basis.
900 7. MERCURIALS
ecular stress" brought about by the formation of S — Hg — S bridges in the
membrane; when this stress reaches a critical level, the membrane disinte-
grates and cellular components are released (Rothstein, 1959). Little con-
sideration is given to the possible effects of Hg++ on the active transport
mechanisms by which K+ is accumulated and emphasis is placed on the
structural changes occurring in the membrane. Most of the studies on K+
loss from tissues have been interpreted in terms of an inhibition of active
transport (page 907), and it seems that this would be the more direct and
logical explanation. It should also be pointed out that, as in all studies of
the effects of substances on transmembrane fluxes, it is very difiicult to
distinguish between actions on the membrane and within the cells, and
that therefore these results in themselves cannot be taken as evidence for
a direct or specific membrane effect.
Erythrocyte Permeability and Hemolysis
Organic mercurials increase erythrocyte fragility and promote hemolysis,
often at quite low concentrations, but the effects of Hg++ are more complex,
hemolysis being either favored or inhibited depending on the conditions, of
which the concentration of Hg++ and the type of hemolysis are the most
important. If hemolysis in isotonic glycerol is studied, Hg++ can markedly
delay the hemolysis. Human erythrocytes hemolyze rapidly in isotonic gly-
cerol at pH 7.2; as the concentration of Hg++ is increased, inhibition is first
observed at 0.025 mM and very strong inhibition at 0.05 mM (Wilbrandt,
1941). This was interpreted as an inhibition of glycerol entry into the cells
by Hg++. On the other hand, if hypotonic hemolysis of human erythrocytes
is examined (i.e., hemolysis in Tyrode solution diluted to varying degrees),
Hg++ can either accelerate or slow hemolysis (Fig. 7-39) (Jung, 1947). In
normal or weakly diluted medium, Hg++ favors hemolysis, but at low con-
centration it suppresses hemolysis in markedly hypotonic media. Jung be-
lieved that the resistance to osmotic effects is mediated through a denatur-
ation of the membrane. Arbuthnott (1962) has recently confirmed the dual
action of Hg++, hemolysis of rabbit erythrocytes being promoted by low
concentrations and inhibited by concentrations around 1 mM. Organic mer-
curials {p-MB, ethyl-Hg+, and thimerosal), however, are only lytic, even
at high concentrations. Arbuthnott related this to the number of charges
on the mercurials, although it is more likely a matter of the ability of
Hg+"'" to form S — Hg — S bridges which increase the stability of the mem-
brane. These effects of the mercurials on erythrocytes may or may not de-
pend on metabolic inhibition, but they are important nexvertheless in un-
derstanding the actions of the mercurials on cell membranes in general,
since the mammalian erythrocyte presents an especially simple system for
investigation and has been well studied.
Hg++ appears to have greater lytic potency than the organic mercurials.
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION
901
Minatoya et al. (1960) reported the ED50 for the lytic action on rabbit
erythrocytes to be 0.0034 m.M for Hg++ and 0.0174 m.M for mersalyl, and
Arbuthnott (1962) found that lysis can occur in 1 hr with 0.017 vnM Hg++
whereas it requires 1 mM p-MB or ethyl-Hg+. The effectiveness depends on
the temperature and must also depend on the medium used, since a much
less potent action of Hg++ on rabbit erythrocytes was observed by Joyce
et al. (1954), lysis occurring in 2 hr with 0.13 mikf. p-MB is much more
lytic to rat erythrocytes than is PM, 0.1 vaM of the former lysing almost,
completely in 40-60 min, whereas at this time 0.5 mM PM produces only
about 50% hemolysis (Moore, 1959), and Hg++ is about 3 times as potent
as p-MB, 50% hemolysis being given by 0.4 raM Hg++ and 1.2 mM p-MB
(these values estimated from data given) in 90 min (Tsen and Collier, 1960).
It is obviously difficult to compare results obtained by different investiga-
tors, even when the same species is used, but the definite difference in po-
tency between the various mercurials is clear. Although the role of SH
groups in erythrocytic membrane structure and function is important, ex-
actly how they operate in this capacity is unknown, so it is difficult to
speculate on either the mechanisms of hemolysis by the mercurials or the
reasons for the differences between the mercurials. Other cells do not lyse
so easily in the presence of mercurials, but this does not necessarily prove
that SH groups are of more importance for the erythrocytic membrane,
since the inherent stability may be less.
100
80
60
20
Fig. 7-39. Hemolysis of human erythrocytes by Hg++ at different
fractional dilution of Tyrode solution {r). The control curve shows the
hemolysis in the absence of Hg++. (From Jung, 1947.)
902 7. MERCURIALS
It is interesting to inquire into how much Hg++ must be bound to the
erythrocytic membrane to cause hemolysis. The data of Meneghetti (1922)
indicate about 1.5 X 10' atoms/cell, but Jung (1947) believed this to be
too low and revised the figure on the basis of his results to 1.4 X 10^ atoms/
cell. The data of Vincent and Blackburn (1958) allow a rough calculation
that K+ loss is induced by Hg++ at binding levels around 2 X 10' atoms/
cell, although no hemolysis occurs, while maximal K+ loss and inhibition
of glucose uptake in human erythrocytes were found by Weed et al. (1962)
to be produced by 3.6-4.5 X 10^ atoms/cell. If there are 10 reactive SH
groups for each membrane protein of molecular weight 100,000, one can
estimate there to be around 3 X 10' SH groups per erythrocyte membrane.
However, although stromal SH groups have a greater affinity for Hg++,
hemoglobin SH groups account for around 85% of the total binding, so the
figures given above should be reduced if only membrane binding is desired.
All one can say is that the amount of Hg++ to alter membrane properties
is of the same order of magnitude as the estimated SH content of the mem-
brane. On the other hand, the number of molecules/ceU of the organic mer-
curials required for hemolysis is greater than necessary to cover the sur-
face of the sheep erythrocyte (Benesch and Benesch, 1954). For PM there
is a 4-fold excess and for mersalyl a 24-fold excess. Of course, the organic
mercurials probably do not lie flat on the membrane, but, more important,
it is not known how much of the mercurial is bound to hemoglobin or other
nonmembrane components.
The kinetics of mercurial hemolysis are generally characterized by a lag
period, the duration of which is dependent on the mercurial concentration,
followed by a rather sudden hemolysis (Fig. 7-40). For sheep erythrocytes
there is a lag period of around 80 min when treated with 0.45 milf PM
(Benesch and Benesch, 1954), and for human erythrocytes the lag period
is 90 min at 37^ when exposed to 0.5 mM p-MB (Sheets et al, 1956 a). The
temperature is an important factor, since in the latter case the lag period
is around 200 min at 25°. The lag period is partly due to the slow binding
of these organic mercurials. Washing the erythrocytes 1 min after exposure
to p-MB protects completely, after 30 min protects partially, and after
60 min there is no protection (Sheets et al., 1956 a). On the other hand,
there is maximal uptake of Hg^"^ by erythrocytes within 5 min (Weed et
al., 1962). The osmotic fragility is altered after 3-min exposure to Hg++:
At 5.2 X 10'-5.5 X 10^ atoms/cell there is a decrease in the fragility, at
4.5 X 10^ atoms/cell there is an increased fragility and hemolysis. The ki-
netics for Hg++ and the organic mercurials appear to be quite different. The
very marked effects of PM concentration on the kinetics of hemolysis may
be seen in Fig. 7-40 for sheep erythrocytes, and a similar dependence has
been noted in rat erythrocytes (Moore, 1959). This might indicate a rather
critical level of membrane binding to produce hemolysis. The uptake of
THE CELL MEMBRANE AS A SITE FOR MERCURIAL ACTION
903
Hg+''" by erythrocytes or ghosts is very rapid but the situation with chlor-
merodrin is different, in that binding to ghosts is rapid but uptake into
erythrocytes continues for 2 hr or more; the binding of chlormerodrin is
also perhaps more specific for certain SH groups (Rothstein, 1964). Chlor-
100
Fig. 7-40. Hemolysis of sheep erythrocytes in a 2% suspension by PM
at pH 7.4 and 37°, showing the marked differences over a narrow con-
centration range. (From Benesch and Benesch, 1954.)
merodrin thus might be useful in separating the effects of membrane and
internal binding.
Is hemolysis by the mercurials in any way related to effects on glucose
uptake or metabolism? This question cannot be satisfactorily answered
since there has been little work where metabolic and hemolytic actions can
be compared, and the results available are divergent. The utilization of
glucose by human erythrocj'tes is inhibited moderately within a range of
Hg++ concentration, the inhibition disappearing as the amount of Hg++
bound is increased (Fig. 7-41) (Weed et al., 1962). In the reversal range, a
change in the hemoglobin was observed and some agglutination of the cells
occurred. The question arises as to whether the effects on K+ loss and glu-
cose utilization result from some action on all the cells or are due to hemol-
ysis of a few cells. Weed et al. (1962) assumed that high Hg++ concentra-
tions denature the membrane, causing a decrease in permeability, which
could explain the reversal of the effect on K+ loss, but is difficult to reconcile
with the disappearance of the effect on glucose utilization. It is interesting
to compare their results on osmotic fragility with these actions. At the Hg++
904
7. MERCURIALS
level producing the maximal K+ loss and inhibition of glucose utilization
(6.0-7.5 X 10"^^ mole Hg/cell), it is claimed that the fragility is decreased,
which it is in very hypotonic media, but examination of the curves shows
that some hemolysis (probably around 2-5%) has occurred in normal me-
dium. At a higher level of bound Hg++, where the effects on K+ and glu-
cose have been partly reversed (7.6 X 10~^^ mole Hg/cell), fragility is
definitely increased, and some 20% hemolysis has occurred in the normal
medium within 3 min. It seems clear, therefore, that the K+ loss and sup-
pression of glucose utilization at lower levels of bound Hg++ (left of the
50-
40
30
20
10
1000
MOLES Hg BOUND/RBC « 10
Fig. 7-41. Effect of Hg++ on the loss of K+ from human erythrocytes
and the uptake of glucose. (From Weed et al., 1962.)
maximum in Fig. 7-41) are due almost entirely to effects on all the cells
and not on lysis of a fraction of the cells. However, at higher levels of
bound Hg++, lysis must contribute to both K+ loss and interference with
glucose utilization; e.g., at 7.6 X 10~^^ mole Hg/cell bound, there is 20%
lysis and around 20% loss of K+. If this is so, the disappearance of the
effect on K+ loss from intact cells must occur even more precipitously than
appears in the figure (the reversal is, of course, really not precipitous, since
it is a logarithmic scale; to reverse these effects appreciably it requires the
binding of about 10 times that amount of Hg++ necessary for maximal K+
loss). The results of Jacob and Jandl (1962), also on human erythrocytes,
are quite different, since they observed that p-MB does not inhibit glucose
uptake or lactate formation — indeed, stimulates these somewhat — up to
5 //moles p-MB/ml of cells, which is around 2 x 10"^^ mole 2>-MB/cell.
There is also no reaction of p-MB with the intracellular glutathione. These
results point to a failure of p-MB to penetrate through the membrane.
Hemolysis occurs and is presumably due to an action on the membrane.
However, it is rather strange that sufficient mercurial can be bound to the
THE CELL MEMBKANE AS A SITE FOR MERCURIAL ACTION 905
membrane to cause lysis and yet have no inhibitory effect on glucose uptake.
One might conclude that Hg++ penetrates into the erythrocytes more readi-
ly than 2?-MB, which is undoubtedly the case, but it is also possible to spec-
ulate that the bifunctional Hg++ can distort the membrane pores by form-
ing S — Hg — S bridges in such a way that glucose penetration is slowed
while K+ permeability is increased, as in the concept of critical pore sizes
formulated by Mullins (1960). Another problem is how these results can
be reconciled with those of LeFevre (1948), who showed that glucose utiliz-
ation by human erythrocytes is inhibited by 0.002 mM p-MB and abolished
by 0.01 mM.
The very rapid loss of K+ from human erythrocytes observed by Weed
et al. (1962) — maximally 50% of the total cell K+ in 3 min at 7.5 X 10-i«
mole Hg/cell — is not seen in rabbit erythrocytes, from which there is a
slow loss of K+ in the presence of Hg++, a rapid loss occurring only upon
hemolysis, a result which is quite reasonable (Joyce et al., 1954). It is dif-
ficult to compare the results of these two groups of investigators because
the Hg++ concentrations are expressed differently. However, it is possible
to estimate that when 7.5 X 10~^^ mole Hg/cell is bound, the free Hg++
concentration is roughly 0.01 mM (see Fig. 2 of Weed et al.). In the work
of Joyce et al., 0.032 mM Hg++ caused a 30% loss of total K in 4 hr, so
that apparently there is a very marked difference in the response of rabbit
and human erythrocytes to Hg++.
The question as to the relation of glucose metabolism to hemolysis is
still unanswered. There is one observation which suggests a relation, the
finding by Moore (1959) that 10-100 roM glucose inhibits the hemolysis
induced by p-MB, this being manifest mainly in a lengthening of the lag
period. No reaction between j^-MB and glucose can be detected spectrosco-
pically and it is not an osmotic effect. Certain other sugars, e.g. fructose
and sorbose, are also effective. It was postulated that glucose may combine
with some component of the rat erythrocyte and protect it from the mer-
curial; if so, this would probably be the transport system for glucose, which
is inhibited readily by p-MB, the situation being similar to the protection
of enzymes by substrates. Sheets et al. (1956 a) had found that glucose exerts
no protection against hemolysis of human erythrocytes by p-MB, but the
glucose was added at various times after the p-MB and was only 3.3 mM.
Membrane or transport ATPase is inhibited by the mercurials but the
Hg++ or chlormerodrin which is initially bound is without effect (Roth-
stein, 1964). Chlormerodrin can bind to about 3% of the total SH groups
without inhibition of ATPase, but by the time of maximal binding with
25% of the SH groups the ATPase is inactivated, possibly leading to the
loss of K+, although some increase in permeability may also play a role.
The possibility that mercurial hemolysis is related to the reaction of
erythrocytic glutathione with SH reagents was considered by Tsen and
906 7. MERCURIALS
Collier (1960). However, Hg++ and p-MB can produce hemolysis without
significant loss of glutathione, whereas iodoacetate and iV-ethylmaleimide
reduce the glutathione completely without lysis. Jacob and Jandl (1962)
also showed that p-M3 reacts readily with glutathione in solution, but does
not attack erythrocyte glutathione, and concluded that p-MB does not
penetrate into the cells. However, Weed et at. (1962) found that of the three
major sources of SH groups in the erythrocyte — stroma, hemoglobin, and
glutathione — the last has the lowest affinity for the mercurial and consti-
tutes only 5% of the total SH groups. It is thus possible that glutathione
would be reacted only when aU the other SH groups are saturated. In any
event, it is evident that glutathione does not play a significant role in
hemolysis.
Hg++ is able to produce structural changes in the erythrocytic membrane
which are detectable by electron microscopy (Jung, 1947). Isolated hemo-
globin-free membranes treated with high concentrations of Hg++ (37 ulM)
show gross changes in structure — a crumpling with increased density and
apparent thickness — but with lower concentrations (0.37 vaM) the pic-
ture is different, a network of holes appearing in the otherwise unaltered
membrane. Intact erythrocytes treated with 1.85 ruM Hg++ for several
hours no longer lyse in distilled water, and the membrane is seen to have
been replaced by a thick mass of coagulated protein. Certain changes in
the over-all erythrocyte configuration were also observed by Vincent (1958)
in preparations allowed to bind Hg++ for 5 min, especially sphering and
crenation. Possibly a more detailed study of structural changes induced by
low prohemolytic concentrations of the mercurials would be useful in clari-
fying the mechanism of hemolysis.
We have assumed with others in this discussion of hemolysis and permea-
bility changes brought about by the mercurials that SH groups only are
attacked. Certain nonelectrolytes, such as glucose and glycerol, enter the
erythrocyte by facilitated diffusion and, since the transport is usually ef-
fectively blocked by SH reagents, it has been thought that SH groups are
involved in some manner. We have seen that Wilbrandt (1941) claimed a
marked -reduction in glycerol permeability with 0.05 vaM Hg++. Further-
more, it was believed that inhibition of glycerol penetration occurs only
while the Hg++ is entering the cells, i.e., when the Hg++ is bound to the
membrane. When the Hg++ has been picked up by the hemoglobin, there
may be little left in the membrane and the permeability to glycerol is re-
stored. LeFevre (1948) established that p-MB likewise blocks glycerol entry
into the human erythrocyte. However, Barnard and Stein (1958) have sug-
gested that an imidazole group is involved in this transport. The fact that
histidine as well as cysteine can reverse the inhibition (it requires a 5- to
10-fold excess of histidine) is not valid evidence; it simply shows that mer-
curials are bound to histidine. It was also claimed that mercurial action
EFFECTS ON PERMEABILITY AND ACTIVE TRANSPORT 907
was characterized by a lag period, this being due to the preferential binding
to SH groups before the imidazole groups are attacked; however, there are
other possible reasons for such a lag period, and indeed Wilbrandt (1941)
claimed the inhibition occurs before the mercurial is bound intracellularly.
They also point out that Cu++ is much more potent than p-MB in depressing
glycerol entry and that this favors an imidazole group; p-MB is, however,
rather ineffective relative to Hg++, which exerts an effect at 0.025 mM, due
possibly to steric factors. It does not appear that the evidence is sufficient
to establish an imidazole group as involved in the glycerol transport, but
one cannot argue against this theory, and it is quite possible that in the
complex mechanisms of penetration there are both SH and imidazole groups.
In either case, one cannot attribute an active role to these groups in the
transport on the basis of the evidence available.
EFFECTS ON PERMEABILITY AND ACTIVE TRANSPORT
The general discussion of the mechanisms by which transport systems
in the membrane may be affected by SH reagents (see III-1-171, 180) is
applicable to the mercurials. We shall confine our attention to certain
important problems and interesting results, as far as possible, and only
summarize most of the studies in Table 7-18. The effects of the mer-
curials on renal transport wiU be taken up in the following section. It
is clear from the results in the table that the mercurials often cause a loss
of intracellular substances, e.g., K+, carbohydrate, and amino acids. It is
likely that a good many substances leak out of cells treated with the mer-
curials as a result of not only interference with active transport but direct
distorting effects on the membrane structure leading to increases in per-
meability. Possibly coenzymes, enzymes, and other large molecules may be
lost. Ohr (1960) observed the release of some ultraviolet-absorbing material
from diaphragm exposed to Hg++, and Weed et al. (1962) detected the early
release of some Hg++-complexing material from human erythrocytes, this
altering the binding kinetics at low concentrations of Hg++. It is not easy
to determine if the action is primarily on active uptake or on outward dif-
fusion, even with labeled substances. For example, if one is studying K^^
efflux, an inhibition of a pump involved in maintaining a high intracellular
K+ level might alter this efflux, either by changing the membrane potential
or directly if part of the K+ efflux is mediated by the pump, since a Na+
pump might not be completely specific for Na+ and might carry some K+
out of the cell. If K*^ influx is measured, an alteration of the permeability
could easily change the rate at which active transport occurs, particularly
if K+ loss accelerates the pump. In most cases there seems to be a decrease
in the intracellular K+/Na+ ratio, but the mechanism is not clear. Further-
more, a decrease in transport is occasionally not accompanied by a signifi-
908
7. MERCURIALS
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EFFECTS ON PERMEABILITY AND ACTIVE TRANSPORT
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7. MERCURIALS
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EFFECTS ON PERMEABILITY AND ACTIVE TRANSPOET
911
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912 7. MEECURIALS
cant depression of respiration, as in the uptake of I" by Fucus, where 0.05
YoM p-MB inhibits transport 50% but 0.2 mM does not affect respiration
(Klemperer, 1957), or the accumulation of K+ by Porphyra, where p-MB
decreases the number of ions pumped per Og consumed (Eppley, 1960), or
the active transport of Na+ through frog skin, which is blocked by Hg++
at a concentration not altering respiration (Linderholm, 1952). In such
cases it has generally been assumed that the action is on the transport
system itself, but this is not necessarily true. It is rather surprising that so
few have reported instances of decreased permeability brought about by
the mercurials, particularly the organic ones, inasmuch as they might be
expected to react with SH groups in or around the membrane pores to
impede the passage of substances across the membrane; perhaps this would
be observed more often if lower concentrations were examined. The per-
meability of frog skin to Cl~ is decreased by p-MB, and Janacek (1962)
postulated that the mercurial sterically hinders the movement of anions
through the pores.
Certain results occasionally point to an effect of mercurials on the end-
ergonic phase of transport rather than a simple depression of ATP forma-
tion. The fact that Hg+^ at 0.5 mM inhibits the 20-sec uptake of acetate
by diaphragm without a lag period (Foulkes and Paine, 1961), taken with
the rather slow depression of metabolism, is indicative of an action directly
on the membrane. We have also seen that p-MB lyses erythrocytes without
reacting with intracellular glutathione or inhibiting glycolysis, in contrast
to A^-ethylmaleimide, and that this has been attributed to a failure to pene-
trate into the cells, so that the effects observed must involve an attack on
the membrane (Jacob and Jandl, 1962). Hg++ very potently inhibits stro-
mal ATPase — 50% inhibition at around 0.00125 mM, and plots of log
{vjvi — 1) against log (Hg++) suggest that 3 Hg++ ions are required for
each ATPase molecule (Laris et al., 1962). The same type of behavior was
observed for the inhibition of glucose uptake, and the concentrations of
Hg++ required to inhibit are comparable (LeFevre, 1954). Laris et al. re-
plotted LeFevre's data and found that roughly 6 ions of Hg++ are necessary
for the inhibition of the transport of each glucose molecule. The similar
sensitivities and kinetics allowed them to postulate that the two inhibi-
tions may be closely related. The stimulation of the uptake of certain sugars
(e.g., D-xylose and L-arabinose) into diaphragm by p-MB, and the inhibition
of the stimulation produced by insulin, may well be on the muscle mem-
branes (Kono and Colowick, 1961). There is certainly no correlation with
the level of ATP, and p-MB actually seems to increase ATP somewhat. A
block between ATP and the transport system was considered a possibility.
The reaction of mercurials with a membrane carrier was adduced to ex-
plain the inhibition of phosphate transport in Micrococcus pyogenes by Hg++
and PM (P. Mitchell, 1953). An inhibition of 50% is given by 2.2 //moles
EFFECTS ON PERMEABILITY AND ACTIVE TRANSPORT 913
PM/g cells and by 4.7 //moles Hg++/g cells. The inhibition-concentration
curves are said to conform to the equation ^ = (I) (1 — i)ji and hence to
suggest reaction of the mercurial with a phosphate carrier X, according to
I + X :±5: IX.* This equation is simply that for noncompetitive inhibition
and it is difficult to understand how it would serve to indicate any partic-
ular mechanism by which transport is depressed. Mitchell then proceeds
to calculate the number of carrier molecules for 100 molecules of intra-
cellular phosphate; since the cells contained 147 //moles P,/g cells, 1.5 and
3.2 molecules of PM and Hg++, respectively, are required for 50% inhibition
per 100 molecules of internal P^. It was apparently assumed that if 50%
inhibition is given by these numbers of molecules, 100% inhibition would
be given by twice these, namely, 3.0 and 6.4 molecules of PM and Hg++,
respectively. If this were true titration or zone C inhibition, this would be
correct, but inspection of the curves shows that it is not; indeed, it is evi-
dent that approximately 20 and 40 //moles/g cells of PM and Hg++ are
needed for 90% inhibition (curves do not reach complete inhibition, which
would require appreciably more of the mercurials). Therefore, his conclusion
that the number of carriers is not more than 3% of the internal P, is not
valid. In addition, the mercurials must be bound to cell components other
than a hypothetical carrier, so that under any circumstances it would be
difficult to estimate the relative amount of carrier present, just as it is
impossible to calculate the amount of an enzyme present in a complex
mixture by the quantity of mercurial required for 50% inhibition.
Transmembrane and Transcellular Transports
The uptake of a substance into a cell is often a process different from
the transport across a layer of the cells. If a substance is moved against
a concentration gradient from one medium into a similar medium, it is
an active transport, whereas accumulation of a substance within a cell can
be the result of binding. A good example of this is the transport of triiodo-
thyroacetate by rat intestine (Herz et al., 1961). The mucosal -^ serosal
transport is inhibited 93% by 1 raM Hg++, but the accumulation in the
tissue is actually accelerated 16%. The cellular uptake was postulated to
be due mainly to binding. The accumulation of Fe+++ (Saltman et al, 1955)
and Cu++ (Saltman et al., 1959) by rat liver slices is slightly stimulated by
p-MB, and it is very likely that these are instances of binding to intracel-
lular ligands. There is sometimes not so clear a separation of transmembrane
and transcellular transports. Rat intestinal slices accumulate Ca++ to a tis-
* Mitchell gives the equation as ^ = (I)i7(l — i), changing his symbols to those
used in the present work, which is obviously incorrect, since it would mean that the
inhibition would vary inversely with the inhibitor concentration, so I have taken the
liberty of rewriting it.
914 7. MERCURIALS
sue/medium ratio of 5.8, and 1 mM Hg++ reduces this to 1.5 (Schachter
et al., 1960). The transport across the intestinal wall leads to an inside-
outside ratio of 4.6, and 1 mM Hg++ drops this to 1.1 (Schachter and Rosen,
1959). Thus in this instance there is no significant difference in the mercurial
inhibition, but certain other inhibitors affect the transintestinal process
more strongly. The results wiU often depend on the location of the active
transport mechanism. We shall see that this is an important point in con-
sidering the effects of the mercurials on the kidney.
Mitochondrial K+
Mitochondria isolated from rat liver in 0.25 M sucrose contain 620-640
-//moles K+/g N and this can be lost if the mitochondria are placed in hypo-
tonic media or treated with saponin, 2,4-dinitrophenol, or Hg+"'" (Spector,
1953). Most of the K+ appears to be free, but a fraction may be bound; the
retention of the free K+ is dependent on oxidative phosphorylation. How-
ever, Gamble (1957) found that mitochondrial fragments catalyzing oxida-
tive phosphorylation can bind K+. This binding is not dependent on ATP
but is abolished by 2.4-dinitrophenol, and a relation between the K+ bind-
ing and the sites for oxidative phosphorylation was postulated. Hg++ at
0.01 mM and p-MB at 0.03 mM produce a 5-fold increase in the K+ ex-
change rate. This was later investigated in detail (Scott and Gamble, 1961),
and Hg++ in concentrations around 0.01 mM was found to increase the ex-
change rate, reduce the bound K+ by 50%, and inhibit phosphorylation
50%. The organic mercurials are less effective. These three actions are pre-
sumably not mediated through the same mechanism, since EDTA prevents
the effects of Hg++ and p-MB on K+ binding, has no effect on the stimulation
of the exchange rate, and protects oxidative phosphorylation from Hg++
but not from p-MB. These complex relationships are not completely under-
stood at the present time, but obviously are of importance in certain cases
of K+ accumulation and transport.
Gastric Acid Secretion
Reduction of gastric acidity by 0.25 mM Hg++ introduced into the stom-
ach was shown by Mann and Mann (1939), and the mechanisms involved
were studied by Davenport and his group at Utah. Gastric secretion of
HCl is depressed to a basic level by 1 mM p-MB; if the secretion is stimul-
ated by carbachol or histamine, the inhibition appears to be greater but
the rate is reduced to the same level (Fig. 7-42), i.e., p-MB effectively abol-
ishes the secretion brought about by these drugs (Davenport, 1954, Da-
venport et al., 1954). There is thus a basal level of secretion (around 30%
of maximal) resistant to the mercurials. Graphical analysis indicated that
2 SH groups are involved in the inhibition. Lactate formation when glu-
EFFECTS ON PERMEABILITY AND ACTIVE TRANSPORT
915
cose is the substrate is inhibited by p-MB, one SH group being involved
here, but it is unlikely that glycolytic inhibition is the mechanism by which
acid secretion is depressed, inasmuch as inhibition occurs when pyruvate or
acetoacetate is the substrate. Respiration associated with secretion is also
inhibited by p-MB; it was felt that this is not a generalized effect on oxi-
0 72
0 63
0.54
0.45
0 27
0 09
CARBACHOL (0 1 MG X)
0.2 0.4 0.6 0.8 1.0 1.2 1.4 18
Fig. 7-42. Inhibition of gastric acid secretion by
p-MB, in the presence of 20 raM glucose and in
the absence and presence of carbachol. (From
Davenport et al., 1954.)
dative reactions, but that the site of attack is some unknown system inti-
mately concerned with the secretory process. The relation between the inhi-
bitions of respiration and secretion by p-MB is reasonably linear (Fig. 7-43),
in contrast to the results with antimycin and 2,4-dinitrophenol (Davenport
and Chavre, 1956). Possibly the primary inhibition by j^-MB is on the trans-
port system itself, the respiration being reduced secondarily. Other SH
reagents inhibit secretion but apparently act at somewhat different sites
than p-MB (Davenport et al., 1955), so that it is difficult to correlate the
blockade of SH groups with the secretory suppression or to locate exactly
these SH groups.
916
7. MERCURIALS
Intestinal Transport
The transports of Na+, water, and glucose across the rat intestine are
inhibited by Hg++ (Clarkson and Cross, 1961). The transintestinal electric
potential is dependent on the Na+ transport and the ionic permeabilities
of the lumenal membranes. Hg++ 0.01-1 mM causes a rapid brief elevation
of the potential which is followed by a fall, the rapidity of which is deter-
mined by the Hg++ concentration. There are two phases in the response:
(1) an immediate loss of K+ and phosphate from the intestinal wall and a
marked inhibition of glucose uptake, and (2) a delayed (occurring after
20 min or longer) inhibition of transintestinal transport of Na+, water, and
0 4
0.3
0.2
0.1
\
-^/
ANTIMYCIN /
/p-MB
)
" //
'
^/dnp
' ^ -^
.^
0.1
0.2
03 0.4 0.5 0.6 0.7 0.8
Fig. 7-43. Effects of inhibitors on the respiration and acid secretion
of mouse gastric mucosa stimulated by carbachol. (From Davenport
and Chavre, 1956.)
glucose, with a suppression of lactate formation. The uptake kinetics of
Hg++ show two phases, a fast component dominant during the initial 20-
30 min of exposure (k^ = 0.0032 min'^) and a slower component (^2 —
0.0017 min-^). It was pointed out that the system is so complex that it is
difficult to interpret the uptake data, but possibly there is some correlation
with the initial and delayed responses discussed above. Analysis showed
that the potential changes are produced when certain quantities of Hg++
are bound to the intestine; i.e., when different concentrations of Hg++ are
applied to the intestine, the potential changes occur at different rates which
are related to the uptake rates. Since the transports and potential are de-
EFFECTS ON THE KIDNEY 917
pendent on glucose, it was suggested that the inhibitions of transintestinal
transport are perhaps all the result of interference with glucose uptake into
the cells. This does not explain the immediate responses, which may be
due to the direct effect of Hg++ on the cell membranes. Indeed, all the
changes observed may arise from modifications in the permeability prop-
erties of the lumenal membranes.
EFFECTS ON THE KIDNEY
The clinical effectiveness of the mercurials in certain edemas has stimulat-
ed much investigation directed at discovering the mechanism by which diu-
resis is produced. It is now clear that the action is on tubular transport
processes. Since these transports are mainly active and depend on tubular
cell metabolism, as well as on certain specific carrier and enzyme systems,
we shall be primarily concerned with the possible effects of the mercurials
on renal metabolism as a basis for their diuretic activity. The pharmacology
textbooks and recent reviews (Beyer and Baer, 1960; Farah and Miller,
1962; Kessler, 1960; Mudge and Weiner, 1958; Orloff and Berliner, 1961;
Pitts, 1958, 1959) cover the general renal actions of the mercurials and also
discuss many of the controversial points. We shall confine ourselves here to
a summary of these actions and then proceed to the mechanisms which may
be involved in the alteration of the transport systems. The structures of the
four most common mercurial diuretics used experimentally are shown in
their ionic forms. The preparations provided for clinical use are complexed
with different ligands (OH^, Cl^, thioacetate, or theophylline), but once
introduced into the body or experimental media, the mercurials usually
establish new equilibria with the available ligands, as discussed previously
OCHjCOO" H3C CH3
r,:^ OCH, bOC^N (-CH, OCH3
L ))— CONH-CH2— CH— CHj— Hg V— CONH— CH— CH— CH2— Hg"
Mersalyl (Salyrgan) Mercurin (mercaptomerin, Thiomerin)
OCH3
I
HjN— CONH— CHj— CH— CH2 — Hg
Chlormerodrin (Neohydrin)
OCH3
I
OOC— CH,CH,— CONH— CONH— CH,— CH— CHj— Hg
Meralluride (Mercuhydrin)
918 7. MERCURIALS
(page 742). Thus mercurin (Mercuzan) is the un-ionized acid, the Na+ salt
is Novurit, the complex with thioacetate is mercaptomerin (Thiomerin),
and the complex with theophylline is Mercurophylline (Mercuzanthin), but
the fundamental active structure is the same in all cases. The complexers
alter the solubility, local actions on tissues, and rates of absorption, but
probably do not significantly affect the basic effects on the tubular trans-
port systems.
Summary of the General Renal Actions of the Mercurials
(A) The diuretic action is entirely renal. Since the classic transplantation
and unilateral injection studies of Govaerts in 1928 and Bartram in 1932 it
has been clear that any extrarenal actions of the mercurials are unimportant
for diuresis. Effects on tissues other than the kidneys may of course occur,
especially at high doses, but do not contribute significantly nor are they
necessary for diuresis.
(B) The primary action is on tubular transport rather than glomerular fil-
tration. Clearance studies have demonstrated that glomerular filtration rates
are not altered much or at all during marked clinical diuresis. Analyses of
the changes in composition of the filtrate as it passes down the nephron
show that the increased flow of urine can be accounted for entirely by the
depression of certain tubular transport processes. Mercurials given intramus-
cularly or orally to human subjects do not affect renal blood flow or glom-
erular filtration (Brun et al., 1947), but injected intravenously in animals
they produce definite effects which may possibly modify the primary diu-
retic action. Vasoconstriction and a reduction in blood flow are usually ob-
served. Jackson (1926 b) reported that intravenous mersalyl causes a rise
in blood pressure and a profound constriction of the kidney (measured on-
cometrically), while Farah (1952) observed the renal blood flow to fall 50%
from mersalyl at 10 mg/kg intravenously and 70% from 4 mg/kg intraarte-
rially, these changes occurring before the onset of diuresis. Kessler et al.
(1957 b) invariably found a significant decrease in glomerular filtration rate
following various mercurial diuretics given intravenously, and such often
persists long after the maximal diuretic effect occurs. Occasionally a trans-
ient antidiuresis is noted after intravenous diuretics and this could be the
result of renal vasoconstriction and a reduced glomerular filtration rate
(Vargas and Cafruny, 1959; Cafruny and Palmer, 1961). It is interesting to
note that the nondiuretic p-MB likewise produces these vascular changes.
Thus the effects on glomerular filtration when observed experimentally
would be antidiuretic and might reduce the over-all diuretic response in-
stead of favoring it.
(C) The diuretic action is mainly due to an inhibition of active iVa+ resorp-
tion in the proximal tubules. It is now generally agreed that the mercurials
EFFECTS ON THE KIDNEY 919
are primarily natriuretics, and that the resorption of Cl~ and water by the
proximal tubules follows the movement of Na+ for electrostatic and osmotic
reasons. The evidence for the proximal site of action of the mercurials will
be presented later (page 920). The fundamental mechanism of action must
therefore be sought in the modifications of active Na+ transport by the
mercurials.
(D) The mercurials can also act elsewhere on the nephron to modify urine
composition and flow rate. Sufficient evidence has been accumulated to show
that the mercurials can exert minor effects throughout the nephron — on
the loop of Henle, the distal tubule, and the collecting duct — to further
alter the urine volume, and that transport processes and exchange reac-
tions of various types can be inhibited. That is, the primary site of action
may be on active Na+ transport in the proximal tubules, but this is by no
means the sole site of action. The transports of a variety of substances, in-
cluding K+, Ca++, urate, p-aminohippurate, amino acids, and various dyes,
are depressed by the mercurials.
(E) Only a relatively small effect on Na+ resorption need be exerted to pro-
duce marked diuresis. Inasmuch as 98-99% of the filtered Na+, CI", and
water is resorbed, it is evident that a reduction of this to 90-95% would
cause up to a 10 fold increase in excretion rate. Consequently one might
predict that only a small metabolic disturbance by the mercurials would be
necessary for diuretic action, and that so small an effect might be difficult
to detect under the usual conditions. Furthermore, only 15-30% of the
total NaCl resorption can be inhibited by the mercurials, the remainder
presumably being mediated through mercurial-resistant systems (Pitts,
1958).
(F) The selective action of the mercurials on the kidney is mainly a conse-
quence of the accumulation of mercurial. Mercurials exert demonstrable ef-
fects only on the kidneys over a dosage range, and this is primarily due to
the relatively high concentrations of mercurial reached in the renal tissue,
whether this is achieved by tubular secretion or filtrate resorption (page
928). The transport systems are probably no more sensitive than in other
tissues to the mercurials (at least there seems to be no clear evidence for
this). However, the point mentioned in the previous paragraph that only
small effects on renal transport need be exerted may be a factor in increas-
ing the apparent sensitivity of the kidney.
(G) The reported renal responses to the mercurials are quite variable. One
cannot fail to be impressed by the general lack of agreement on certain
basic actions of the mercurials despite the great amount of work done over
many years, and it is disturbing that almost every hypothesis can be re-
futed by evidence of apparent validity. It may be helpful to list some of the
reasons for these disagreements. (1) Work done with different species can
920 7. MERCURIALS
frequently not be compared. For example, mercurials in diuretic dosage
inhibit glucose resorption and p-aminohippurate secretion in man, but have
no effect on these transports in the dog; also p-MB is not diuretic in the
dog, but increases urine flow in the rat (Cafruny and Palmer, 1961). (2)
Animals in different states of water load, ion load, or pH wiU respond dif-
ferently to the mercurials. (3) The use of theophylline-containing mercurials
has often confused interpretation, since theophylline itself is a diuretic act-
ing by a mechanism quite different than the mercurials. Thus Goldstein et
al. (1961) found Mercuhydrin (meralluride complexed with theophylline)
to produce two phases of diuresis, the first due to the theophylline. Certainly
some of the results attributed to the mercurials have had their origin in the
theophylline present, and for this reason it is always advisable to use mer-
curials complexed with inactive substances if a pure mercurial action is to
be investigated. (4) Much of the work on distribution of the mercurials and
their actions on enzymes in the kidney has been done with toxic or lethal
doses or concentrations. If a mechanism for the normal diuretic effect is to
be found, one must use mercurial concentrations which do not deviate ap-
preciably from those producing maximal diuresis. (5) Different routes of ad-
ministration often lead to different results. We have seen that intravenous
injection causes changes in blood flow and glomerular filtration not seen
with the usual routes of administration.
Sites of Action in the Nephron
Several types of evidence have been used to locate the major sites of
mercurial action on renal transport processes; these will be discussed briefly,
since they also provide interesting information on the mechanisms involved.
(A) Inhibition of transport processes located in different regions of the
nephron. The mercurials interfere with the transport of a variety of sub-
stances by the proximal segment of the nephron. This includes the resorp-
tion of glucose, amino acids, urate, phosphate, bicarbonate, Na+, K+, and
Ca++, and the active secretion of p-aminohippurate, iodopyracet (Diodrast),
tetraethylammonium ion, phenol red, and various dyes. Izar (1909) noted
an increase in urinary urate in dogs given HgClg intravenously, and Dale
and Sanderson (1954) demonstrated that urate excretion in man rises rap-
idly following administration of mersalyl. However, if oliguria is produced
by lethal doses of HgCla, urate excretion is impaired and the tissue concen-
tration will rise (Wells, 1916). Mild poisoning by mercurials leads to an
aminoaciduria in man (Clarkson and Kench, 1956). There has been dis-
agreement with respect to the effects on glucose resorption, and perhaps in
man there is little reduction at diuretic doses, but Vander (1963) has shown
a very definite inhibition in the dog, A Tm being around — 100 during max-
imal diuresis. Mercaptomerin in dogs lowers the bicarbonate threshold of
the proximal tubules by 35% without significant effect on the distal tubules
EFFECTS ON THE KIDNEY 921
(Gardier and Woodbury, 1955). Since the resorption of bicarbonate is about
equal in the proximal and distal segments, this indicates an exclusively
proximal action. Increased excretion of Ca++ and Mg++ in both man and
dog treated with mercurials has been reported, and it is likely that the site
is proximal (Wesson, 1962). The inhibition of the secretion of p-aminohip-
purate, tetraethylammonium ion, and phenol red has been shown not only
in intact animals but with low concentrations of the mercurials in isolated
tubules or slices (Forster and Taggart, 1950; Farah and Rennick, 1956;
Koishi, 1959 b). Thus 0.01 mil/ Hg++ completely blocks phenol red trans-
port in the flounder tubule. These and other observations all point definitely
to a major site of action in the proximal segment of the nephron. However,
there is also evidence that more distal transports can be affected. For exam-
ple, the secretion of K+ and the H+ and NH4+ exchanges in the distal seg-
ment (Dale and Sanderson, 1954), and the resorption of solute-free water
by the loop of Henle (Lambie and Robson, 1960) and the distal segment
(Goldstein et al., 1961), are depressed by the mercurials. It is difficult to
compare the actions on proximal and distal portions of the nephron be-
cause of the different magnitudes of the transport processes; i.e., effects
on proximal transport would be much more marked because of the major
role of this segment in resorption.
(B) Disappearance of renal SH grovps. Histochemical determination of
the free SH groups in different regions of the kidney in normal and mer-
curial-treated animals might provide some information on the site of ac-
tion if clear-cut differences are observed. Cafruny et al. (1955 b) determined
the free protein SH groups in rat kidney sections by treatment with the SH
reagent DDD (2,2'-dihydroxy-6,6'-dinaphthyldisulfide), coupling of the
naphthol moiety with the azo dye Fast Blue RR, and photometric analysis.
Following injection of a large dose of mersalyl (20 mg/kg), reduction of SH
groups was observed in all portions of the nephron except the proximal and
distal convoluted portions (see accompanying tabulation). Even at the
markedly nephrotoxic dose of 40 mg/kg there is no decrease in SH groups
Extinction
values
% Change
Control
Mersalyl
Proximal tubules (convoluted)
0.619
0.613
- 1
Proximal tubules (terminal)
0.415
0.227
-45
Brush borders (terminal)
0.701
0.545
-22
Loop of Henle (descending)
0.355
0.238
-33
Loop of Henle (ascending)
0.373
0.229
-39
Distal tubules (convoluted)
0.592
0.596
+ 1
Collecting duct (medullary)
0.279
0.136
-51
922 7. MERCURIALS
in the convoluted segments. With low doses (2.5 mg/kg), disappearance of
SH groups was observed only in the terminal portion of the proximal tu-
bules and the ascending loops, the latter being the most sensitive region
of the nephron. Time studies showed that the terminal proximal tubules
are affected first and up to 1 hr show more reduction than the loops of
Henle. Incubation of kidney sections with 20 mM mersalyl (Cafruny et al.,
1955 b) or saturated HgClg (Cafruny et al., 1955 a) produces marked non-
specific reduction in free SH groups, indicating that the pattern seen in the
whole animal is due in part to the factors involved in the resorption and
secretion of the mercurials. Cafnmy and Farah (1956) later used dogs so
that correlation between diuresis and SH group disappearance might be
made. Kidneys were removed at the peak of diuresis (around 90 min) from
10 mg/kg of mersalyl, urine flow and Na+ being increased 5- to 6-fold, and
the changes given in the accompanying tabulation were observed, indicat-
Extinction
. values
0/
/o
Change
Control
Mersalyl
Proximal tubules (convoluted)
0.537
0.541
+ 1
Proximal tubules (terminal)
0.410
0.267
-35
Loop of Henle (ascending)
Distal tubules
0.360
0.495
0.261
0.510
-28
+ 3
Collecting ducts
0.242
0.192
-21
ing that selective reaction with SH groups in certain regions of the kidney
does occur. It should be pointed out that the nature of these SH groups is
not known; they may be on enzymes, carriers, or nonfunctional proteins.
Farah and Kruse (1960) used seven mercurials at 4 mg Hg/kg in rats
and found moderate reduction of the protein SH groups (around 20-30%)
in the terminal proximal tubules, the loops of Henle, and the collecting
ducts, and it was concluded that maximal diuresis occurs when 20% of the
protein SH groups of the proximal tubule are reacted, and thus that no
more than this can be related to the diuresis. However, there is no correla-
tion between diuresis and decrease in the SH groups, since p-MB and MM,
both nondiuretic, produce similar changes in these groups. HgClg and mer-
salyl at equimolar doses cause comparable decreases in renal SH groups in
rats, and this was noted particularly at the bases of the proximal tubular
cells (Gayer and Partowi, 1962). Renal S — S groups do not change for sev-
eral hours after injections of HgClg or chlormerodrin, but from 6 to 24 hr
there is a,n increase in S — S groups at the expense of SH groups (Shore and
Shore, 1962). This may be related to the potent inhibition of protein disul-
fide reductase, but is probably not correlated with diuresis since it occurs
EFFECTS ON THE KIDNEY 923
some time after maximal urine flow. Renal damage and conversion of SH
to S — S groups could be related in some as yet unexplained way.
These results all demonstrate that mercurials react with renal SH groups,
and that some selectivity on certain regions may be exerted, but do not
necessarily have any bearing on the site of transport inhibition, since the
SH groups involved in the transport (assuming they are) may be only a
very small fraction of the total in the tissue; indeed, it is quite possible that
only 1-2% of the total SH groups need be reacted to produce maximal
diuresis.
(C) Reduction of electrical potentials of tubular cells. There are two elec-
trical potentials of the proximal tubular cells of the isolated Necturus neph-
ron, a transmembrane potential of —72 mv and a transtubular potential
of —20 mv (lumen negative) (Giebisch, 1958, 1960, 1961). Chlormerodrin
in a concentration around 220 //g Hg/g tissue reduces both potentials; in
the perfused nephrons the transmembrane potential is decreased 62% and
the transtubular potential 63%. Since these potentials are dependent on
active ion transports, quite possibly they relate to renal function. These
results show that mercurials can affect the proximal tubules, but whether
this is related to the diuretic effect is impossible to say.
(D) Pattern of accumulation of mercurials in the kidney. The kidneys of
rats poisoned with HgClj (3 mg/kg intraperitoneally) were examined from
5 min to 48 hr afterward by the silver sulfide method, and mercury was
found to be deposited first in the endothelial cells of the interstitial capilla-
ries, then in the glomerular tufts, and eventually in the epithelium of the
proximal tubules, beginning apically and progressing toward the bases of
the cells (Wockel et al., 1961). The mercury in the proximal ceUs is partic-
ularly associated with the basally situated mitochondria. It was conclud-
ed that Hg++ is filtered through the glomerulus and picked up by the tu-
bular cells during resorption, which is the most obvious route for Hg++
and one which explains the early and marked effects on the proximal tu-
bule. However, it has recently been claimed that another route is more
important. Brun et al. (1947) suggested that mersalyl is secreted by the tu-
bular cells, and that this accounts for the high concentration of mercury in
the tubules and the selective effects on proximal transport. It was claimed
by Borghgraef et al. (1956) that the excretory rate of chlormerodrin is
too fast for glomerular filtration, especially considering that a large frac-
tion of the plasma mercurial is bound and not filtered, and that tubular
secretion is responsible for essentially all the mercury in the tubules. This
theory has also been proposed by Weiner et al. (1956), Kessler et al. (1957
a, b), and Campbell (1959). Greif (1960) held that the uptake of Hg^o^
by Phascolosoma nephridia is an active transport, presumably because it is
inhibited by cyanide; however, no inhibition by '2,4-dinitrophenol, azide,
or iodoacetate was noted. Despite the evidence for the tubular secretion of
924 7. MERCURIALS
mercurials, I am not convinced that it is more important than resorption
from the glomerular filtrate. If 1-5% of the total plasma mercurial is in a
freely diffusible form, this fraction will certainly be filtered readily and
concentrated in the lumen; renal accumulation occurs with many drugs
which are bound to the plasma proteins. Second, only the free mercurial is
available to the tubular cells from the peritubular fluid, and it is difiicult
to envision a mechanism by which the cells can pick up or actively secrete
protein-bound mercurial. The tubular cells undoubtedly accumulate mer-
curials from the plasma as do other tissues, and may secrete them to some
extent, but the rates of excretion are not such as to imply secretion as the
major pathway. It is also strange that mecurials of so many different struc-
tures would be actively secreted; those with carboxylate groups might be
carried by the transport system for acids, as Campbell (1959) postulated
for mercaptomerin, but neither probenecid nor p-aminohippurate interferes
with the excretion of mercurials in the dog (Kessler et al., 1957 b). In any
event, there is little likelihood that the pattern of mercurial distribution in
the kidney can be directly correlated with the site of action. Weiner et al.
(1959) emphasized that there is no obvious relationship between diuresis
and the total amount of mercurial in the kidney or its parts, and stated,
"Diuresis is a consequence of the action on a specific renal receptor by a
very small amount of mercury."
(E) Stop-flow technique. Serial sampling of urine following ureteral clamp-
ing allows an analysis of the composition changes throughout the nephron,
and such studies have uniformly pointed to a proximal rather than a distal
site of mercurial action (Kessler et al., 1958; Vander et al., 1958). This ap-
plies exclusively to the site of inhibition of Na+ resorption and diuresis.
(F) Differential damage to renal cells. It has been thought that those por-
tions of the nephron most readily damaged by toxic doses of the mercurials
might be the same portions primarily affected to produce diuresis. Suzuoki
(1912) was the first to show by adequate methods that mercurials can dam-
age rather selectively the more terminal portions of the proximal tubules.
Edwards (1942), on the other hand, claimed that Hg++ exerts selective
damage on the central region of the proximal convoluted tubule, injury
to the distal convolution being rarely seen. The loops of Henle are too thin
and squamous to permit satisfactory examination. Simonds and Hepler
(1945) confirmed the observations of Suzuoki in finding selective damage
to terminal proximal tubules. More recent work has not greatly extended
our knowledge. Relatively little damage to the glomeruli has been confirm-
ed (Staemmler, 1956) even when a severe nephrosis is produced, although
Schorcher and Loblich (1960) found some changes in the glomerular filtra-
tion membrane by electron microscopy. Tubular cells show apical edema
and vacuolization, these occurring primarily in the proximal segment in
rats injected with meralluride (Sanabria, 1963). The brush border may show
EFFECTS ON THE KIDNEY 925
a separation of the villi, and mitochondrial disintegration occurs. With
large doses, damage may be observed with the electron microscope within
10 min. The maximal diuretic dose of mersalyl (6 mg/kg) in the rabbit
produces nuclear pycnosis, mitochondrial changes, and vacuolization in the
convoluted tubule (Dejung, 1963). Again, these results show selective ef-
fects, but may be the result of differential distribution and may not relate
to the diuretic site of action.
The evidence taken together suggests that various portions of the nephron
are affected in one way or another by the mercurials, and what portion may
be involved will depend on the particular transport considered. The diuretic
action, i.e., the inhibition of Na+ resorption, appears to be limited mainly
to the proximal tubule; whether this is primarily in the convoluted or ter-
minal segments is not known. Analyses of th6 filtrate composition through-
out the nephron in the presence of mercurials are very difficult to interpret,
but most of the data are compatible with a proximal action (Welt et al.,
1953). More indirect evidence will be provided by studies on enzyme inhi-
bition in the following section, but it is clear that the basic mechanism of
mercurial action must be sought in the Na+ transport system of the prox-
imal tubules.
Effects on Enzyme Activity in the Kidneys
Much of the work has unfortunately been on succinate dehydrogenase,
presumably because the activity is easily measured, but this enzyme is
probably not directly involved in renal transport and, in fact, is much less
sensitive to the mercurials than are many other enzymes. The results, how-
ever, may be taken as a rough indication that the mercurials can in diuretic
doses inhibit various renal enzymes, and to some extent provide evidence
for the primary site of action. Handley and Lavik (1950) found that mer-
alluride injected intravenously at a dosage of 8 mg Hg/kg in dogs and rats
reduces the succinate dehydrogenase activity around 45% in the kidney
at the peak of diuresis, whereas no inhibitions were observed in the liver
or heart. Fawaz and Fawaz (1951), on the other hand, could detect no
changes in succinate oxidation by renal homogenates from rats given the
diuretic dose (4 mg Hg/kg) of mersalyl, and concluded that if the mercurial
is acting on an SH enzyme, succinate dehydrogenase is not involved. Mus-
takallio and Telkka (1953) reported that high doses (10-60 mg Hg/kg) of
mercurophylline lead to loss of succinate dehydrogenase activity in the distal
tubules, using a tetrazolium staining technique, and later (Telkka and Mus-
takallio, 1954) found some inhibition in the proximal tubules and loops of
Henle, their conclusion being that there is little correlation between such in-
hibition and transport inhibition. Somewhat different results were obtained
by Rennels and Ruskin (1954), who found marked inhibition of succinate
dehydrogenase in the proximal tubules of the rat following 40 mg Hg/kg
of meralluride, little or no effect being exerted on the distal tubules or loops
926 7. MERCURIALS
of Henle. However, this is a very high toxic dose and the maximal inhibi-
tion occurred at 24-48 hr, although some effect could be observed at 3 hr.
In doses under 10 mg Hg/kg, no inhibition could be demonstrated, so that
again there is no reason to relate the inhibition to diuresis. Wachstein and
Meisel (1954) also observed succinate dehydrogenase inhibition in the prox-
imal tubules of the rat following large nephrotoxic doses of meralluride.
Bahn and Longley (1956) confirmed these results in that diuretic doses of
meralluride (4 mg Hg/kg) produce insignificant inhibition of succinate de-
hydrogenase while toxic doses (12 mg Hg/kg) inhibit moderately, especially
in the terminal proximal tubules. Finally, Bickers et al. (1960) found that
meralluride at 3-4 mg Hg/kg does not inhibit succinate dehydrogenase,
whereas 5 mg Hg/kg does to some extent after 10 hr. Later changes in en-
zyme activity, at a period of tubular damage, may be the result of secondary
changes and necrosis, so have little bearing on the mechanism of the diuretic
action.
a-Ketoglutarate oxidase is more sensitive to mercurials than is succinate
dehydrogenase, and is inhibited 64% at 3-4 hr and 91% at 6 hr in rats
given HgClg at 3 mg Hg/kg (Shore and Shore, 1960). In animals fed sucrose,
the inhibitions are less, and sucrose also protects somewhat against the
nephrotoxic effects of Hg++, the mechanism being unknown. But again it
is impossible to correlate this action with the diuretic effect, since mersalyl
in diuretic dose (4 mg Hg/kg) does not alter the renal concentration of
a-ketoglutarate, although toxic doses (10 mg Hg/kg) produce an early and
prolonged rise in a-ketoglutarate (Dziirik and Krajci-Lazary, 1962). No
changes in sorbitol dehydrogenase activity in the kidneys of rats given
nephrotoxic doses of mercurin are detectable, and it would probably not
be very significant if they were (Pietschmann et al., 1962). Moderate inhibi-
tions of NAD and NADP diaphorases are found after toxic doses of the
mercurials (Bickers et al., 1960; Wachstein and Meisel, 1957), but in all
cases where enzyme inhibition is seen, there is histologically demonstrable
damage to the tubules. Protein disulfide reductase is apparently rather
potently inhibited in the kidney during mercurial action, but this is un-
doubtedly unrelated to the diuretic effect (Shore and Shore, 1962).
Various phosphatases have been the subject of investigation although
there is little reason to expect a relation to ion transport. Nephrotoxic
doses of HgClg inhibit renal phosphatase slightly (10-20%) without affect-
ing serum phosphatase, but subtoxic doses somewhat increase the phospha-
tase activity (Hepler et al., 1945). There is no correlation between glucosuria
and phosphatase inhibition (Hepler and Simonds, 1946). Very high doses
(175-260 mg/kg) of HgClg decrease tubular phosphatase, although not in
capillaries or glomeruli, but low doses (6-10 mg/kg), which are lethal in 3
weeks, do not alter the phosphatase (Sachs and Dulskas, 1956). Diuretic
doses of various mercurials do not inhibit alkaline phosphatase, |5-glycero-
EFFECTS ON THE KIDNEY 927
phosphatase, or glucose-6-phosphatase (Shore et al., 1959; Bickers et al.,
1960), but toxic doses lead to a diffuse distribution of various phosphatass
in the tubular cells, and some inactivation (Wachstein and Meisel, 1957).
Very little work has been done on ATPase, but there is shght evidence that
it can be inhibited in the kidney by both mersalyl (DeGroot et al., 1955)
and p-MB (Padykula and Herman, 1955). It is probably fair to say that
none of these investigations of phosphatases has established a relation to
their role in transport or mercurial diuresis.
Effects on Renal Metabolism
Mercurials can certainly depress the respiration of kidney at fiufficiently
high doses, and this may possibly be a factor in the renal damage produced,
but there is little evidence that nontoxic diuretic doses of the mercurials
reduce Og consumption. The early work of Gremels (1929) indeed showed
that, in the heart-lung-kidney preparation, mersalyl actuaUy increases renal
respiration during diuresis. Cohen (1953 a,b) showed particularly well the
difference between diuretic and toxic doses. Mercaptomerin at a dosage of
10.7 mg Hg/kg in rats produces a marked diuresis. After 1 hr the animals
were sacrificed and a mitochondrial suspension prepared from the kidneys;
no change in the Og uptake was noted, and the P : 0 ratio may actually
have been increased. However, with a toxic dose of 26.7 mg Hg/kg, the O2
uptake was depressed 35% and the P:0 ratio dropped from 0.55 to 0.092.
Very similar effects were observed on kidney slices, only toxic doses reduc-
ing the respiration. Even toxic doses do not affect the O2 uptake or P:0
ratio of liver mitochondria obtained from poisoned animals. That mercurials
exert very little effect on the cycle when given in diuretic doses was demon-
strated by Fawaz and Fawaz (1954). Rats were poisoned with fluoroacetate,
and citrate accumulation was determined in the heart and kidney of control
animals and those given mersalyl at 4-5 mg Hg/kg; no significant change
in citrate formed was noted, so that the operation of the cycle would seem
to be unaltered. Although these results indicate no appreciable interference
with coenzyme A, Leuschner et al. (1957) claimed that the coenzyme A
induced acetylation of sulfanilamide is reduced by HgCla, mersalyl, and
mercaptomerin in the same order of potency as for diuresis. Toxic doses
reduce the coenzyme A reaction, but it is not certain if diuretic doses are
able to do this. The mercurial diuresis is less in coenzyme A-deficient rats,
but the significance of this is unknown. Eesults on oxidative phosphoryla-
tion are contradictory. Greif and Jacobs (1958) found that even large doses
of chlormerodrin (up to 20 mg Hg/kg, which is 20 times the diuretic dose)
do not alter the P : 0 ratio of kidney mitochondria with glutamate as the
substrate, but Shore and Shore (1960) reported a marked fall in P:0 ratio
with a-ketoglutarate as the substrate following toxic doses (3 mg Hg/kg)
of HgCl^.
928 7. MERCURIALS
The results obtained on kidney slices are generally in agreement that,
although respiration may be reduced by reasonably high concentrations
of mercurials (0.25-1 mM), the inhibition of various transports is much
greater. Thus Cross and Taggart (1950) found that 1 mM Hg++ depresses Og
uptake of rabbit kidney slices 35% while inhibiting p-aminohippurate ac-
cumulation 89%, and Mudge (1951) showed that respiration is scarcely af-
fected by Hg++ at concentrations markedly altering Na+ and K+ transport.
Mendelsohn (1955) confirmed that Hg++ can reduce p-aminohippurate ac-
cumulation as much as 70% without affecting respiration in kidney slices.
Kobinson (1956) believed that the inhibition of respiration by mercapto-
merin might be the basis for the swelling of rat kidney slices and the inter-
ference with water transport, but there was no direct evidence for a relation-
ship. Maizels and Remington (1958) also observed moderate respiratory
inhibition with mercaptomerin and meralluride, but did not feel that this
was the chief factor in the increase in water and Na+ of the slices. Further-
more, the lowest concentration of the mercurials which exerts an effect in
vitro is much greater than the maximal tolerated plasma concentration in
rats in vivo, so it is doubtful if these inhibitions of respiration are relevant
to the diuretic action.
Summarizing all the data obtained on enzyme and metabolic inhibition
in the kidney, it is disappointing that no system particularly sensitive to
the mercurials has been found, and that no correlation between inhition
and transport processes has been demonstrated. If the basis of mercurial
diuresis is metabolic, no clear evidence for this has yet been provided.
Accumulation and Excretion of Mercurials by the Kidneys
All mercurials are rather slowly accumulated by the kidneys and the high
levels are sustained for periods of several days. The kidney/plasma ratio is
maximally around 1000 in the rat and dog when chlormerodrin is given
(Borghgraef and Pitts, 1956; Giebisch and Dorman, 1958), but these ratios
are reached only after many hours, and are in part due to the retention by
the kidney with falling plasma levels. The correlation between distribution
and excretion of a mercurial and the diuresis is well shown in the results of
Borghgraef et al. (1956) (Fig. 7-44). The loss of mercurial from the plasma
is divided into three components: that excreted in the urine, that entering
the various tissues, and that accumulated by the kidneys. Some mercurials
are excreted fairly rapidly and others slowly; meralluride administered in-
tramuscularly in man is 50% excreted in 3 hr (Burch et al., 1950) but
chlormerodrin given by the same route to rats is only 21% excreted after
24 hr, 67% remaining in the kidneys (Borghgraef and Pitts, 1956). We have
discussed the theories of the excretory mechanisms (page 923) and the
possible role of tubular secretion. The mercurials are not excreted entirely
in the form administered. Some may be split into inorganic Hg++ but most
EFFECTS ON THE KIDNEY
929
is excreted as a complex with cysteine, or other thiols (Weiner and Miiller,
1955). The origin of this cysteine complex is not known, and it may be in
the kidney or the blood. The various mercurials are distributed differently
throughout the tissues, as one might expect from the different properties of
their molecules, and this must play some role in the effects they produce,
lOOfc -w
Fig. 7-44. Distribution of Hg^"^ after intravenous injection of
chlormerodrin in dogs at 1 mg Hg/kg. The curve shows the
change in urine flow (diuresis) estimated from the figures given.
(From Borghgraef et al., 1956.)
not only on the kidney but on other tissues in higher doses. The most
thorough study has been made by Kessler et al. (1957 a) and some of their
results are summarized in Table 7-19. Hg++ behaves quite differently than
the organic mercurials; it does not enter the kidney rapidly but eventually
reaches very high levels after several hours. It may be noted that the dis-
tribution of p-MB is not markedly different from the diuretic mercurials.
Little is known about the cellular fractions of the kidney which accumulate
the mercurials, but it is somewhat surprising that Greif et al. (1956) found
that by far the most mercury after injection of chlormerodrin to rats, fol-
lowed by fractionation of kidney homogenates in sucrose solutions, is in
the soluble fraction, about one third the amount in granules, and much
less in the nuclei. Although all of these distribution studies are important
in understanding many facets of mercurial action, they do not appreciably
contribute to our knowledge of where or how the mercurials produce dis-
turbances in the renal function.
930
7. MERCURIALS
Table 7-19
Concentration of Mercury in Tissues of the Dog After Intravenous
Administration of Mercurials "
Tissue
concentration
(//g/g wet
weight)
Tissue
Chlormerodrin
Meralluride
Mersalyl
HgCl,
p-MB
Renal cortex
155
19
7.7
113
36
Renal medulla''
125
13
4.0
79
—
Renal papilla
4.5
1.4
0.4
2.3
—
Liver
3.9
0.8
0.8
6.6
2.8
Heart
0.3
0.2
0.4
0.5
0.7
Spleen
1.4
0.6
1.1
74
2.1
Lung
1.2
0.9
0.9
2.0
2.8
Diaphragm
0.2
0.2
0.1
0.3
0.8
Intestine
0.7
0.5
0.7
1.1
2.5
Skin
0.4
1.0
0.7
0.8
1.0
Plasma
1.1
1.3
1.2
2.9
—
" At 2 mg Hg/kg and sacrifice of the animals at 160 niin. (From Kessler et nl.,
1957 a; p-MB data from Kessler et ah, 1957 b.)
'' Only outermost sections of medulla considered here.
Active Form of the Mercurials and Relation of Action to Structure
The concept that the organic mercurials in order to be active diuretics
must dissociate into inorganic mercury is an old one and has been revived
recently to explain some of the differences between mercurials and the ef-
fects of pH on the activity. Most diuretic mercurials have the structure:
OCH3
I
R— C H2— CH— CH2— Hg+
The methoxy group arises because these mercurials are synthesized by the
oxymercuration of alkenes in methanol; the nature of this group is not par-
ticularly important for the activity. It is possible that the reverse of this
reaction
OCH3
1
R— CH2— CH— CH2— Hg+ + H+ -► R— CH2— CH = CHj + CH3OH + Hg++
might occur under physiological conditions, as suggested by Hughes (1957).
EFFECTS ON THE KIDNEY
931
Such a splitting would occur more rapidly the lower the pH. Certainly this
release of Hg++ is not generally responsible for the actions and toxicity of
the organic mercurials, and most of the mercurials, such as p-MB, PM,
and MM, are quite stable. Hepp (1887) emphasized long ago that alkyl
mercurials do not release Hg++ in the body and exert a toxic action much
different than Hg++. However, the diuretic mercurials present a different
situation and the theory of Hg++ release must be given serious consideration.
The possibility that the above reaction might be catalyzed or accelerated by
thiols through the formation of mercaptides was presented by Benesch and
Benesch (1952) as a result of their polarographic investigations of the reac-
tion between mersalyl and dimercaprol. In this scheme, free Hg++ may not
be produced directly; instead, a cyclic mercaptide is formed, which could
OCH,
I
-S-Hg— CHg— CH— R
hS-Hg— CH2— CH— R
OCH,
-s.
Hg
hS
/
CH,= CH — R
CH,OH
OCH3
I +
R— CH— CHj— Hg
conceivably be the inhibiting complex in renal transport, although it is also
possible that monothiols can act similarly:
OCH3
— S— Hg— CH2— CH— R + H^
-S— Hg+ + CH2=CH— R + CH3OH
since Mudge and Weiner (1958) showed that cysteine increases the split-
ting of mersalyl in acid medium. Other acid-stimulated types of splitting
would be the simple reactions:
\\ //
Hg
v\ //
Hg
R— CH,— Hg + H
R-CH, - Hg*
but these usually occur fairly slowly, especially for the alkyl mercurials.
932 7. MERCURIALS
The hypothesis that organic mercurial diuretics to be active must release
inorganic Hg++ in the kidney was proposed by Mudge and Weiner (1958)
and the evidence was presented by Weiner et al. (1962). This had been
suggested occasionally ever since the first use of the organic mercurials but
very little evidence either for or against was reported, and the idea generally
was not taken seriously because all the other metal compounds used clini-
cally had been shown to act directly without splitting off the metal ion. The
evidence now accumulated impels one to consider this possibility. If such
splitting occurs, it is important not only for the diuretic action but for many
other effects of the mercurials, even in vitro. The evidence is mainly of two
sorts: (1) a correlation between acid lability of organic mercurials and their
diuretic activity, and (2) the potentiation of diuretic activity by the acid-
ifying NH^Cl.
Mudge and Weiner (1958) pointed out that mersalyl does not split* in
acid medium over 3 hr, but in the presence of cysteine the half-time for
splitting is 105 min and with dimercaprol 5 min. The acid lability of 32
mercurials was tested by Weiner et al. (1962) by incubating the mercurial
at 1 WlM with cysteine at 2 mM in an Og-free medium at pH 4 and 37°
for 3 hr. The diuretic activity was expressed as A CI (//moles/min/kg). All
of the 22 mercurials which are diuretic are acid-labile, while of the 9 non-
diuretic mercurials 6 are stable and 3 labile (1 mercurial is indeterminate
in diuretic activity). There is thus a reasonably good correlation between
lability and diuretic activity. The 3 labile nondiuretic mercurials are all
of the ether series with structures of the type R — CHg — 0 — CH2CH2 — Hg+,
and possibly their distribution is such that Hg++ is not released in the
proper region. The pH dependence of the splitting, according to the reac-
tions of Benesch and Benesch (1952), indicate the rate of splitting to be
approximately one thousandth as fast at pH 7 as at pH 4. If one assumes
that X = XqC'^', where X is the amount of organic mercurial, it may be
calculated that at pH 7 around 0.14% splitting would occur in 3 hr, since
the mean splitting of the labile mercurials is about 75%. Since maximal
diuresis occurs in 1-2 hr, approximately 0.1% would be split in this time.
Now, this calculation is not very valid because one does not know the con-
ditions for splitting in the kidney; e.g., dimercaptides may be formed there
and split more rapidly than the cysteine complexes. Diuretic activity was
examined by injecting the mercurials with a 10-fold excess of cysteine, but
presumably in the kidney there would be a transfer of the mercurial from
cysteine to other thiols. Thus it is difficult to obtain an idea of the amount
of inorganic mercury which is released in the kidney. If much splitting oc-
curs, one might expect to find Hg++ excreted in the urine in some form.
* The term "split" will be used to designate the dissociation of the mercurial into
inorganic mercury so that there will be no confusion with the term "dissociation"
which is used to indicate the reaction R — Hg — X ±^ R — Hg+ + X~.
EFFECTS ON THE KIDNEY 933
Moyer et al. (1957) and Handley and Seibert (1956) could detect no inor-
ganic mercury after administration of meralluride, but Weiner et al. (1962)
pointed out that Hg++ would be excreted mainly as the cysteine complex
and this would be included with the organic mercurial in their chromato-
graphic fractionations. It is also possible that very tight binding of the ac-
tive Hg++ in the kidney would occur, so that the excretion would be slow.
Weiner et al. (1962) could detect Hg-cysteine in the urine following injec-
tion of 3 particularly labile mercurials, but of course this is not valid evi-
dence that the Hg++ is the active form.
Clinical diuretic refractoriness to the mercurials has been known for years
and it is often possible to restore the diuretic response by giving NH4CI.
The potentiating action of NH4CI has been the subject of much work and
speculation, but the mechanism is still unknown. One hypothesis is that
the urinary acidification is the major factor. Weiner et al. (1962) assume
that this acidification increases the splitting of the labile mercurials. Ad-
ministration of NH4CI drops the pH of the urine below 5 and an optimal
effect is usually seen around 4.5; thus the splitting of the mercurial in the
urine will be significantly accelerated. However, the Hg++ in the urine wiU
presumably be complexed with cysteine or other simple thiols and the rate
of splitting will not be very great. It has also generally been assumed that
the splitting occurs in the tubular cells. Although the intracellular pH un-
doubtedly falls after NH4CI, the decrease is certainly not as great as in
the urine. Pending determinations of intracellular changes, one cannot esti-
mate the effect this would have on mercurial splitting. The mercury con-
tents of the proximal tubules in the dog were determined histochemically
by Cafruny (1962), using di-/?-naphthylthiocarbazone, and acidosis was
shown to increase the levels for chlormerodrin and Hg++, although a de-
crease occurs with p-MB. He felt that acidosis either increases the available
receptors for mercurials or somehow alters the affinity of the mercurial for
the receptors. Change in the acid-base balance does not alter the excretion
of the mercurials, so it is presumably not a matter of the tubular concen-
tration of mercurial. If the fall in pH is responsible for increased splitting
of the mercurials and thus a greater diuretic effect, NH4CI administration
or other acid-base changes should not affect the diuresis produced by Hg++
complexes. Mudge and Weiner (1958) and Levy et al. (1958) reported that
the action of meralluride is altered more than Hg-cysteine by variations in
the urinary pH. However, Hg-cysteine diuresis is increased 2.4-fold in go-
ing from alkalosis to acidosis, so that the results are not as clear-cut as one
might wish. If acidosis is responsible for greater mercurial action, one
would also expect that any type of acidosis would be effective. However,
Kessler (1960) points out that inhalation of 12% CO., actually decreases
mercurial diuresis, although not as much as alkalosis produced by HCOg"
infusion. But inhalation of 12% CO2, although it produces an acidification
934
7. MERCURIALS
of the plasma (pH 7.4 to 7.14), does not alter urinary pH significantly, as
does administration of NH4CI; no one knows what happens to intracellular
pH in the tubules. It may also be pointed out that alkalinization of the
urine with acetazolamide or K+ does not alter the diuretic response to mer-
curials (Pitts, 1958).
It is not immediately apparent why Hg++ must be formed from organic
mercurials to inhibit renal transport, since in most cases the organic mer-
curials react readily with SH groups which may be involved. If mercaptides
or dimercaptides participate in the splitting of the mercurials, the Hg++
formed must dissociate from these SH groups and attach to others, because
the cell component originally binding the mercurial must be blocked and
there would be no necessity for splitting. If Hg++ is necessary for diuresis,
it must be that either (1) a cyclic mercaptide is required, or (2) the impor-
tant SH groups are not sterically accessible to the larger organic mercur-
ials. Weiner et al. (1962) assume that the specific receptor for the diuretic
action contains two groups, one being an SH group and the other either an
SH group or some other ligand complexing with Hg++ (e.g., an amino
group). The complete scheme as outlined by Weiner et al. (1962) is shown
NONSPECIFIC
PROTEIN
R-Hg-S-PROTEIN
4=
R-Hg-S-CYST ■
CYST-S-Hg-S-CYSTj
R-Hg-S-CYST
iCYST-S-Hg-S-CYST
SPECIFIC
RECEPTOR
Fig. 7-45. Scheme of mercurial re-
actions in the kidney. (From Weiner
et al., 1962.)
in Fig. 7-45. It is strange that this transport component would not be inhib-
ited by organic mercurials bound to one SH group, or that certain potent
SH reagents, such as MM or p-MB, would not inactivate it. The nondiuretic
p-MB prevents and reverses the diuretic effects of the mercurials, while
MM does not do this (Miller and Farah, 1962 a). A competition between
p-MB and Hg++ for the receptor SH groups was suggested. Miller and Farah
EFFECTS ON THE KIDNEY 935
(1962 b) also postulate a two-group receptor, one group being SH; mer-
curials which are diuretics attach to the SH group, split, and the resulting
Hg++ makes a two-point attachment. The block by p-MB is due to its
binding to the SH group; being stable it does not split. If this is so, p-MB
might be expected to displace mercurials in the kidney, and this was dem-
onstrated using Hg^^^-labeled chlormerodrin. The decrease in radioactivity
of the kidneys parallels the antagonism of the diuresis by p-MB.
Another hypothesis for diuretic mechanism was outlined by Kessler et
al. (1957 b), who assumed that the organic mercurials act as intact molecules
by a two-point attachment to a receptor. The basic structure for diuretic
activity was given as R — C — C — C — Hg+, i.e., a hydrophilic group sepa-
rated from the Hg by three carbon atoms, the R group interacting in some
manner with a group spaced appropriately in relation to an SH group. This
hypothesis in its simple form has had to be abandoned in the light of fur-
ther work showing that various mercurials not conforming to this structure
are diuretic, e.g., some of the aryl mercurials (Weiner et al., 1962). How-
ever, the idea that there is some relationship between structure and diuretic
action should not be given up, inasmuch as the situation may be more com-
plex than originally assumed. If one considers the three simple alkyl mer-
curials (see accompanying tabulation), one sees that diuretic activity is
Alky] mercurial Diuretic activity Lability
CH3— CH2— Hg+ - -
HO— CH2— CH2— Hg+ + +
HO— CH2— CH2— CHj— Hg+ - -
correlated with acid lability, and that HO— CH2CH2CH2— Hg+, which
should be diuretic in the scheme of Kessler et al., is not. It is intriguing
that HO— CH2CH2— Hg+ is 95% split under the conditions of the lability
test, whereas the other two compounds are completely stable; it is also
surprismg that HO^— Hg+ and HgN— <^— Hg+ are 73% and 88% split,
respectively, while "OOC — (f — Hg+ and (f — Hg+ are not split at all. The
structural requirements of lability seem to be very rigid. Could it be that
the structural requirements for splitting are the same as for combination
with a receptor to produce diuresis, splitting not being a necessary prelude
to an effect? It is clear that a final decision as to these important matters
cannot be reached at this time and that more data must be accumulated;
one would like to have some information on the rates of splitting of mer-
curials in homogenates or extracts of kidney, and the effects of pH on this.
936 7. MERCURIALS
Mechanism of Transport Inhibition
If one knows essentially nothing of the cellular site of action of the mer-
curials, and is completely ignorant of the molecular nature of ion transport,
it is difficult to discuss possible mechanisms of inhibition without becoming
ethereal. All the evidence points to a lack of significant depression of the
exergonic phases of renal metabolism at concentrations markedly affecting
transport, so it is likely that the action is on some component of the func-
tional system. If the postulated specific receptor for mercurials is a carrier,
then there is the problem of accounting for the inhibition of many types
of transport; furthermore, it is not at all certain that a carrier is involved
in ion transport. On the basis of what was said in the previous section on
the effects of mercurials on membranes and permeabilities, it is most likely
that the site of action is the tubular cell membrane. The Na+ pump is prob-
ably located in the peritubular membrane and the diffusion of Na+ across
the lumenal membrane is passive along concentration and electrical gra-
dients. The fact that the transmembrane potential is around — 43 mv at
the lumenal surface and — 64 mv at the peritubular surface was interpreted
by Giebisch (1960) as indicating the greater Na+ permeability of the lu-
menal membrane. Mercurials could thus depress Na+ resorption by acting
in three ways: (1) inhibition of the Na+ pump, (2) decrease of the permeabi-
hty to Na+ of the lumenal membrane, or (3) increase of the permeability
to Na+ of the peritubular membrane. It has frequently been assumed that
the mercurials inhibit active transport directly but, as has been discussed
for other systems, it is possible that the primary effect is on the membrane
structure controlling permeability. The evidence from the changes in elec-
trical potentials brought about by chlormerodrin shows that both mem-
branes are affected (Giebisch, 1961). The potential across the peritubular
membrane is decreased to —25 mv and, since the transtubular potential
is simultaneously decreased to — 7 mv, it would seem that the lumenal
membrane potential is decreased to —18 mv. These changes can be in-
terpreted as due to increases in the Na+ permeability of both membranes,
but it is also possible to conclude that there is a general decrease in the
ionic permeability. Until information on the effects of mercurials on ion
fluxes is available, one cannot distinguish between these possibilities. The
results of Mudge (1951) on rabbit kidney slices point either to an increase
in permeability to Na+ and K+, or to an inhibition of active transport.
However, the relationship of these in vitro results to mercurial diuresis is
obscure, especially as Auditore and Holland (1956) found that diuresis can
be produced without appreciable loss of cell K+, although the latter can
occur with minimal toxic doses. That mercurials can alter ion permeabilities
without appreciably depressing active transport is demonstrated by studies
on other tissues, such as atria (page 945). Mercurials could conceivably
alter pore sizes by distorting membrane structure, or actually clog ion-
EFFECTS ON TISSUE FUNCTIONS 937
transporting pores by reacting with SH groups on the walls, or interfere
with the open-closed transition postulated to occur in the membrane by
Kavanau (1963). White et al. (1961) have provided the only direct evidence
that mercurials increase the permeability of the proximal tubule cells to
Na+. They infused Na^' into the renal artery during mannitol diuresis and
found the Na+ flux to be increased by meralluride at diuretic doses. It was
thus concluded that the net resorption of Na+ is decreased because of the
augmented backward leak.
EFFECTS ON TISSUE FUNCTIONS
Surprisingly few thorough or quantitative investigations of the effects of
mercurials on tissue function have been made, especially considering the
long-known toxic and therapeutic actions of these substances, and most of
them are on the heart. Since in essentially no case have functional and
metabolic disturbances been correlated, most of the results will be treated
cursorily and presented principally to point out some fields in which inter-
esting work may be done. There is great need for the study of the metabolic
changes produced in tissues isolated from animals treated with mercurials,
since only in this way can one be certain that the effects observed are re-
lated to the in vivo interference with function. This has been done with
the kidney but is notably absent with other tissues. Much of the in vitro
work with mercurials has been done with relatively high concentrations
(around 1 mM or higher) so that it is impossible to determine if the
results are applicable to the effects seen in the whole animal. Indeed,
studies of the simultaneous changes in fuction and metabolism under
any conditions are very rare.
Skeletal Muscle
Resting rat diaphragm treated with 1-2 niM mersalyl soon exhibits fibril-
latory twitches accompanied by rapid small (1 mv) variations of the mem-
brane potential, which persist for several minutes, followed by a rise in
the resting tension during the next 10 min, and finally by a further slowly
developing irreversible contracture and loss of excitability (Kuschinsky et
al., 1953). Stimulation during the early action of mersalyl produces a nor-
mal contractile tension but there is a marked retardation of relaxation,
the duration of contraction increasing 5- to 10-fold. Hg++, on the other
hand, causes only the slow contracture and loss of excitability. Decame-
thonium and curare abolish the fibrillation due to mersalyl, indicating that
the action of the mercurial is on the end-plate. Furthermore, chronically
denervated muscle does not show mersalyl fibrillation. Inasmuch as physo-
stigmine and neostigmine cause a similar fibrillation, and since mersalyl
938 7. MERCURIALS
inhibits muscle cholinesterase, it was concluded that the fibrillation results
from inhibition of cholinesterase, allowing acetylcholine to accumulate
(Kuschinsky and LiiUmann, 1954). None of the other actions of mersalyl
appears to be related to this inhibition. The delayed relaxation was claimed
to be similar to that produced by veratrine but, if so, it does not provide
much information on the mechanism, since one is ignorant of how vera-
trine acts.
Contracture of frog muscle by Hg++ had been noted by Bacq (1942),
Beck and Bein (1948), and Krueger (1950). Bacq assumed this to be an
effect such as that given by iodoacetate, i.e., a Lundsgaard effect due to
glycolytic inhibition, but Krueger showed that Hg++, in contrast to iodo-
acetate, does not bring about a reduction in lactate concentration during
rigor. Kuschinsky and Liillmann (1954) found that mersalyl causes a rapid
loss of muscle K+ and attributed the initial rapid contracture to a depola-
rization of the fibers, a conclusion shared by Muscholl (1958), who demon-
strated a fall in the membrane potential coincident with contracture. How-
ever, the delayed contracture must have another origin and, although Kus-
chinsky and Liillmann postulated a Lundsgaard mechanism, there is no
evidence one way or the other. The action potential traces presented by
Muscholl (1958) show that mersalyl reduces the magnitude so that the over-
shoot is lost, slows both depolarization and repolarization, and hence pro-
longs the duration of the action potential, effects quite different than those
seen in heart (page 945). Frog muscle after-potentials seem to be unaffect-
ed by mercurials between 0.1 and 2 mM (Macfarlane and Meares, 1958).
Contracture by mersalyl is dependent on Ca++ in the medium, but this
may be true for most types of contracture (Kutscha, 1961).
The contraction of glycerinated muscle fibers by ATP is inhibited by
Hg++ at concentrations around 0.01-0.1 mM (Godeaux, 1944, Korey,
1950; Hasselbach, 1953; Edman, 1958) and by mersalyl at similar con-
centrations (Portzehl, 1952; Edman, 1959). A preparation contracted by
ATP is relaxed upon addition of the mercurial. These effects are irrever-
sible by washing or treatment with cysteine. Weber and Portzehl (1954)
suggested that the inhibition of the ATP effect is due to a block of ATPase
so that ATP can act only as a plasticizer, but there is also, according to
Edman (1958), a direct effect since there is some relaxation in the absence
of ATP.
A great deal of work has been done on the behavior of muscle contractile
proteins exposed to mercurials, and the importance of SH groups has been
conclusively demonstrated. The effects of the mercurials are summarized in
Table 7-20. Both actin and myosin possess SH groups of differing degrees of
reactivity and function. In the complexing of actin and myosin to form
actomyosin, it is the SH groups of myosin which are important (Bailey
and Perry, 1947; Kuschinsky and Turba, 1951). Bailey and Perry felt that
EFFECTS ON TISSUE FUNCTIONS
939
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940 7. MERCURIALS
the binding might be through SH groups but, as Gergely et al. (1959) point-
ed out, it only indicates that SH groups are in the vicinity of the binding
groups. The myosin SH groups concerned with the binding of actin react
with mercurials more readily than the SH groups upon which ATPase ac-
tivity depends (Fig. 7-22) (Barany, 1959). However, MM presents an ex-
ception, in that it inhibits ATPase and actomyosin formation in a parallel
fashion (Barany and Barany, 1959 a), possibly indicating that the size
of the group on the mercurial is important. G-actin is more reactive than
F-actin, due perhaps to shielding of the SH groups in the polymerized form.
G-actin cannot polymerize unless ATP is present and, since mercurials re-
lease ATP from actin, the possibility of the effect on the G-actin ±^ F-actin
transformation being due to an interference with ATP binding was examin-
ed, but most mercurials were found to cause a rapid loss of polymerizability
without appreciable loss of ATP (Drabikowski and Gergely, 1963). When
the ATP is finally lost, the actin has been changed irreversibly, and there
is further evidence from optical rotation that structural changes are pro-
duced (Tonomura and Yoshimura, 1962). Katz and Mommaerts (1962) con-
sider the six SH groups of G-actin to fall into three categories: two rapidly
reacting, two intermediately reacting, and two slowly reacting, only the
last two being necessary for polymerization. It is interesting that the SH
groups of G-actin are made more reactive to p-MB by Mg++ and less reac-
tive by Ca++ (Katz, 1963). It was postulated that Mg++ brings about an
open configuration whereas Ca++ tends to produce a closed configuration,
the SH group being in a crevice.
The effects of the mercurials on extracted muscle proteins are certainly
interesting and often obtained at low concentrations, but there is at pres-
ent essentially no way of determining if they are at all responsible for any
of the changes observed in intact muscle. It would be particularly important
to know if rigor is related to any of the actions on actomyosin, but actually
most of the actions described above could not very well explain why a
muscle goes into contracture. The mechanisms by which mercurials alter
muscle function are thus obscure, but it is not unlikely that the earliest
effects are on the permeability and transport systems in the membrane.
More information will be provided in the following section in which the
effects of the mercurials on cardiac muscle will be discussed.
Heart
The detrimental effects of the mercurials on the heart have long been
recognized and many cases of clinical deaths from intravenous injections
of mercurial diuretics have been reported. It is generally agreed that death
is attributable to the direct action on the heart during temporary high
plasma concentrations of the mercurials, whereas at the usual low concen-
trations required for diuresis there are no detectable cardiac effects. The
EFFECTS ON TISSUE FUNCTIONS 941
actions of the mercurials on the heart have been well studied but the
mechanisms involved, and whether these actions involve metabolic distur-
bances, are not known. Hepp (1887) observed that the toxic effects of the
organic mercurials differ from those produced by Hg++, and that ethyl-
Hg++ stops the frog heart in diastole. Dreser (1893) studied the cardiac
effects of several complexes of Hg++, both in vivo and on perfused isolated
frog hearts. Cardiac depression was noted after injection of around 2.5 mg
of the rhodanate, succinimide, and cyanide complexes of Hg++, and stand-
still of the isolated heart was brought about by 0.45 mM of the thiocyanate
complex. These and other early investigations showed only that the heart
can be depressed by mercurials and that the relative potencies depend on the
substances with which the Hg++ is complexed. For example, Miiller et al.
(1911) showed that compounds of the type R — Hg — OH or R — Hg — CN are
around 10 times as cardiotoxic in cats as compounds of the type R — C —
Hg — C — R in which the Hg is bonded to two C atoms, and that the toxicity
is related to the rate at which these compounds react with sulfide. The first
serious study of the effects of the mercurials on the heart was undertaken
by Salant and his co-workers in Georgia from 1921 to 1931, the results of
which will be discussed throughout the following sections.
(A) Isolated heart preparations. The effects of the mercurials depend on
the species, the type of preparation, the mercurial used and its concentra-
tion (Table 7-21). As would be expected, there is generally depression of
contractile amplitude, rate of beating, and rate of conduction, the last lead-
ing to varying degrees of a-v block and dissociation of the atria and ven-
tricles. Ventricular standstill often occurs before the atria cease to beat.
Tachycardia and fibrillation are frequently seen in animals poisoned acutely
with the mercurials, but are not noted in isolated preparations, although
occasionally with low concentrations, or initially, some increase in rate and
contractile amplitude may be observed. The ventricular dysrhythmias in
vivo may be in part the result of altered a-v and ventricular conduction,
but in isolated preparations there is little evidence for the appearance of
rapidly discharging ectopic foci. Most of these effects are irreversible, or
only partially reversible at the lowest concentrations and with short expo-
sures, but dimercaprol or cysteine is occasionally able to reverse rather ad-
vanced degrees of depression (Ruskin and Johnson, 1949). The selective de-
pression of the rat atrial rate is marked; at 0.0013 vaM Hg++ the rate is
reduced 35% in 15-20 min while the amplitude is unaffected, and at 0.0025
raM the rate may be inhibited 90% and the amplitude some 15% greater
than normal (Berman, 1951). Glutathione and dimercaprol effectively pro-
tect both the atria and ventricles.
The development of contracture is not nearly as common with the mer-
curials as with iodoacetate, and in fact has been noted only in frog and turtle
hearts. Rat atria treated with p-MB are slowly and markedly depressed
942
7. MERCURIALS
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but no elevation of resting tension is seen even after 2 hr (Webb and Hollan-
der, 1959). It may well be that the mercurials so readily depress membrane
functions — pacemaker discharge, conduction, etc. — that the hearts cease
beating before they are depleted of ATP. Mendez (1946) noted that frog
heart treated with 0.56 mM p-MB stops beating quite soon and before full
contracture has developed; however, if the heart is electrically stimulated
it can be put into complete rigor. It has been observed many times that the
heart stops in diastole in vivo following intravenous injections of the mer-
curials.
(B) Cardiac effects in whole animals. Sudden death during or following
the intravenous injection of diuretic mercurials clinically has usually been
attributed to ventricular fibrillation. In animals (usually cats and dogs)
the mercurials produce the following cardiac effects: initial cardiac depres-
sion, disturbances in a-v conduction leading occasionally to a temporary
ventricular bradycardia, atrial flutter or fibrillation (rarely), slowing of
ventricular conduction, and often ventricular tachycardia before the ter-
minal fibrillation (Jackson, 1926 a, b; Salant and Nadler, 1927; Macht,
1931 b; DeGraff and Lehman, 1942). The effects may be quite complex and
are undoubtedly due to a variety of actions. McCrea and Meek (1929) felt
that one of the major actions is a descending stimulation followed by a de-
pression of the cardiac conducting tissue. The innervation of the heart
probably plays a role in the initiation of the dysrhythmias, since atropini-
zation or cutting the vagi in dogs prevents ventricular fibrillation due to
mersalyl (Jackson, 1926 a). It is also known that epinephrine potentiates
the fibrillatory action of the mercurials. The fall in blood pressure invariably
observed during intravenous infusion of the mercurials must induce sym-
pathetic activity and a rise in plasma catecholamines. Various salts of Hg++
apparently are not so likely to induce dysrhythmias as the organic mer-
curials and are more directly depressant (Salant and Kleitman, 1922).
These effects are not dependent on the vagi since they occur after atropini-
zation. However, Hg++ can produce conduction disturbances and dysrhyth-
mias in dogs, and occasionally ventricular fibrillation (McCrea and Meek,
1929). The intravenous lethal dose depends on the mercurial, the species
used, and the rate of injection; in most cases it falls between 10 and 50 mg
Hg/kg for the common diuretic mercurials (DeGraff and Lehman, 1942;
Chapman and Shaffer, 1947; Lehman et al., 1950; Farah et al., 1951). Inor-
ganic Hg++ is somewhat more toxic, the lethal dose usually being around
one third to one half that for the organic mercurials. The toxicity of Hg++
is dependent on the blood pH, being least between 7.4 and 7.6, and increas-
ing on either side, especially between 7.14 and 7.35 (Salant and Nadler,
1927). Since comparable experiments have not been run on isolated
hearts, it is impossible to understand the mechanism for this sensitive pH
dependence; it is difficult to accept that a direct effect of pH could
EFFECTS ON TISSUE FUNCTIONS 945
alter 2- or 3-fold the sensitivity of the heart to the Hg++ ion, and it is
more likely that secondary changes due to the alteration of the pH are
responsible.
The electrocardiographic changes in dogs are similar for all the diuretic
mercurials tested and for HgCl,, and are primarily the result of conduction
disturbances. They may be summarized briefly as follows: depression and
change of configuration of the st segment, increase in height of the t wave,
widening and notching of the qrs complex, widening of the p wave, and
increase of the p-r interval (McCrea and Meek, 1929; Farah et al., 1951).
In the rat there is an initial flattening of the t wave, and eventually the
p wave may disappear (Gessler and Kuner. 1960). Most of these changes
are, of course, simply due to the slowing of conduction throughout the
myocardium. The t wave changes are different from those seen with most
metabolic inhibitors and are perhaps related more to a membrane effect
than a metabolic disturbance. It is interesting that p-MB acts differently
than the mercurial diuretics in that no qrs changes are seen, even at doses
2-3 times the lethal doses of the other mercurials, and death is not due to
fibrillation but to ventricular asystole (Farah et al., 1951). The lethal dose
of p-MB is also about 4 times as great as for the other mercurials.
(C) Transmembrane potentials and ionic shifts. The membrane charac-
teristics of rat atria are changed markedly by 0.05 mM p-MB, although
the rate of action is rather slow (this is probably not due to slow penetra-
tion into the atria since the potentials are recorded from cells at the sur-
face) (Webb and Hollander, 1959). During the first 20-30 minutes there is
no significant alteration of the contractile behavior, but there is a progres-
sive reduction in the magnitude of the action potential, an acceleration of
the repolarization rate, and a slowing of conduction. It is possible that these
early effects arise from selective action on the cell membranes. During the
next hour these changes continue but, in addition, contraction becomes im-
paired. At 1 hr the changes may be summarized as follows: no significant
change in resting potential ( + 2.1%), a severe depression of the action
potential magnitude ( — 29%), a faster repolarization ( + 51%) leading to
a shorter action potential (—60%), a decrease of the developed tension
( — 48%), a slowing of conduction (—38%), and a prolongation of the
latent period (+61%). Even during this later period it appears that the
contractile depression is due mainly to the shortening of the action potential,
and to some extent to its reduced magnitude, and there is little evidence
for direct effects on the contractile systems. It may well be that p-MB
penetrates into the cells rather poorly and that some of the other mercurials
would not have so selective an action on the membrane. Stein et al. (1960)
reported that mersalyl (0.2 mM) causes a faster repolarization and contrac-
tile depression in guinea pig atria, but no changes in either the resting or
action potential magnitudes were observed. The failure of the mercurials
946 7. MERCURIALS
to affect the resting potential in either rat or guinea pig atria indicates
that no marked changes in intracellular K+ occur during the duration of the
experiments, so that appreciable depression of ion pumps or increase in ion
permeabilities seem not to be a characteristic of the action on the heart.
It is difficult to interpret the cardiac ionic changes noted by Gessler and
Bass (1960) in rats poisoned with HgClg, because the electrolyte changes
resulting from the renal effects (either polyuria or anuria) probably com-
plicate the picture. However, with a dose of HgClg sufficient to produce a
long-lasting polyuria there is only a minor fall in the myocardial K+/Na+
ratio (1.78 to 1.60), and, although plasma K+ rises, the tissue/plasma ratio
for K+ certainly does not drop very much, although Gessler and Kuner
(1960) felt that the qt changes are perhaps correlated with alterations of
this ratio. The results on isolated atria support the concept that the major
effect is on the ionic flux rates during membrane activation.
The various regions of the heart respond differently to the mercurials
as they do to other inhibitors and drugs. Isolated pig ventricle fibers are
not appreciably affected by 0.26 milf p-MB but the Purkinje fibers are
more sensitive (Kleinfeld et al, 1964). The action potential magnitude in
the ventricle may fall around 10% within 15 min but there is little further
change, while the resting potential and action potential duration are not
significantly modified. In the Purkinje fibers, on the other hand, the magni-
tude of the action potential is rapidly depressed, falling approximately 25%
within 10 min, after which another rapid fall occurs between 20 and 30 min.
Since the resting potential is unchanged for 20 min, there is initially a
marked decrease of the overshoot; the resting potential later falls gradually.
The duration of the action potential is surprisingly not altered in contrast
to the results in atria. Although no evident explanation for these differences
is at hand, it was considered that the greater glycolytic activity of the
Purkinje fibers might predispose them to inhibition. We have seen, however,
that the glycolytic pathway is probably less sensitive than the cycle to the
mercurials.
(D) Cardiac innervation and responses to acetylcJwline and epinephrine.
Salant and Kleitman (1922) noted that Hg++ exerts some vagal blocking
action in the cat heart, but Jackson (1926 b) could find no acceleration of
the heart and no evidence of vagal block by mersalyl in dogs. Salant and
Brodman (1929 a) reinvestigated this question and established that Hg++
first sensitizes the heart to the vagus and later blocks the vagal endings.
It is during the first sensitization phase that dysrhythmias are apt to occur,
which is reasonable since acetylcholine is profibrillatory as a result of its
marked shortening of the action potential duration. Hg++ and p-MB at
0.01-0.1 niM antagonize the effects of acetylcholine on the frog heart, the
mercurials being allowed to act for 3-40 sec and then washed out (Pohle
and Matthias, 1959). It was concluded that the acetylcholine receptors may
EFFECTS ON TISSUE FUNCTIONS 947
contain SH groups, a conclusion previously reached by Turpaev (1955).
Nistratova and Turpaev (1959) titrated the SH groups in a frog ventricle
homogenate and found that the presence of acetylcholine alters the shape
of the titration curve, but not the total number of SH groups titrated —
part of the SH groups becomes less reactive in the presence of acetylcholine.
Inasmuch as cholinesterase is inhibited to some extent by mercurials (Ta-
ble 7-13). it is possible that this can account for the vagal sensitization, a
secondary blocking of the receptors for acetylcholine reversing this effect.
There is no evidence for specific interference with the action of .the cate-
cholamines on the heart, but epinephrine potentiates the profibrillatory
action of the mercurials (Jackson, 1026 a). Yet Salant and Brodman (1929
c) claimed that the cat heart is most sensitive to Hg++ when the sympa-
thetics are blocked by ergotamine, and that high concentrations of epineph-
rine actually protect the heart against the mercurials. In any event, it is
likely that the over-all effects of the mercurials on the heart, especially in
the whole animal, must to some extent involve the sympathetic and para-
sympathetic innervation. Mercurials can release catecholamines from ad-
renal medulla granules (D'lorio, 1957) but it is not known if such a release
can occur in the heart or other tissue.
(E) Consideration of some mechanisms of cardiac action. There is little
justification for discussing mechanisms by which the mercurials affect the
heart because essentially no basic work to elucidate the cellular actions has
been done. The interesting observations of Salant and Nagler (1930, 1931)
on the relation of the response to Hg++ of the frog heart and the level of
Ca++ in the medium may provide some clue. If the Ca++ is reduced to around
one half normal, the heart is depressed much more readily by Hg++, but if
the Ca++ is reduced further (this in itself suppressing contractions), Hg++
may then actually stimulate the amplitude. High Ca++ somewhat antago-
nizes the action of Hg++. Increasing the K+ slows the rate and then Hg++
accelerates the heart and seems to have less effect on the contraction. The
authors suggested that the alterations in response to Hg++ might be due
to permeability changes brought about by Ca"^+, but we now know that
Ca++ has other, perhaps more important, effects on the heart. It would
be interesting to know how mercurials affect the positive inotropic action
of Ca++. It would also be worthwhile to determine if mercurials inhibit the
various ATPases of the heart. Padykula and Herman (1955) showed histo-
chemically that p-MB strongly inhibits cardiac ATPase, but it is not known
if this occurs in vivo.
For the purpose of this volume it would be of some importance if the
cardiac effects could be correlated with any of the well-known enzymic or
metabolic inhibitions exerted by the mercurials, but this cannot be done
because there are no investigations of metabolic changes during mercurial
action. Even the results reported on respiratory inhibition, for example by
948 7. MERCURIALS
Ruskin and Ruskin (1953), where 5.8 mM meralluride depresses rat heart
slices 57%, are scarcely pertinent to understanding how the mercurials act.
We have postulated previously that the site of mercurial action on the heart
is mainly at the membrane to alter ionic fluxes. This does not imply that
the action is nonmetabolic, since enzymes in the membrane may be the
ultimate vulnerable points of attack.
Smooth Muscle
It is necessary to consider briefly the effects of the mercurials on smooth
muscle, if only because calomel was used for centuries as a purgative. We
have not discussed Hg+ because little is known about its actions on meta-
bolic systems. Hg++ salts are also capable of causing diarrhea, and it is
likely that Hg+ is active after being oxidized to Hg++. Hand et at. (1943)
developed histochemical tests for Hg, Hg+, and Hg++, and found in various
animals that within a few minutes of the intravenous injection of mercurous
acetate they could detect both Hg+ and Hg++ in the parenchymal and en-
dothelial cells of the kidney, the latter predominating. Whole blood oxi-
dizes Hg+ to Hg^^ quite rapidly. Many theories of the mechanism of the
purgative action, several rather fanciful, have been advanced, but very
few justify even serious criticism. HgClg did not achieve its name of "cor-
rosive sublimate" in vain; it is a direct irritant of tissues, by which is meant
that it induces cellular damage of both metabolic and nonmetabolic origin,
this initiating an inflammatory sequence, which in the intestine causes in-
creased activity, depression of the ability to absorb water and various sub-
stances, and consequently colitis and diarrhea. In severe mercury poison-
ing there is a hyperemic and hemorrhagic appearance of the intestine, with
erosion and necrosis. One is thus tempted to attribute the diarrhea to such
a nonspecific action, but there is some evidence against this. Isolated in-
testine is stimulated in a characteristic way by Hg++ in low concentrations
(0.004-0.02 mM), tonic contractions being markedly increased with sup-
pression of rhythmic activity (Salant and Brodman, 1929 b). Organic mer-
curials apparently can stimulate similarly; e.g., merbaphen augments peri-
staltic activity of isolated cat intestine (Govorov, 1936), and p-MB at
0.0057 mM stimulates the rat intestine around 10% (Goodman and Hiatt,
1964), although p-MB was reported to inhibit rabbit intestine (Haley, 1945),
perhaps because of too high a concentration (not given, but around 0.02
mM). The stimulations produced by both Hg++ and merbaphen are readily
blocked by atropine, indicating that the action is to some extent mediated
through the vagal nerves in the intestine. Govorov believed merbaphen to
be a parasympathomimetic substance. There is also some increase in the
sensitivity of the intestine to vagal stimulation when the tissue is treated
with Hg++, recalling similar actions on the heart, and it is possible that
inhibition of cholinesterase is involved. However, it seems unlikely that
EFFECTS ON TISSUE FUNCTIONS 949
mercurials in vivo can produce intestinal stimulation by such a selective
inhibition, and the subject needs further investigation.
Another mechanism which must be given serious consideration is hista-
mine release. Bachmann (1938) showed that the isolated cat intestine ex-
posed to Hg++ releases a substance which behaves like histamine pharma-
cologically, and felt that at least some of the action on the intestine can be
explained by this release. It may be mentioned that Hg++ has been report-
ed to release histamine from perfused dog Kver (Feldberg and Kellaway,
1938) and p-MB to release histamine from rat mast cells (Bray and Van-
Arsdel, 1961), but in both cases the concentrations used were too high to
enable correlation with in vivo effects; it is quite likely that any substance
at high enough concentration or any irritant histotoxic agent will release
histamine.
Nervous System
Neurological dysfunction is common in mercury poisoning (page 951 ) but
it is not known if the action is axonal or synaptic. One usually assumes that
metabolic disturbances affect primarily junctional transmission. Halasz et
al. (1960) have shown that transmission in the cat superior cervical gan-
glion is rapidly and reversibly depressed by p-MB at 0.0056-0.02 mM, while
simultaneously the effects of injected acetylcholine are potentiated. If the
concentration is increased toward 0.028 mM, this potentiation of acetyl-
choline is lost. The stimulatory action of K+ is unaffected by lower and
depressed by higher concentrations. Inhibition of acetylcholine synthesis is
apparently not involved since there is a store of acetylcholine and the de-
pression of transmission is immediate, so the authors postulate a reduction
of the response of the postganglionic cells to acetylcholine. However, at the
time of the initial suppression of transmission there is actually a potentia-
tion of the acetylcholine response, which is difficult to explain, particularly
since cholinesterase inhibition is not a likely hypothesis for ganglia. It is
possible that SH groups of the acetylcholine receptors are reacted at higher
concentrations of the mercurial, as has been suggested for cardiac receptors.
Recordings of the postganglionic membrane potential changes are needed
to interpret these results.
Axonal conduction is also depressed by p-MB at low concentrations (H.
M. Smith, 1958). Conduction in the frog sciatic nerve is blocked in 4 min
by 0.002-0.02 mM p-MB and in lobster giant axon in 3 min by 0.045-0.07
mM p-MB. There is a gradual depolarization of the axon but block occurs
long before the potential is lost. The post-tetanic hyperpolarization of sym-
pathetic C fibers is more sensitive to metabolic inhibitors than the magnitude
of the action potential, and is decreased by mersalyl at 0.34 mM (Greengard
and Straub, 1962). However, the nature of such a hyperpolarization and its
significance for conduction are not understood. The injection into the squid
axon of 6 X 10~^ ml/mm of 7.5 mM p-MB is without effect on the action
950 7. MERCURIALS
potential (Brady et al., 1958), which may indicate that the mercurial must
act on the external surface of the membrane to block conduction. The
changes in the structure of the myelin sheath of nerves brought about by
Hg++ at high concentrations (around 10 mM) are certainly not relevant to
acute experiments with low concentrations, but in chronic mercury poison-
ing it is possible that sufficient Hg++ is incorporated in the myelin to disturb
nerve function (Millington and Finean, 1958, 1961). Thus at the present
time we cannot decide whether the primary action of the mercurials is on the
axon or on the synaptic regions, or on both, especially in chronic poisoning.
Skin
Various types of skin reaction to the mercurials administered both sys-
temically and topically have been recognized for years. Some of these are
undoubtedly of the allergic or sensitivity category and need not concern us.
Mercurial diuretics, like mersalyl, when injected in small amounts into the
skin cause blisters, and Hahn and Taeger (1931) concluded that there is a
relationship between diuretic activity and vesication. Almkvist (1922) had
claimed that mercurials cause vascular dilatation in the skin, with result-
ing edema, by a paralysis of the sympathetic nerves, but there is little evi-
dence that this is a significant factor. Hellerman and Newman (1932) noted
that alkyl mercurials are powerful vesicants and can cause a severe der-
matitis. These early observations are of interest in the light of the relation-
ship between SH group reaction in the skin and vesication established by
work on the arsenical war gases, and one might postulate that the mercur-
ials have a metabolic basis for their effects on skin, perhaps an inhibition
of the cycle. Hg++ reduces frog skin potentials across both borders and in-
creases the outer membrane resistance (Lodin et al., 1963).
EFFECTS OBSERVED IN THE WHOLE ANIMAL
It is difficult to present the toxicology of the mercurials concisely be-
cause the effects depend on the type of mercurial, the species considered,
whether the poisoning is acute or chromic, the route by which the mercurial
is taken into the body, and many other factors. The symptoms of chronic
mercury poisoning (mercurialism) in man are quite variable and usually
not correlated with the blood or urinary levels of mercury. Frequently
urinary mercury may be considerably higher than the normal range and
yet no symptoms occur; however, definite symptoms may sometimes be ob-
served in those whose level is in the normal range. This lack of correlation
with urinary levels and the protean nature of the poisoning not only make
diagnosis frequently difficult but indicate that the individual pattern of
response must relate to a number of obscure factors, such as hereditary
EFFECTS OBSERVED IN THE WHOLE ANIMAL 951
constitution, vitamin intake, electrolyte balance, protein nutrition, and
other imponderables. The concentrations of mercury in the blood or urine
are, of course, not the critical determinants in poisoning when the mercury
has been slowly taken into the body over a period of months or years.
Mercury is picked up by the various tissues at different rates and to dif-
ferent degrees, and it is the eventual levels of mercury in these tissues which
determine the toxic response. Such accumulation may occur over a long
time and several weeks be required before a balance between intake and
excretion is achieved. One factor which must be of importance, but about
which little is known, is the concentrations of the various thiols in the blood,
since this will not only alter the over-all tissue uptake but will modify the
pattern of distribution in the body. Most mercurialism in adults is industrial
in origin and due to the inhalation or ingestion of small amounts of metallic
mercury or mercury compounds daily over a prolonged period.
General Symptoms of Mercury Poisoning
We have discussed the most im])ortant aspects of acute poisoning by the
inorganic and organic mercurials, namely, the effects on the cardiovascular
and renal systems, and little more need be added. Slow inhalation of mer-
cury vapor produces typical poisoning of the kind commonly seen with the
inorganic mercury salts, because the metallic mercury is oxidized during
and after absorption. However, when the concentration of mercury vapor
is high, absorption is faster than oxidation and unique symptoms are ex-
hibited, e.g., hyperthermia, tachypnea, cough, nausea, dizziness, and weak-
ness (Carpenter and Benedict, 1909). These may be due primarily to the
greater uptake of mercury by the central nervous system under these con-
ditions. It is important to emphasize that the character of mercury poison-
ing depends greatly on the tissue distribution, and hence on the physico-
chemical properties of the mercurial. Thus the more or less volatile, lipid-
soluble, alkyl mercurials produce quite a different picture from the inor-
ganic or diuretic mercurials (Miiller et al., 1911; Hunter et al., 1940). Alkyl
mercurials act rather selectively on the central nervous system to produce
ataxia, paralysis, and depression — in higher concentrations they act much
like certain anesthetics — and acutely these effects are possibly unrelated
to mercury or reactions with SH groups.
The acute effects of mercurials on the central nervous system are often
marked but have not been analyzed in detail. When HgCL, is injected sub-
cutaneously into rats at the high dose of 17 mg/kg, there is progressive loss
of the reflexes and all have disappeared after 54 hr, this being reversible
upon administration of a Hg++-binding thiol (Galoyan and Turpaev, 1958).
Conditioned reflexes are suppressed partially at the much lower dose of
3.7 mg/kg (Galoyan, 1957). The respiration is usually affected and may be
taken as an index of certain central actions. Respiratory stimulation by
952 7. MERCURIALS
Hg++ (Hanzlik, 1923 c) and mersalyl (Jackson, 1926 b) has been noted,
this being attributed to a direct medullary effect at low mercurial concen-
tration, but lethal doses of both HgClg and PM cause dyspnea and depres-
sion of the respiration (Wien, 1939). The injection of certain mercurials can
produce a very rapid fulminant type of reaction characterized by dyspnea,
convulsions, and death, and this was also attributed to a central effect on
the respiratory centers (Fourneau and Melville, 1931). Direct effects on the
central nervous system were observed by Pentschew and Kassowitz (1932)
following suboccipital injections of HgClj at the minimal lethal dose (around
0.2 mg); tremors, convulsions, and other motor disturbances occur after
16 hr and last for several days. It would be interesting to know the form,
or forms, in which Hg++ penetrates into the nervous system, whether mainly
as the uncharged HgClg or in combination with thiols and other substances
in the blood.
The most characteristic symptoms of mercurialism, regardless of the type
of mercurial responsible, may be summarized as follows (Hunter et al., 1940;
Cumings, 1959, p. 78; Noe, 1960; Kantarjian, 1961). (1) A fine intention
tremor, starting in the fingers and hands, and progressing to the feet, eye-
lids, cheeks, tongue, and neck. The motor activity is primarily affected and
usually there is little if any disturbance in sensation. (2) Insomnia, anorexia,
and various emotional alterations, such as mood depression and timidity.
There is generally little effect on intelligence or memory. (3) Erethism, or
blushing, is often common, but whether it is due to emotional disturbances
or alteration of the autonomic vascular control is unknown. Sometimes,
especially in infants, the skin may become red; such erythema is most likely
vascular in origin. (4) Stomatitis, salivation, and gingival swelling are fre-
quent and possibly due to the secretion of mercurials in the saliva. Of these
symptoms, and others less common, only the nervous system changes lend
themselves to an analysis of the mechanisms which may be involved, but we
shall see that regrettably little can be concluded.
Urinary mercury excretion in normal individuals is usually between 1
and 15 //g/day, but may be so low as to be imdetectable or considerably
higher without obvious symptoms. In patients with evident mercurialism,
the urinary mercury may vary widely — excretions between 3 and 8000
//g/day have been reported — but it is generally above 250 //g/day. The
level will depend on the daily uptake and whether the individual is in com-
plete balance or not. The fact that the same degree of severity of symp-
toms may be observed in patients with very different urinary levels sug-
gests that the susceptibility to mercury varies widely, but possibly the
tissue concentrations of mercury are much more uniform than the urinary
concentrations.
In view of the selective effect of iodoacetate on the retina and visual
function, it is interesting to inquire as to whether other SH reagents, such
EFFECTS OBSERVED IN THE WHOLE ANIMAL 953
as the mercurials, possess this action. Since the mercurials seem to lack
marked effects on glycolysis in intact tissues, one would not expect visual
disturbances if these are indeed due to glycolytic inhibition. Sorsby et al.
(1957) could detect no retinal degeneration in rats or rabbits given HgClg
or p-MB under conditions where lesions are produced by iodoacetate. How-
ever, in poisoning with certain organic mercurials there may be a marked
constriction of the visual field (Hunter et al., 1940). It is not known
whether this is a retinal effect or due to nerve degeneration. The sjaithesis
of rhodopsin from opsin and retinenci is blocked by 0.1 mM p-MB though
an action on opsin (Wald and Brown, 1951, 1952), and the synthesis of
iodopsin is likewise suppressed (Wald et al., 1955). The mercurial does not
alter rhodopsin directly but readily bleaches iodopsin. If this effect on the
regeneration of visual pigments occurs in vivo it has not been reported.
For many years a rather uncommon disease of infants has been recog-
nized and called pink disease, infantile acrodynia, or erythredema, and is
characterized by a redness and swelling of the extremities and certain other
skin areas, along with photophobia, irritability, loss of reflexes, and muscular
hypotonia. Dr. Warkany of the Children's Hospital in Cincinnati in 1945
examined an infant suffering from this disease and found a urinary mercury
concentration of 360 ^g/liter. A summary of 20 cases showed that most
infants with pink disease had definitely elevated mercury levels — 75%
had more than 50 //g/liter and 10% more than 400 //g/liter ■ — whereas
most control infants showed undetectable levels (Warkany and Hubbard,
1948). It is now generally agreed that the majority of cases of pink disease
are due to mercury poisoning, which manifests itself somewhat differently
in infants than in adults although some of the neurological changes are
similar. The source of the mercury is usually calomel or mercury ointments.
Of the group of 54 studied by Zellweger and Wehrli (1951), around 80% had
been given such drugs, but in the remainder there was no obvious source
of mercury. The situation is probably more complex than believed originally.
Some of the symptoms may not be directly due to the mercury but are
predisposing conditions to mercury poisoning (Barrett, 1957). Thus a high
intestinal alkalinity, due in infants to faulty acid secretion in the stomach,
may accelerate the oxidation of calomel and increase its toxicity, and simul-
taneously reduce absorption of certain fatty acids, this latter possibly be-
ing responsible in part for the acrodynia. Since acrodynia in animals may
be induced by pyridoxine deficiency, there is also some possibility that the
mercury either inhibits some phase of pyridoxine metabolism or blocks a
pyridoxal-P enzyme to produce symptoms similar to deficiency. Since typi-
cal pink disease can occur in the absence of excess mercury intake or signifi-
cant urinary levels, one must assume that some basic metabolic disturbance
is the basis of this malady and that it can be brought about in various ways.
That aU the symptoms are not immediately due to mercury seems to be
954 7. MERCURIALS
indicated by the fact that administration of dimercaprol is not remarkably
successful, although certain clinical improvement has been noted (Bivings
and Lewis, 1948).
Histological Changes
The neurological picture of fasciculations, hyperreflexia, tremor, and mo-
tor weakness, followed by muscular atrophy, often seen in chronic poison-
ing with the organic mercurials — as reported, for example, by Kantarjian
(1961) in individuals eating bread treated with the fungicide Granosan M
(ethylmercuri-p-toluene sulfonanilide) — accompanied by occasional numb-
ness or paresthesias, may clinically resemble amyotrophic lateral sclerosis
(Brown, 1954). Rats and monkeys chronically poisoned with MM show
Wallerian degeneration of the peripheral nerves (Hunter et al., 1940). The
peripheral nerves and the posterior spinal roots are affected first, and later
the posterior columns and granular layer of the middle lobe of the cerebel-
lum. The ataxia and tremor could result from the cerebellar lesions. Bila-
teral cortical atrophy in the area striata was associated with the reduction
of the visual field. A good review of the neurological changes has been pro-
vided by Noe (1960). The renal and intestinal changes have been described,
and we shall only note that degeneration of the liver has also been observed
(MacNider, 1918 b). Dogs given HgClg orally (15 mg/kg) exhibit a deposi-
tion of fat in the cells surrounding the central vein, followed by cloudy
swelling and necrosis, with eventual extension to the periphery of the lobule.
Such hepatic changes could well be responsible for some of the over-all
metabolic disturbances observed in animals.
Foulerton (1921) believed that Hg++ has an affinity for lipids, not only
because of the solubility of HgClg in fat but also due to the formation of
oleates, and is transported to the liver in the circulating blood fat. The liver
damage then results in a defective lipid metabolism. There are certainly
definite disturbances in lipid metabolism — for example, Ogilvie (1932)
found an immediate and considerable rise in blood lipid following adminis-
tration of HgCla; subsequently there is a fall and a second rise — but no
evidence to indicate the mechanism involved. Toxic doses of mersalyl in
rats produce hypoglycemia and reduce the liver glycogen to essentially zero
(Dzurik et al., 1963). It was stated that these effects are secondary to renal
dysfunction and not a manifestation of a direct action of the mercurial on
the tissues, but this seems unlikely and there are possibly several factors
of importance, including epinephrine release as a result of a nonspecific
stress reaction. Free amino acids in the livers of rats given HgClg for 10
days were determined by Thoelen and Pletscher (1953), and it was shown
that although serine, leucine, and phenylalanine do not change significantly,
cystine rises to 3 times the control value. This was interpreted as a detoxi-
fication response. One must question the ability of animals to increase the
synthesis of specific amino acids for the purpose of complexing with a non-
EFFECTS OBSERVED IN THE WHOLE ANIMAL 955
physiological metal ion. Might it not be assumed, with as little evidence, that
reduction of the free cysteine-cystine concentration by the formation of
Hg++ complexes would stimulate synthesis of these amino acids?
Toxic and Lethal Doses
A few of the results on different types of mercurial are summarized in
Table 7-22. The lethal dose will depend on the time interval chosen for de-
termination of the mortality, since death from mercurial poisoning may
occur several days following the administration, and this accounts for some
of the variability seen in the table. It is clear that the organic mercurials
are generally less toxic than HgCl,. It is difficult to estimate average doses,
but roughly the LD50 for HgClg is near 10 mg Hg/kg (0.05 millimole/kg)
and for the organic mercurials around 40 mg Hg/kg (0.2 millimole/kg);
there is so much species variation and differences between the organic mer-
curials that these figures are to be taken only as a crude basis for compari-
son. Differences between routes of administration are not as marked as one
might expect. One of the most important factors determining the toxicity of
mercurials given intravenously is the state of dissociation of the R — Hg — X
bond, where X represents any ion or thiol either introduced with the mer-
curial or present in the blood. In other words, the concentration of the free
R — Hg+ ion, which is able to react with the SH groups of the tissue cell
membranes, is a major toxicity determinant. If the mercurial is already
complexed with a thiol, as in mercaptomerin, the dissociation of the Hg — S
bond will be slow and little of the mercurial will be bound to SH groups,
in either the blood or the tissues. A second factor of undoubted significance
is the distribution of the mercurials, in both the R — Hg — X and R — Hg+
forms. An R group or a slowly dissociating Hg — X bond will favor penetra-
tion into the central nervous system in some instances, and this may alter
the pattern of toxicity from a rapid cardiovascular death to a slowly de-
veloping degeneration of certain nervous pathways.
Distribution, Metabolism, and Excretion of Mercurials
Certain aspects of the fate of the mercurials in the body have been dis-
cussed relative to the diuretic action, and these will be briefly summarized.
(1) All mercurials are accumulated in the kidney and reach much higher
concentrations in this tissue than in others, although the rate and degree
of accumulation depend on the structure of the mercurial. (2) Mercurials
are to a great extent bound to the plasma proteins and erythrocytes so
that only a small fraction is free to enter the tissues or be filtered through
the glomeruli. (3) Some of the mercurials may be secreted by the renal
tubular cells. (4) Mercurials are excreted in the urine mainly complexed
with thiols such as cysteine. (5) Some organic mercurials are split to form
Hg++ in the body, but there is no agreement as to the degree to which this
956
7. MERCURIALS
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958 7. MERCURIALS
occurs or whether it is important for the actions of the mercurials. (6) Hg+
is quite rapidly oxidized to Hg++ in the body and probably acts on the tis-
sues in the oxidized form.
The results of distribution studies are shown in Table 7-23. Further data
on the early distribution of several mercurials may be found in Table 7-19;
rough values for tissues levels of Hg++ in fatal human poisonings were pre-
sented in Table 1-8-1. Although quantitative comparisons are difficult due
to the widely different doses and the various routes of administration, the
general picture is clear. The concentrations in most tissues are not mark-
edly different from those in blood, but there is slight accumulation in the
spleen, moderate accumulation in the liver, and striking accumulation in
the kidney. The central nervous system levels are generally low, as expected,
and this must be due mainly to the small unbound fraction in the blood,
.since even the more lipid-soluble mercurials (e.g. MM) do not readily pene-
trate into the brain. Berlin and Ullberg (1963) gave single doses of Hg^^^Clg
intravenously to mice and determined the changing tissue levels over 16
days. The greatest amount in the central nervous system occurs in the brain
stem in the area postrema, in the hypothalamus, and in sites adjacent to the
lateral ventricles, and the retention in these regions is greater than in other
tissues. Essentially no Hg++ appears in the fetus so that the placenta pre-
sents a barrier to penetration, most of the Hg++ being bound to proteins
and the cellular elements of the blood. With the exception of the kidney
and brain there appears to be no obvious correlation between tissue levels
and pharmacological or toxic actions, and it is possible that the acute ef-
-fects, as on the heart, may be due to the initial binding to the cell mem-
branes rather than the result of intracellular uptake. Most of the results in
the tables were obtained with subtoxic doses, with the exception of those
of Galoyan and Turpaev (1958) and the cases of human poisoning, so it is
not possible to obtain a complete picture of the tissue levels during periods
of toxic reactions, but it is evident that rather low over-all concentrations
occur in most tissues. When it is considered that probably a major fraction
of the tissue mercurial is boiind to metabolically or functionally inert com-
ponents, it appears that very little mercurial is required to alter tissue
activity.
Loss of mercurials from the body by urinary excretion is usually slow.
Rothstein and Hayes (1960) determined the total body content of Hg^"*
in rats given small doses of HgCL over a period of 100 days, and found
three distinct phases: 40% is lost in 5-10 days, 45% more during the next
40-50 days, and not over 5% more in the next 50 days, so that at 100 days
there is still some 10-15% of the administered mercury in the body. When
jjg203Qj^ is infused intravenously for periods up to 4 hr in rabbits, it is found
that the renal excretion does not exceed 10% of the total amount of Hg^°^
passing through the kidneys. About 50% of the total dose is taken up in
EFFECTS OBSERVED IN THE WHOLE ANIMAL
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960 7. MERCURIALS
the kidneys (Berlin and Gibson, 1963). Less than 1% of the plasma Hg^"^
is filtered, due to both protein binding and the fact that nearly 50% of the
Hg203 is in the erythrocytes and is slowly exchangeable with the plasma.
Thus much of the Hg^''^ found in the kidney tissue must come directly from
the blood since the glomerular filtration cannot account for it. In the case
of the mercurial diuretics, around 20-40% is excreted during the first day
(Borghgraef and Pitts, 1956; Calesnick et at., 1960), but progressively smaller
amounts are lost each day. The kidney retains appreciable mercurial for
many days; following injection of only 0.2 mg Hg/kg of HgClg into rats, the
renal level is around 12.4 //g/g at 52 days (Rothstein and Hayes, 1960), this
constituting 90% of the body mercury. MM is surprisingly well retained in
the kidney, at 32 days the level being only 30% reduced from that after
1 day (Swensson et al., 1959). It is obvious that cumulation invariably oc-
curs when a mercurial is administered daily. Indeed, HgClg given intrave-
nously every 21 days leads to a marked cumulation, the total body mercury
after 5 doses being twice that from a single dose (Rothstein and Hayes,
1960). Rats given HgClg subcutaneously daily cumulate mercury in several
tissues and require 2 weeks for the rates of intake and excretion to be equal
(Friberg, 1956).
Normal human tissues contain mercury because there is a daily intake
in the food. Bread, flour, milk, pork, and beef contain 2-4 //g% mercury,
and certain vegetables a good deal more, depending on soil conditions and
sprays used (Szep, 1940). Forney and Harger (1949) reported a wide va-
riation in kidney mercury levels in normal human subjects (from 0 to 12.7
mg% in 92 autopsies) with two thirds having concentrations greater than
0.1 mg%. Liver levels were less (from 0 to 1.72 mg%) with values greater
than 0.1 mg% in one third. Those having received mercurial medication
ranged from 0.94 to 27.5 mg% mercury in the kidney. Similar results were
obtained by Griffith et al. (1954), the mean values in nonmercurialized cases
being 0.45 mg% in kidney, 0.10 mg% in liver, and 0.026 mg% in spleen
(these values are in terms of wet weight to compare with the results of
others, and were calculated from the dry weight figures with the data in
Table 1-8-3). Patients receiving large amounts of mercurials for some time
prior to death (mean of 4.7 g total) had much higher concentrations in the
kidney (3.4 mg%) and liver (0.36 mg%). It is very interesting that normal
human liver contains around 1 //g/g wet weight of mercury. From the values
for liver in Table 7-23 it is seen that in many cases the levels are lower even
though the animals had been given mercurials; since control concentrations
have seldom been obtained, it is questionable how much of the mercury in
most of the tissues is due to the administered mercurial and how much to
other sources. Of course, this does not apply to studies with Hg^*'^. It would
appear that human liver normally contains more mercury than the rodent
liver, but whether this is of dietary origin or a species difference is not known.
EFFECTS OBSERVED IN THE WHOLE ANIMAL 961
The metabolism of the alkyl mercurials and PM in the body is of some
importance in understanding the pattern of their effects. PM is one of the
least stable mercurials in the body and much of it is split to inorganic
mercury, little being retained in any tissue but the kidneys (Miller et al.,
1960; Gage and Swan, 1961). Very little MM, on the other hand, is split to
inorganic mercury; it is slowly excreted and cumulates in certain tissues,
such as the brain. Ethyl-Hg is also retained well by the tissues, but is ap-
parently split to inorganic mercury at a moderate rate (i.e., faster than MM
and slower than PM), so that by the seventh day only 21% of the total
mercury in the kidney is ethyl-Hg (Miller et al., 1961). It is odd that at the
seventh day all the mercury in the liver is ethyl-Hg, so that one concludes
that splitting does not occur in the liver. However, only 70% of the blood
mercury is ethyl-Hg, so it seems that the liver takes up some inorganic
mercury. It is possible, of course, that all the nonethyl-Hg mercury is not
inorganic mercury.
Toxicity to Aquatic Organisms
A discussion of the effects of mercurials on animals would not be complete
without mentioning briefly some of the interesting work done with marine
invertebrates, mainly in connection with antifouling programs, and with
fish. One would expect sea water not to be a favorable medium for the ac-
tion of Hg++ because of the high concentrations of complexing anions and
the elevated pH. The importance of the medium is apparent in the study
of the amphipod crustacean Marinogammarus marinus by Hunter (1949).
The minimal toxic concentration of Hg++ in sea water is 0.074 mM, while
in distilled water it is only 0.0093 vaM. Other factors, such as altered trans-
port activity, may contribute to the increased susceptibility. Hg++ at 0.18
milf does not depress the respiration of this organism, indicating that the
toxic effect is not to be attributed to a general metabolic inhibition.
Marine invertebrates often show marked changes in susceptibility to Hg++
during development. This is well illustrated by the results obtained on the
barnacle Balanus balanoides, the sensitivity to Hg++ reaching a minimum
during the free-swimming cyprid stage (see accompanying tabulation) (Pye-
(Hg++) for 50% lethality
(mM)
Nauplii
Stage III 0.00033
Stage IV 0.00085
Stage V 0.0011
Stage VI 0.0011
Cyprids 0.011
Barnacles 0.0026
962
7. MERCURIALS
finch and Mott, 1948). Although the cyprids are not killed so readily by
Hg+"'", their settlement on the substratum is reduced appreciably by 0.000037
mM and completely by 0.00018 mM. The lack of an open gut in the cyprids
is suggested as a possible explanation for the reduced sensitivity. Support
for the law that nothing is simple or predictable in the response of organisms
to metal ions, although no further support is needed, is the fact that the
cyprids are less sensitive to Hg++ in 50% diluted sea water relative to normal
sea water, in contrast to the amphipod discussed above. Another interesting
and complex phenomenon was discovered by Barnes and Stanbury (1948)
in studying the effects of Hg++ and Cu++ on the harpacticid copepod Nitocra
spinipes. Hg++ is over 1000 times more toxic than Cu++, but when these
metal ions are present together at certain concentrations the lethal effect
is greater than would be expected on the basis of their actions alone (Ta-
ble 7-24). An isobologram for 50% lethality provides a curve characteristic
Table 7-24
Lethal Effects of Mercury, Copper, and Their Combinations on Nitocra spinipes'^
Animals
killed in 24 hr
(%)
Hg++
(mM)
No Cu++
Cu++
Cu++
Cu++
Cu++
0.0041 mM
0.041 mM
0.41 mM
4.1 mM
0
0
1.3
11.3
21.2
42.5
0.00026
0
9.1
11.9
—
—
0.00056
1.4
14.5
20.0
78
—
0.0011
10.0
12.7
45.6
82
—
0.0015
16.7
50.0
93.7
98
—
0.0022
60
61.8
100
100
—
0.0026
72
76.4
100
100
—
0.0056
78
87.3
100
100
—
0.011
84
100
100
100
—
0.016
100
100
100
100
—
" From Barnes and Stanbury (1948.)
of very definite synergism (see Fig. 1-10-8), i.e., the curve is extremely con-
cave upward. It was postulated that lowered vitality due to one metal ion
may not allow the animal to deal effectively with the other metal ion. For
example, Hg++ might impair the excretory system so that Cu++ would be
retained, since it is known that certain crustaceans and mollusks excrete
Cu++. The synergistic effects indicate that Hg++ and Cu++ may act by dif-
EFFECTS ON MITOSIS, GROWTH, DIFFERENTIATION 963
ferent mechanisms, and possibly simultaneous attacks on different metabolic
or functional systems would be particularly toxic.
Jones (1946) studied the effects of several metabolic inhibitors on the
fresh-water stickleback Gasterosteus aculeatus. When Hg++ is present at 0.02
mikf, there is a temporary stimulation of respiration (+ 20-30%) at 10-20
min, followed by a depression that reaches 50% at 55 min, the fish surviv-
ing for 110 min. During the phase of respiratory increase, there is accelerat-
ed motility, a greater opercular activity, and a faster heart rate; it is quite
possible that the rise in respiration is associated with the greater functional
activity. If the Hg++ is removed after the respiration has been depressed
50%, recovery is slow and erratic, and the respiration never recovers its
normal level, although after 1 day the fish appear normal. The mechanism
by which the fish are killed is unknown but possibly it is asphyxial.
EFFECTS ON MITOSIS, GROWTH, AND DIFFERENTIATION
If SH groups are particularly important in cell cleavage, as many have
believed, the mercurials should be effective growth inhibitors and perhaps
useful agents to determine if these SH groups are enzymic or involved in
cytoplasmic structure. The rather potent inhibition of the proliferation of
many microorganisms by mercurials has been known for almost 100 years
and will be discussed in the following section, while here we shall attempt
to analyze the mechanisms by which mitosis of plant and animal cells is
disturbed by the mercurials.
Eggs and Embryos
Mercurials at concentrations in the range 0.001-0.01 mM usually inter-
fere with cleavage, even in sea water and despite the fact that much of the
mercurial in most work is removed from the medium because of binding.
Thus Mathews (1904) showed that the formation of embryos from Fundulus
heteroditus eggs is 50% blocked by 0.0048 mM Hg++, 90% blocked by
0.0095 mM, and completely blocked by 0.014 raM. The effects of mercurials
on eggs and embryos at different stages of development may be quite com-
plex. Hg++ may be parthenogenetic in that it induces membrane elevation
in Arbacia eggs and initiates a form of cleavage at 0.01-0.1 mM (Heilbrunn,
1925). The membranes begin to rise 3-5 min after addition of Hg++ and
after 12 min the cells may become constricted unequally or cleave, but the
relation of this to normal division is not clear. Hoadley (1930) studied these
effects more closely and observed that 0.025 mM Hg++ (a concentration
several times that suppressing cleavage completely) caused, after mem-
brane elevation, a clumping of the cortical pigment to one side of the egg,
followed by an unequal constriction which pinches off a small fragment
964 7. MERCURIALS
containing all the pigment, this fragment later cytolyzing. Hoadley claimed
that no true cleavage occurs and called the Hg++-induced behavior pseu-
docleavage. Both Heilbrunn and Hoadley felt that the Hg++ acts primarily
on the cortical region, and Hoadley, in addition, thought that the Hg++
may react with the pigment itself.
Kriszat and Runnstrom (1952) reported a strange phenomenon occurring
in Arhacia eggs treated with 0.028 mill p-MB. This concentration of mer-
curial rapidly inactivates the spermatozoa, but some fertilization can occur
before this is complete. Fertilization causes a strong contraction of the cor-
tical layer, squeezing out the cytoplasm into a number of pigment-free
lobes, the cortex, containing all the pigment, shrinking to a small folded
sac. It was postulated that p-MB blocks rather specifically those processes
reversing the surface contraction occurring during normal fertilization, it
is quite possible that these effects are exerted directly on the SH groups
of the protein components of the cortex (or plasma membrane, since it is
difficult to differentiate them), rather than on enzymes. The concentration
of mercurial is very critical. Hg++ at 0.02 n\M acting for 20 min on Arhacia
eggs prevents development beyond the early blastula stage, but acting for
6 min has no effect on motility or larvae; however, 0.025 vaM acting for
3 min reduces cleavage and interferes with development (Hoadley, 1930).
If the mercurial is added some time after insemination, the effects are
modified. Thus 0.05 vaM p-MB 30 min after fertilization scarcely interferes
with cleavage, the delay in onset being only 1-2 min and 95% of the eggs
dividing (Zimmerman et al., 1957). Mersalyl is less inhibitory and at 2 mM
the eggs cleave normally, although there is a 10-15 min delay; only 40%
develop to the blastula stage. PM, on the other hand, is very potent, indi-
cating possible permeability factors (Macfarlane and Nadeau, 1948). De-
velopment of Tripneustes esculentus (sea urchin) embryos, exposed at the
2-4 cell stage for 1 hr to 0.001 mill PM, is inhibited and only 2% reach a
motile blastula stage. Many of the embryos are abnormal and partial cytol-
ysis occurs. Even 0.00038 m3I PM slows yolk absorption although cleavage
is not affected. Echinus miliaris larvae exposed to 0.0005 mill Hg++ meta-
morphose, but there is dedifferentiation of the tissues so that the young
echinus is often abnormal, perhaps possessing rudimentary tube-feet or
spines (Huxley, 1928). Gastrulation is a process generally sensitive to toxic
substances and this is true for the mercurials. For example, frog dorsal lip
explants are depressed rapidly by 0.1 milf p-MB so that little further de-
velopment takes place (Ornstein and Gregg, 1952; Gregg and Ornstein,
1953). The mercurial seems to prevent certain movements and spreading
associated with gastrulation, e.g., the stretching of the mesoderm within
the endoderm and the ectodermal flow over the endoderm. Not much has
been done on later embryonic development, but p-MB injected into newborn
mice brings about varying degrees of neuroblastic necrosis, chiefly in the
EFFECTS ON MITOSIS, GROWTH, DIFFERENTIATION 965
outer cortical zones, this to some extent simulating radiation injury (Hicks,
1953).
There is no evidence that mercurials disturb development by generally
depressing metabolism. Ornstein and Gregg (1952) observed no effect of
p-MB on dorsal lip explant respiration at a concentration blocking differen-
tiation, and Brock et al. (1939) found that to inhibit sea urchin egg respira-
tion requires 20 times the Hg++ concentration necessary for cleavage block.
The latter concluded that this points to a nuclear effect as the basis for the
inhibition of division, but this does not follow. However, the difference be-
tween SH reagents is well shown here, in that arsenite depresses respiration
more readily than division. Haas (1941) also inclined to a primary nuclear
effect, since Hg++ produces demonstrable damage to the nucleus at 0.00074
vaM, while 0.037 mM is required for cytoplasmic damage in Anodonta
(fresh-water clam) eggs. Without denying that Hg++ can damage the nu-
cleus, one must be pessimistic as to the reliability of determining the site
of action of an inhibitor by visual inspection; e.g., the action could have been
on the plasma membrane and be microscopically undetectable, or a good
deal of disturbance in the cytoplasm might have been caused without being
immediately evident. Landau et al. (1954) stated that mersalyl is an effec-
tive ATPase inhibitor and hence was tried on the cleavage of Arbacia and
Chaetopterus eggs. Fertilized eggs placed in 2 mM mersalyl complete the
first 3-4 cleavages, but the furrowing strength is reduced, as measured by
the pressure increases required to prevent furrowing, so an inhibition of the
gelation of the cortex in the equatorial region by mersalyl was postulated,
this presumably being mediated through an interference with ATP utiliza-
tion. Since the ATPases from different sources vary a good deal in sensi-
tivity to mercurials, one does not know what inhibition to expect in these
eggs, and the mercurial concentration is so extremely high that many me-
tabolic and functional processes must be affected. Heilbrun and Wilson
(1955) explained the block of cleavage by mercurials as an inhibition of the
proteolytic enzyme system involved in gelation, without obvious evidence.
The direct effects of Hg++ on fibrous proteins extracted by sea urchin eggs
by Sakai (1962) are meaningless because a concentration of 10 mM was
used.
Plants
Many organic mercurials are applied to seeds, bulbs, or plants as fungi-
cides, but occasionally the plant tissues may be damaged and growth de-
pressed. The persistence of the mercurial in the plant is often remarkable.
For example, carnation seeds treated with radioactive PM at a concentra-
tion causing growth abnormalities in the seedlings produces plants which
at 8-9 weeks contain the mercurial in the cotyledon leaves, the hypocotyl,
and the root adjacent to the hypocotyl (Robson and Fenn, 1961). A very
interesting effect, and one illustrating that the mechanisms by which mer-
966 7. MERCURIALS
curials act may often be unexpected, is the zinc-deficiency disease of coffee
trees in Kenya due to spraying with mercurial fungicides (Bock et al., 1958).
Not only do the plants exhibit typical signs of zinc deficiency — chlorosis,
abnormal growth of shoots and leaves, and short internodes — but the zinc
content is reduced to 25% of normal. In the promotion program for mer-
bromin, Macht (1931 a) purported to show that organic mercurials are less
toxic than Hg++ to plants, but his data are equivocal, since Hg++ is toxic
around 0.1 mM, while merbromin inhibits growth slightly at 0.0013 mM,
50% at 0.043 mM, and 81% at 1.29 mM. The growth of Avena coleoptiles
and of pea stems is readily inhibited by the mercurials, PM being much
more potent than p-MB (see accompanying tabulation) (Thimann and Bon-
Concentration for
Tissue
Mercurial
50% inhibition
(mM)
Avena coleoptile
p-MB
0.035
PM
0.007
Pea stems
p-MB
0.4
PM
0.02
ner, 1949). It is likely that the carboxylate group prevents the p-MB from
penetrating as well as PM. These results will suffice to demonstrate growth
inhibition by the mercurials, and we shall turn to what little evidence is
available for the mechanisms involved.
Onion roots exposed to 0.0075 mM PM develop terminal swellings and
growth is immediately stopped; however, after a day new growth starts
distal to the swelling (Macfarlane and Nadeau, 1948). Doubling the con-
centration leads to 100% mortality of the roots. Hg++ even at 0.05 mM
does not cause terminal swelling, inhibits growth only 15%, and does not
kill any of the roots. Macfarlane (1951) pointed out that PM acts on onion
roots cytologically like colchicine, in that spindles are abnormal, chromo-
some movement is impeded, and polyploidy results in the zone of cell en-
largement proximal to the meristem. In addition, there is chromosome
stickiness, fragmentation, and aggregation. Although mitotic and chromo-
somal disturbances certainly occur, there may be some question as to the
validity of terming these effects mutagenic or radiomimetic (Macfarlane,
1953). Meyer (1948) also observed such changes in the root tips of Crepis
capillaris exposed to 0.013 mM Hg++, the sister telophase nuclei often being
connected by chromatin bridges, with some breakage and recombination,
leading to 80% diploid metaphases and 1.5% tetraploid metaphases. The
formation of cell wall material in the microspores of excised Lilium henryi
anthers is reversibly blocked by 0.01 mM p-MB and the progress of meiosis
EFFECTS ON MITOSIS, GROWTH, DIFFERENTIATION
967
is slightly retarded (Pereira and Linskens, 1963). Similar chromosomal
changes induced by mercurials have not been reported, as far as I know,
for animal cells. The growth stimulation by auxin applied to Avena coleop-
tiles is inhibited by p-MB at 0.3 raM (Cleland and Bonner, 1956), but the
effects on auxin transport in sunflower stem section are complex in that
0.01 mM p-MB accelerates transport 125%, 0.1 mM depresses it 25%, and
1 mM blocks it completely (Niedergang-Kamien and Leopold, 1957). It is
not known if interference with auxin transport or action is involved in
growth inhibition.
The structure-action relationships of mercurials acting on the sporelings
of the marine red alga Flumaria elegans, reported by Boney et al. (1959),
were believed to demonstrate the importance of lipophilicity and penetra-
tion (Table 7-25). The alkyl mercurials are often 200-300 times more toxic
than HgClg, the branched chain compounds being less toxic than the straight
Table 7-25
Lethal Concentrations for Plumaria Sporelings Exposed
TO Mercurials for 18 Hr."
Mercurial
Concentration for
50% lethality
(mM)
Potency
relative to HgCl.
0.0115
1
0.000344
33
0.000176
65
0.000097
119
0.000046
250
0.000043
268
0.000041
280
0.000099
116
0.000060
192
0.000173
67
0.000260
45
HgCl,
Hgl,
Methyl-HgCl
Ethyl-HgCI
n-Propyl-HgCl
n-Butyl-HgCl
«-Amyl-HgCl
Isopropyl-HgCl
Isoaniyl-HgCl
Phenyl-HgCl
Phenyl-Hgl
« From Boney et al. (1959).
chain. Some correlation between potency and the distribution ratios be-
tween ether and water, and between methyloleate and water, was claimed,
but discrepancies exist. Similar relationships have been reported for certain
marine crustaceans (e.g., Artemia salina), but in others (e.g., Acartia clausi)
there is little difference in toxicity between the mercurials. Inasmuch as
968 7. MERCURIALS
there has been no work on the relative inherent or direct toxicities to cel-
lular processes, metabolic or functional, or adequate comparison of their
abilities to react with relevant SH groups, it is impossible to be certain
that the differences are due solely to variation in the penetration. Indeed,
one is not sure that the mercurials all act by the same mechanism. One
factor which is often ignored is the role of the size of the side chain in
membrane processes, assuming that they all react with identical SH groups
in the membrane. Nevertheless, such quantitative studies are valuable in
establishing a necessary basis for understanding the mechanisms by which
the mercurials act; further work will undoubtedly allow these results to
be interpreted more readily.
Mammalian Cells and Tissue Cultures
We must note that growth, in common with many other processes, may
be accelerated by low concentrations of the mercurials, as noted by Hira-
shima (1934) in chick fibroblast cultures, and by others. This has also been
reported for plant tissues, p-MB at 0.001-0.005 mM stimulating the growth
oi Arena coleoptiles some 20-25% (Thimann and Bonner, 1949). It is inter-
esting to recall that Mallus (1931) gave HgClg at 0.25-0.3 mg/kg to atrophic
but otherwise healthy children and observed certain changes — increased
chest measurements, rise in erythrocyte count and hemoglobin, and eleva-
tion of urinary nitrogen — indicating a stimulation of growth and metabo-
lism. The mechanisms by which mercurials can stimulate growth are un-
known, but it is possibly not a specific action since many types of cells ex-
hibit an increased proliferative activity when disturbed slightly by irritant
substances (our terminology in this field is admittedly inadequate).
Inhibition of growth is invariable when the mercurial concentration is
increased beyond a certain level, which is frequently quite low. Many of
the experiments with tissue cultures have been done of necessity in complex
media (e.g., embryo extract) which must bind a large fraction of the mer-
curial, so the true inhibitory potency must be much greater than is indi-
cated by the concentrations used. Fibroblastic and leucocytic migration is
depressed 50% by Hg++ at concentrations near 0.08 mM, and growth is
somewhat modified even at 0.0037 mM (Meier, 1933). Chick embryo heart
cultures fail to grow in 0.08 mM Hg++ (Salle and Lazarus, 1936) and here
the organic mercurials are less toxic (Salle, 1943). Pulsations of the cardiac
cells are stopped by Hg++ before growth is affected, but the organic mer-
curials (thimerosal and nitromersol) stop growth and cause cytolysis with-
out previously interfering with the contractile activity. The concentration
of Hg++ for 50% inhibition of Eagle's KB strain of human carcinoma cells
is 0.037 mM (Smith et al., 1959), and 75% reduction in the mitoses of mouse
ear epidermis requires 0.01 mM p-MB (Gelfant, 1960). We may thus con-
clude that mercurials at concentrations around 0.005-0.05 mM seriously
EFFECTS ON MITOSIS, GROWTH, DIFFERENTIATION 969
restrict growth of mammalian cells in culture, higher concentrations usually
killing the cells directly'. If the mechanism of this growth inhibition is to be
elucidated and correlated with metabolic alterations, it will be necessary
to work within this range if the conclusions are to be valid.
Some General Aspects of the Effects of Mercurials on Mitosis and Growth
A few general comments on the localization of the sites of growth inhi-
bition were made on page 1-531 and reference to these will make it evident
that at the present time we have little hope of explaining the actions of the
mercurials. Essentially nothing is known of the possible effects of mer-
curials on protein, nucleic acid, or coenzyme synthesis, or whether the
demonstrated inhibitions of active transport are in any way related to the
growth depression. We have seen that respiration is not significantly de-
pressed during growth inhibition in the few instances in which it has been
determined; however, in view of the rather potent inhibition of the cycle,
it would be worthwhile to pursue this question further. Mercurials are not
efficient uncoupling agents and could scarcely act in this way. The signifi-
cance as a possible metabolic mechanism of mercurial action of the obser-
vation by Hirashima (1935), that glucose reduces the toxicity of Hg++ for
fibroblast cultures, cannot be evaluated.
A direct action on the sol-gel transformations and protoplasmic move-
ments during furrowing, spindle formation, and cleavage has been postulat-
ed and discussed briefly in previous sections. Mazia (1959) believes that the
mitotic apparatus may be an S — S bonded structure because of the ability
of agents splitting S — S bonds to dissolve the structure, and states that
p-MB and mersalyl bring about the dissolution of the freshly isolated spindle.
The question is whether such an action can be exerted at the concentrations
occurring within cells during mitotic inhibition. We have also mentioned
that several workers favor a nuclear site for the mercurials but that the
evidence is insufficient, as is that of Meyer (1960), who showed that the
conidia of Fusarium decemcellnlare incubated with 0.037 mM Hg++ accu-
mulate mercury in some form either on or within the nucleus, since there
is no necessary correlation between relative intracellular concentrations and
the site of action. It is worth noting that p-MB interferes with the synthesis
of RNA from nucleosides, as determined by the uptake of cytidine into the
nuclear RNA of HeLa cells, and an inhibition of the RNA polymerase was
suggested (Srinivasan ct al., 1964). HeLa cells are blocked in metaphase by
0.02 mM 7)-MB. In connection with the selective accumulation of mercurials,
it is worth noting that merbromin appears to be localized in tumor tissues
of both mouse and man, as indicated by fluorescence several days after
initiation of intravenous or oral administration (Katsuya et al., 1963). Al-
though kidney exhibits the highest concentration of mercurial initially, it
and other normal tissues lose the merbromin much faster than tumors. It is
970 7. MERCURIALS
somewhat surprising that tumors could contain a component holding mer-
curials more tightly than in normal tissues, or at least more of a component
binding the mercurial strongly, but this would support ideas which have
been advanced relative to the abnormal state of SH group-containing sub-
stances in tumor cells.
EFFECTS ON THE GROWTH OF MICROORGANISMS
Mercurials have been used for years to control the growth of many types
of microorganism, invertebrate, and plant, and have been applied commer-
cially as fruit sprays, paint preservatives, mothproofers, grain insecticides,
anthelmintics, as well as antiseptics and disinfectants, and in antifouling,
crab grass control, bacterial plant diseases, and nematode control. One of
the most important commercial uses at the present is as fungicides in the
treatment of seeds, fruits, and plants, and for the most part certain organic
mercurials have been developed for this purpose. The clinical use of mer-
curials as antiseptics, first popularized by Koch (1881), has dechned some-
what due to the discovery of generally more selective antibacterial agents,
but most of the experimental work has been done with this group of aro-
matic mercurials. Inasmuch as the relation between these actions and met-
NO,
\^ /^ Hg NO3
H3C
Phenylmercuric nitrate Nitromersol
(Merphenyl nitrate, Merphene) (Metaphen)
HgOH
Merbromin Thimerosal
(Mercurochrome) (Merthiolate)
abolic interference is vague, only a cursory treatment of the mercurials
as inhibitors of microorganism growth will be given.
EFFECTS ON THE GROWTH OF MICROORGANISMS 971
Bacteria, Fungi, and Yeast
Koch originally claimed that Hg++ possesses the ability to kill various
bacteria and their spores, but it was soon shown by Geppert (1889) that
the proliferative activities of the treated bacteria can be restored by remov-
ing the Hg++ with sulfides, and that the action of the Hg++ is primarily
bacteriostatic, a concept confirmed many times and extended to other or-
ganisms. Dilution or washing is not sufficient to extract the Hg++ bound
to the cells (Chick, 1908), but sulfides and various thiols (Fildes, 1940) can
readily reverse the bacteriostatic action. With high enough concentrations
and prolonged exposure, of course, bacteria may be killed, especially at
elevated temperatures. Yeast cells incubated with 0.93 mM Hg++ are killed
progressively over an hour as determined by staining with methylene blue
or Congo red (Rahn and Barnes, 1933). One of the first effects noted at
minimal concentrations of the mercurials is a prolongation of the lag phase
of proliferation (Cook and Steel, 1959). Increasing the concentration of mer-
curial progressively delays and slows the growth, and eventually stops it,
at which point the cells can remain viable often for quite extended periods
of time, as exhibited by the renewal of proliferation when the mercurial is
removed with a thiol.
The sensitivities of various microorganisms to the mercurials are shown
in Table 7-26, in which only a few of the reported results have been pre-
sented because the purpose is mainly to illustrate that proliferation is
usually depressed at fairly low concentrations, and that the data often vary
due to the conditions of the testing. One of the most important factors is
the medium used, inasmuch as most growth media contain many substances
capable of complexing with the mercurials and reducing their effective
concentration. Claus (1956) demonstrated that the minimal inhibitory con-
centration of Hg++ varies over a 10-fold range depending on whether pep-
tone media or simple nitrogen sources are supplied, and Cook and Steel
(1959) also presented evidence that the usual culture media exert a pro-
tective action. Another important factor is the temperature, increase in
temperature markedly enhancing the bacteriostatic effect (Cianci, 1940;
Cook and Steel, 1959), the Q^q being 3-4 (Chick, 1908). There is unquestion-
ably species variation in susceptibility but no correlation with bacterial
metabolic or growth habits has been made. It should be noted that very
low concentrations of the mercurials occasionally stimulate growth, this
being perhaps more true for fungi than bacteria. Robertson (1943) noted
that a mercurial used for preserving leather actually accelerated the growth
of certain fungi within a particular concentration range, and Converse and
Besemer (1959) reported that p-MB at 0.00028 mM stimulates the growth
of Coccidioides immitis, although spherulation is definitely inhibited. This
phenomenon would probably be seen more commonly if low concentrations
of the mercurials were more frequently tested, and clinically and commer-
972
7. MEECURIALS
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EFFECTS ON THE GROWTH OF MICROORGANISMS
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974 7. MERCURIALS
cially it might be an important aspect of their use. Despite the many com-
parisons of the relative activities of different mercurials, very few interesting
correlations between structure and effectiveness have emerged. Krahe (1924)
found that the bacteriostatic activity of Hg++ is reduced by increasing the
concentration of NaCl and postulated this to be due to the formation of
HgClg"" and HgCl4= complexes, these being less lipid-soluble, and showed
that the distribution coefficient between ether and water is reduced parallel
with the antibacterial potency. Coleman et al. (1937) observed in the ali-
phatic mercurials that the antibacterial activity increases with the length
of the side chain. Such relationships have been found for many actions of
the mercurials and are probably based on differences in penetration into
the cells rather than to fundamental differences in action on the susceptible
cellular mechanisms.
The uptake and distribution of mercurials have been well studied and
several facts relevant to the mechanism of their action have emerged. The
amount taken up in any case will depend on the relative quantities of cells
and mercurial present. Herzog and Betzel (1911) incubated 10 g of pressed
yeast (2.6 g dry weight) with various concentrations of Hg++ and found
the cellular Hg++ concentration to increase with the total amount of H.g++
present, but the percentage taken up falls (see accompanying tabulation).
Total Hg++ present
(g/100 ml)
Hg++
taken up by yeast cells
Total
(g)
Concentration
(g/g dry weight)
0/
/o
0.092
0.077
0.030
84
0.460
0.168
0.065
37
0.921
0.219
0.084
24
1.341
0.304
0.117
16
3.683
0.449
0.173
12
Since there are roughly 10^'' cells in 1 g of dry yeast, these uptakes would
correspond to between 2 x 10* and 10® Hg++ ions/cell; they would also
correspond to 0.065-0.38 g Hg++/g yeast protein of molecular weight 100,000.
The amount of Hg++ accumulated by yeast is thus very considerable and
only a fraction is likely to be bound to SH groups. E. coli binds even more
Hg++, since Hahn and Remy (1922) found an uptake of around 0.5 g Hg++/g
dry weight (assuming around 75% water content) from a 3.7 mM solution.
McCalla and Foltz (1941) calculated that E. coli possesses around 10* bind-
ing sites/cell, which is close to the figure to be estimated from the uptake
found by Hahn and Remy. Steel (1960) claimed that a cell of E. coli con-
tains about 10* SH groups, but that Hg++ does not react with all of them,
EFFECTS ON THE GROWTH OF MICROORGANISMS 975
p-MB giving a much higher accumulation. It is more difficult to determine
the distribution of Hg++ within the cells. Siipfle (1923) treated anthrax
bacilli with Hg++ and then with HgS, and showed black granules within the
cells. However, the Hg++ enters much more slowly than the antibacterial
action develops, which might be used as evidence for a primary membrane
site. Ruska (1947) examined Hg++-treated streptococci and E. coli with
the electron microscope and observed mercury in the membrane and dif-
fusely distributed in the cytoplasm, but most in small globular masses be-
tween the membrane and the cytoplasm. By a similar technique, Harris
et al. (1954) found no mercury in the membrane or cell wall of E. coli —
confirming the absence of electrophoretic change in cells treated with Hg++
— and most deposited as granules within the cytoplasm. Troger (1959)
localized mercury by the diphenylcarbazone method and found accumula-
tion in certain areas. It should be noted that visualization either electron
microscopically or histochemically is difficult in bacteria and, furthermore,
that generally quite high concentrations have been used so that the pattern
of distribution cannot apply directly to the bacteriostatic situation.
The possible mechanisms for the bacteriostatic action of the mercurials
have been debated for years and many theories have been proposed without
benefit of experimental evidence. The amount of valuable work on the mer-
curials from the standpoint of basic actions is almost negligible, due prob-
ably to the fact that when bacterial metabolism and proliferation began
to be investigated seriously, attention was turned to the sulfonamides and
antibiotics. From Tables 7-13 and 7-17 one might conclude that metabolism
must certainly be depressed in some manner during the action of the mer-
curials on bacteria, and this may well be in many cases, but Yamada and
Yanagita (1957) showed quite conclusively that the growth of staphylococci
is 140 times and 57 times more sensitive than respiration to thimerosal and
Hg++, respectively. Indeed, it is possible to stop growth essentially com-
pletely without affecting respiration significantly. Despite the lack of critical
experiments in other organisms, it is safe to say that the mercurials do not
inhibit the growth of microorganisms by simply suppressing oxidative proc-
esses and the supply of energy for growth and division. Fildes (1940) on
the basis of irrelevant evidence concluded that the antibacterial mechanism
is based on reaction with SH groups, and the impression is gained that he
was thinking of the smaller thiols rather than proteins and enzymes. Lou-
reiro and Lito (1946) put this theory on a better basis by demonstrating
some correlation between the fraction of bacterial SH groups reacted and
the bactericidal activity. However, even here all one can do is to increase
the mercurial concentration so that more and more SH groups are reacted,
and it is not surprising that more and more cells are inhibited or killed;
one cannot say what fraction of SH groups should be reacted before an
effect on the bacteria is observed, and indeed it is very unlikely that it
976 7, MERCURIALS
requires a 1 : 1 ratio of mercurial to SH groups, as assumed by these work-
ers. Nevertheless, in the face of no negative evidence, it is felt that, the
mercurials do inhibit growth by reacting with some SH groups — the prob-
lem is with what SH groups, since there are many different SH-containing
substances in the cell. Do the mercurials inactivate some SH enzyme, or
enzymes, involved in an important metabolic pathway, or react with SH
groups in the membrane to block active transport of necessary substances
into the cell, or alter permeability so that intracellular components are lost,
or directly stabilize the membrane to prevent division, or interfere with the
utilization of ATP, or disturb metabolism by reacting with some thiol co-
enzyme ? Experiments showing that certain substances protect against mer-
curials are not easy to interpret. Thus Pershin and Shcherbakova (1958)
found that histidine, glutamate, methionine, and particularly thiamine pro-
tect E. coli against Hg++, and interpreted this as indicating that the metab-
olism of these compounds is interfered with by Hg++, but it is also possible
that the protection is simply due to complexes formed with the Hg++.
Theories involving various physicochemical properties of Hg++ and other
heavy metal ions — such as solution pressure, solubility products, electro-
negativity, and ionization potential (e.g., Shaw, 1954; Somers, 1959) —
do not warrant serious consideration since they simplify the biological
system beyond recognition and, even if true, would not help us appreciably
to understand how the mercurials act.
Viruses
Most viruses and phages can be inactivated by the mercurials but it
requires fairly high concentrations relative to those inhibiting bacterial
growth (Table 7-27). In most work a virus suspension is incubated with the
mercurial for a certain period and the infectivity is then tested. As with
the effects on microorganisms in general, the degree of inactivation by the
mercurials depends on many factors, particularly the medium in which the
virus is suspended, the temperature, and the exposure time. The rates of
inactivation are quite different for various viruses: ECHO 7 virus is 50%
inactivated by 0.05 roM p-MB in 1 min and 99% inactivated in 6 min
(Choppin and Philipson, 1961), whereas tobacco mosaic virus is not com-
pletely inactivated after 24 hr exposure to 18.5 mM Hg++ (Kassanis and
Kleczkowski, 1944). This would be expected since the virus SH groups
must vary widely in reactivity as do the SH groups of proteins in general.
The inactivation is first order with respect to virus. Staphylococcus phage
infectivity declines exponentially when exposed to Hg++ according to the
equation:
dPIdt - kiRgCU) (Po - Pi)
where Pq is the phage initially present and P, the inactivated phage (Krueger
and Baldwin, 1933). This equation holds fairly well over most of the range.
EFFECTS ON THE GROWTH OF MICROORGANISMS
977
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EFFECTS ON THE GROWTH OF MICROORGANISMS
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980 7. MEKCURIALS
but there appears to be a small resistant fraction which remains infectious
over several days (Krueger and Baldwin, 1934). Similar kinetics of inactiva-
tion have been reported by Moriyama and Ohashi (1941) for E. coli phage,
and by Allison (1962) for fowl plague and vaccinia viruses.
The mercurials usually do not destroy the viruses or produce irreversible
structural changes in them, since reactivation with thiols has been observed
with staphylococcus phage (Wahl, 1939), influenza virus (Klein et at., 1948;
Perez et al., 1949), psittacosis virus (Burney and Golub, 1948), vaccinia
virus (Kaplan, 1959; Allison, 1962), streptococcal phage (Kessler and
Krause, 1963), and various enteroviruses (Choppin and Philipson, 1961). The
results of Krueger and Baldwin (1933, 1934) are particularly impressive;
reactivation with sulfide occurred even after exposure of phage to around
100 mM Hg++ for 9 days at 22o. It is remarkable that viruses, like certain
enzymes, can be reactivated readily with thiols (especially dimercaprol)
although no reactivation occurs by washing or dilution. Dimercaprol is
able to reactivate influenza virus in vivo when injected after the treated
virus in animals (Klein et al., 1948) or chick embryos (Perez et al., 1949).
The reactivation with thiols does not prove that the mercurials react
with virus SH groups, as has been concluded, but there is evidence for the
importance of SH groups. The relative resistance of tobacco mosaic virus
to the mercurials is probably due to the unavailability of the SH groups,
since Anson and Stanley (1941) showed that p-MB reacts with all the SH
groups of denatured virus but does not inactivate native virus. Some steric
factor preventing reaction with p-MB was postulated by Fraenkel-Conrat
(1959), since MM reacts stoichiometrically in a 1:1 ratio with the SH
groups. The restriction may be imposed by hydrogen bonding to adjacent
groups. Some plant viruses are structurally altered by mercurials. Solutions
of potato virus X lose their flow birefringence when treated with p-MB, and
sedimentation studies indicate disintegration into subunits (Reichmann and
Hatt, 1961). It was concluded that the SH groups occur near the linkage
sites holding the units together, rather than participating in the linkage,
and that the bulky p-MB molecule splits the links. Turnip yellow mosaic
virus is also split into subunits by p-MB, and RNA is liberated simultane-
ously (Kaper and Houwing, 1962 a). The artificial top component (empty
virus protein shells) binds 645-660 molecules of mercurial per particle. As
structural changes occur, new SH groups are unmasked and react with
p-MB (Kaper and Houwing, 1962 b). Finally, one must consider the reac-
tion of mercurials with the nucleic acid components of the viruses, since
such complexes have been established (Katz, 1962). Tobacco mosaic virus
RNA complexes with Hg++ (Katz and Santilli, 1962 b) but much of the
infectivity remains in this case, although retention of specific infectivity
was not demonstrated (Katz and Santilli, 1962 a).
We shall now inquire into the particular phases of virus multiplication
EFFECTS ON THE GROWTH OF MICROORGANISMS 981
inhibited by the mercurials. It is clear that there is little or no selective ac-
tion on viruses growing in vivo, and the mercurials have not been found to
be effective virucidal or virustatic agents. Thus, although p-MB depresses
psittacosis virus formation in chick embryo cultures, it also inhibits tissue
growth, and it is quite possible that the effect on the virus is secondary to
that on the host cells (Burney and Golub, 1948). Corneal infections with
herpes virus are not benefited by application of p-MB, although the virus
is readily inactivated in vitro (Sery and Furgiuele, 1961), and mercurials
are not effective in preventing or treating plant virus infections. Kaplan
(1959) concluded that mercurials react with SH groups on the surface of the
virus and thus prevent attachment to the host cell. Certainly the hemagglu-
tinating activity and the adsorption onto erythrocytes are depressed along
with the infectivity (Choppin and Philipson, 1961). Adsorption of entero-
viruses onto renal cells may also be reduced, but p-MB does not prevent
adsorption of influenza virus onto the chorioallantoic membrane. Further-
more, p-MB does not prevent infection of E. coli by phage but inhibits the
proliferation. Allison (1962) holds that the mercurials do not affect the
primary attachment of the virus to the host cell, but may prevent the un-
coating of the virus, an event which precedes multiplication. The studies
of Shug et al. (1960) on T2 phage show that here the inhibition by p-MB is
exerted early in the development, the maximal inhibition being 10-20 min
after infection. This excludes the energy-yielding host metabolism as a pri-
mary site of action, since the energy requirements are greater after 25 min
when the phage is being synthesized and assembled. They conclude that
the site of attack is a protein concerned with the initial phases of replica-
tion and possibly involved in the assembly of the components into an in-
tact phage. It would not be surprising if the site of mercurial action, or
the phase disturbed, is different for the various phages and viruses.
Protozoa
Ciliates are immobilized and killed by the mercurials but the results re-
ported are quantitatively discrepant. Nuhaus (1910) found that 0.15 milf
Hg++ paralyzes paramecia in 50 min and kills them in 70 min, but Wood-
ruff and Bunzel (1910) stated that 0.175 mM Hg++ stops all motion within
2 sec. Even greater sensitivity was reported by Gause (1933), as indicated
in the accompanying tabulation. It should be noted that actually death
Hg++ (milf) Duration of life (sec)
0.01
1080
0.015
360
0.02
116
0.03
30
982 7. MERCURIALS
was not the criterion, but cessation of movement, since no attempt to reac-
tivate was made. When the log of survival time was plotted against log
(Hg++), two linear segments were obtained, which led Gause to conclude
that two different processes are responsible for the paralysis. The treat-
ment is based on an equation of the type 1-12-89 and plotting as in Fig. I-
12-38. A break in the curve would suggest a change of the exponent n and,
since it is possible for one process or response to exhibit different values of
n, it is not necessary to conclude that Hg++ kills by two processes. High
sensitivity of paramecia to Hg++ was also noted by Calcutt (1950), who
found 0.001 raM to paralyze within 6-14 min depending on illumination.
Paralysis of Colpidium colpoda occurs in 3 min after exposure to 0.00087
mM PM (Walker, 1928). Some of the variations in sensitivity are due to
the strains used, and probably some to the media in which the ciliates were
suspended. It is probably safe to say that ciliary movement is very sensitive
to mercurials and is stopped within a few minutes by concentrations in the
range 0.001-0.01 mM. Reversal of ciliary beat by Hg++ is not observed
(Oliphant, 1942). The classic death rate curves for Colpidium exposed to
0.2 mM Hg++ and their relation to population variation were discussed
previously (page 1-593 and Fig. 1-12-36).
The effects of the mercurials on ameboid movement are interesting be-
cause of the possible bearing on muscle contractility. Reznikoff (1926) used
rather high concentrations of Hg++ and consequently found only a pinch-
ing off of the region into which the Hg++ was injected (0.62 mM), or a
break in the membrane (up to 10 mM), or an immediate gelation or coagu-
lation of the protoplasm (up to 200 mM). Kappner (1961) used mersalyl
since this mercurial has been a favorite with myologists, and the changes
he observed in amebas with increasing concentration are worth describing
briefly. With 0.001 mM mersalyl there are no immediate changes but after
several days some damage is evident. At 0.01 mM there is restriction of
normal pseudopodial response and fewer pseudopods are formed, while
clumping of the cytoplasmic crystals occurs. At 0.1 mM the pseudopods
withdraw, the ceUs soon form numerous small pseudopods at the end of
which appear tiny spheres, the cortex appears to be thicker, and eventually
the cells round up. At 1 mM the response is not so specific, the surface
bubbles, the cells round up, and soon the membrane dissolves. Higher con-
centrations produce vacuolization and cytolysis. Abe (1963) reported a sim-
ilar study but with p-MB to which amebas seem to be more sensitive than
to mersalyl, since 0.1 mM causes cytolysis within 8 min. Further investiga-
tion of this interesting problem is warranted and a closer analysis of the
effects of low concentrations on the sol-gel transformation might provide
useful information on the role of SH groups in protoplasmic movement.
DEVELOPMENT OF KESISTANCE TO MERCURIALS
983
DEVELOPMENT OF RESISTANCE TO MERCURIALS
Most types of microorganism appear to be able to adapt to the presence
of mercurials, but usually not as readily or to such a degree as to arseni-
cals, sulfonamides, or antibiotics. Some resistance factors are given in the
accompanying tabulation, but it is likely that greater tolerance could have
Organism
Mercurial
Resistance
factor
Reference
Staphylococcus aureus
Hg++
>50
Benigno and Santi (1946)
1.8
Klimek et al. (1948)
Escherichia coli
PM
2.6
Akiba and Ishii (1952)
Salmonella pullorum
Hg++
12
Severens and Tanner (1945)
Salmonella typhosa
Hg++
6
Severens and Tanner (1945)
Penicillium notatum
Hg++
2.5
Partridge and Rich (1962)
Sclerotinia fructicola
Hg++
2.5
Partridge and Rich (1962)
Yeast
Hg++
>10
Imshenetsky and Perova (1957)
Candida, utilis
Hg++
7
Gerardin and Kayser (1959)
Treponema pallidum
Hg++
75
Nogiichi and Akatsu (1917)
been developed in some instances if training had been prolonged. There are
also naturally occurring resistant strains and species. An interesting exam-
ple is the relative tolerance of Penicillium roqueforti to PM. and this has
bearing on the preservation of groundwood pulp (Russell, 1955). Most fungi
fail to grow in 0.006-0.030 mM PM, but this species grows well in a con-
centration of 0.06 mM and furthermore accumulates sufficient mercurial
to allow the less resistant organisms to grow. The number of serial cultures
in increasing mercurial concentrations required to produce tolerance varies
with the organism: It was 20 transfers for the fungi in the above table
(Partridge and Rich, 1962), 70-100 transfers for the species of Salmonella
(Severens and Tanner, 1945), and up to 500 transfers for yeast (Imshenetsky
and Perova, 1957). Occasionally no transfers are required and the organism
begins to grow normally after a prolonged lag period, as is the case with
Aspergillus glaucus where hyphal inoculations fail to grow for periods of
up to 14 days in 0.033 mM Hg++, and then proliferate without loss of vigor
(Briault, 1956). This indicates that resistance can develop in nonprolif crat-
ing organisms.
Inasmuch as we do not understand how the mercurials depress growth,
it is clear that we cannot immediately postulate logical mechanisms for the
developed resistance. However, some interesting observations may contrib-
ute to the elucidation of the mechanisms of inhibition. The resistance is
apparently not due to a reduction of permeability to the mercurials, as
984
7. MERCURIALS
occurs with the arsenicals, since Benigno and Santi (1946) found that
staphylococci tolerant to Hg++ grow when they have taken up much more
Hg++ than is required to prevent growth of the normal strain. The situation
with respect to the thiol content of the resistant organisms is confused,
since Akiba and Ishii (1952) showed that E. coli tolerant to PM have less
SH groups than normally, and Gerardin and Kayser (1959) found that tol-
erant Candida utilis contained 6 times more SH groups than the normal
strain. Zambonelli (1958 a,b) has obtained evidence that adapted yeast
produces more HgS and believed that this inactivates much of the Hg++.
Normal yeast produces HgS only from sulfite, whereas adapted strains pro-
duce it from sulfate and hyposulfite in addition, although not from cysteine
or glutathione. If the resistant strains are grown with only cysteine or glu-
tathione as the source of sulfur, Hg++ readily inhibits their growth. If
metabolic changes occur during adaptation, they are not marked. Thus
resistant Candida respires normally (Gerardin and Kayser, 1959) and re-
sistant yeast ferments glucose at the normal rate, although the respiration
may be slightly higher (Imshenetsky and Perova, 1957). Claus (1956) has
shown that the respiration of Aerobacter aerogenes is initially depressed by
Hg++ but recovers after several hours and reaches normal levels (Fig. 7-46).
Fig. 7-46. Effects of Hg++ on the respiration of Aerobacter
aerogenes. The O2 uptake is given as mm^ Oj/ml/SO min. A:
Initial exposure to Hg++; B: inoculation of organisms from
3 in A and re-exposure. C, control; 1, 0.0011 mM;
2, 0.0022 mM; 3, 0.0044 mM; 4, 0.0088 mM; 5, 0.0178
mM; 6, 0.037 mM. (From Claus, 1956.)
Adaptation to mercurials is apparently specific in most cases, since Se-
verens and Tanner (1945) found that Salmonella sp. tolerant to Hg++ are
not tolerant to Cu++, and vice versa, while Launoy and Levaditi (1913)
showed that spirochetes tolerant to antisyphilitic mercurials are not
tolerant to arsenicals. However, Blumenthal and Pan (1963) noted that
penicillin-resistant strains of staphylococci are more apt to be resistant to
DEVELOPMENT OF EESISTANCE TO MERCURIALS 985
Hg++ than the normal strains. Resistance to the mercurials is usually not
lost during subsequent culturing in mercurial-free media, as shown for fungi
(Partridge and Rich, 1962), yeast (Zambonelli, 1958 a), and spirochetes
(Launoy and Levaditi, 1913). The most striking case of the retention of
resistance is that of Salmonella, tolerance being unchanged during 55 trans-
fers over a period of 18 months (Severens and Tanner, 1945). It has thus
been concluded that the tolerance is inherited. Morphological changes dur-
ing adaptation generally do not occur, but yeast cells tolerant to Hg++
are smaller, have lost their smooth contours, contain more vacuoles, and are
deficient in lipid (Imshenetsky and Perova, 1957). Growth of tolerant or-
ganisms in normal media is usually not different from that of sensitive
strains, and no instance of dependence on mercurials has been reported.
Development of resistance to mercurials is not confined to microorgan-
isms but is observed in certain mammalian tissues. Gil y Gil (1924) claimed
to have produced renal tolerance to Hg++ in rabbits, but his work was
criticized by Hunter (1929), who repeated and extended this study using
more approijriate dosages. Rabbits given nephrotoxic doses of mercurial
exhibited a regeneration of the renal epithelium around the fourth day,
the new cells being elongated, flattened, with hyperchromatic nuclei, and
somewhat resistant to Hg++. Tsurumaki et al. (1928) confirmed that the
kidneys of rabbits surviving toxic injections of HgCl, are not damaged by
the same doses if given 10-14 days after the disappearance of the original
nephritis. During the period of adaptation, even though repeated subcutane-
ous injections are given, the nephritis disappears, but there is no alteration
in the excretion of Hg++ (Miura, 1934). MacNider (1941) repeated such ex-
periments in dogs and claimed that the newly regenerated renal epithelial
cells are different both morphologically and chemically from the normal
cells. The depressant effects of meralluride on intestinal transport of Na+,
CI", and water also disappear with repeated administration of the drug
(Blickenstaff, 1954), as does the suppression of conditioned reflexes in rats
given HgCL (Galoyan, 1957).
REFERENCES
Abadom, P.N. and Scholefield, P.G. (1962). Can. J. Biochem. Physiol. 40, 1591.
Abe, S. (1963). Biol. Bull. 124, 107.
Abood, L.G. and Romanchek, L. (1955). Biochem. J. 60, 233.
Abraham, E.P. and Newton, G.G.F. (1956). Biochem. J. 63, 628.
Abraham, S., Matthes, K.J., and Chaikoff, I.L. (1960). Biochem. Biophys. Res. Com-
mun. 3, 646.
Abraham, S., Matthes, K.J. and Chaikoff, I.L. (1961). Biochim. Biophys. Acta 49, 268.
Abul-Fadl, M.A.M. and King, E.J. (1949). Biochem. J. 45, 51.
Ackermann, W.W. (1951). J. Biol. Chem. 189, 421.
Ackermann, W.W. and Potter, V.R. (1949). Proc. Soc. Exptl. Biol. Med. 11, 1.
Ackermann, W.W. and Taylor, A. (1948). Proc. Soc. Exptl. Biol. Med. 67, 449.
Acland, J.D. (1962). Biochem. J. 84, 77p.
Ada, G.L. (1963). Biochim. Biophys. Acta 73, 276.
Adams, E. (1954). J. Biol. Chem. 209, 829.
Adams, E. (1955). J. Biol. Chem. 217, 325.
Adams, E. (1959). J. Biol. Chem. 234, 2073.
Adams, E. and Goldstone, A. (1960 a). J. Biol. Chem. 235, 3499.
Adams, E. and Goldstone, A. (1960 b). J. Biol. Chem. 235, 3504.
Adell, B. (1940). Z. Physik. Chem. A 185, 161.
Adkins, H., Cramer, H.I., and Connor, R. (1931). J. Am. Chem. Soc. 53, 1402.
Adler, E., von Euler, H. Giinther, G., and Plass, M. (1939). Biochem. J. 33, 1028.
Agarwala, S.C, Krishna Murti, C.R., and Shrivastava, D.L. (1954). Enzymologia 16,
322.
Agosin, M. and Aravena, L. (1959). Biochim. Biophys. Acta 34, 90.
Agosin, M. and Aravena, L. (1960). Enzymologia 22, 281.
Agosin, M. and von Brand, T. (1953). J. Infect. Diseases 93, 101.
Agosin, M. and von Brand, T. (1955). Exptl. Parasitol. 4, 548.
Agosin, M. and Weinbach, E.C. (1956). Biochim. Biophys. Acta 21, 117.
Agosin, M., von Brand, T., Rivera, G.Fr, and McMahon, P. (1957). Exptl. Parasitol.
6, 37.
Agranoff, B.W., Eggerer, H., Henning, U., and Lynen, F. (1960). J. Biol. Chem. 235, 326.
Ahmed, K. and Scholefield, P.G. (1962). Can. J. Biochem. Physiol. 40, 1101.
Aikawa, J.K., Blumberg, A.J., and Catterson, D.A. (1955). Proc. Soc. Exp. Biol.
Med. 89, 204.
Airan, J.W. and Barnabas, J. (1953). Naturwissenschaften 40, 510.
Airan, J.W., Joshi, J.V., Barnabas, J., and Master, R.W.P. (1953). Anal. Chem. 25, 659.
Aisenberg, A.C., Reinafarje, B., and Potter, V.R. (1957). J. Biol. Chem. 224, 1099.
Ajl, S.J. and Werkman, C.H. (1948). Proc. Natl. Acad. Sci. U.S. 34, 491.
Akamatsu, S., Kiyomoto, A., Harigaya, S., and Ohshima, S. (1961). Nature 191, 1298.
Akiba, T. and Ishii, K. (1952). Japan. J. Exptl. Med. 22, 241.
Alanis, J. (1948). Arch. Inst. Cardiol. Mex. 18, 410.
Albaum, H.G. and Eichel, B. (1943). Am. J. Botany 30, 18.
Albers, R.W. and Koval, G.J. (1961). Biochim. Biophys. Ada 52, 29.
987
988 REFERENCES
Albert, A. (1960). "Selective Toxicity," 2nd ed. Wiley, New York.
Albert, A., Falk, J.E., and Rubbo, S.D. (1944). Nature 153, 712.
Albrecht, A.M. (1957). Federation Proc. 16, 144.
Alburn, H.E. and Whitley, R.W. (1954). Federation Proc. 13, 330.
Aleem, M.I.H. and Lees, H. (1963). Can. J. Biochetn. Physiol. 41, 763.
Alivisatos, S.G.A. and Denstedt, O.F. (1952). J. Biol. Chem. 199, 493.
Alivisatos, S.G.A. , Kashket, S., and Denstedt, O.F. (1956). Can. J. Biochem. Physiol.
34, 46.
Allison, A.C. (1962). Virology 17, 176.
Allison, A.C. and Cecil, R. (1958). Biochem. J. 69, 27.
Allison, A.C, Buckland, F.E., and Andrews, C.H. (1962). Virology 17, 171.
Almkvist, J. (1922). Arch. Dermatol. Syphilol. 141, 342.
Altekar, W.W. and Rao, M.R.R. (1963). J. Bacteriol. 85, 604.
Altszuler, N., Dunn, A., Steele, R., Bishop, J.S., and De Bodo, R.C. (1963). Am. J.
Physiol. 204, 1008.
Amaha, M. and Nakahara, T. (1959). Nature 184, 1255.
Ames, S.R. and Elvehjem, C.A. (1944 a). Proc. Soc. Exptl. Biol. Med. 57, 108.
Ames, S.R. and Elvehjem, C.A. (1944 b). Arch. Biochem. 5, 191.
Ames, S.R. and Elvehjem, C.A. (1945). J. Biol. Chem. 159, 549.
Anagnostopoulos, C. (1953 a). Bull. Soc. Chim. Biol. 35, 575.
Anagnostopoulos, C. (1953 b). Bull. Soc. Chim. Biol. 35, 595.
Anderson, B.M. and Kaplan, N.O. (1959). J. Biol. Chem. 234, 1226.
Anderson, D.G., Rice, M.S., and Porter, J.W. (1960). Biochem. Biophys. Res. Com-
mun. 3, 591.
Anderson, E.P. and Heppel, L.A. (1960). Biochim. Biophys. Acta 43, 79.
Andrews, G.S. and Taylor, K.B. (1955). Brit. J. Exptl. Pathol. 36, 611.
Angielski, S., Rogulski, J., and Madonska, L. (1960a). Acta Biochim. Polon. 7, 269.
Angielski, S., Rogulski, J., Mikulski, P., and Popinigis, J. (1960b). Acta Biochim.
Polon. 7, 285.
Angielski, S., Rogulski, J., and Basciak, J. (1960 c). Acta Biochim. Polon. 7, 295.
Anichkov, S.V. (1953). DoM. na Mezhdunar. Fiziol. Kongr., Montreal 19, 7.
Annau, E. (1935). Z. Physiol. Chem. 236, 58.
Anson, M.L. (1939 a). J. Gen. Physiol. 23, 239.
Anson, M.L. (1939 b). J. Gen. Physiol. 23, 247.
Anson, M.L. (1940). J. Gen. Physiol. 23, 321.
Anson, M.L. (1941). J. Gen. Physiol. 24, 399.
Anson, M.L. and Mirsky, A.E. (1931). J. Gen. Physiol. 14, 605.
Anson, M.L. and Stanley, W.M. (1941). J. Gen. Physiol. 24, 679.
Anthony, D.D., Starr, J.L., Kerr, D.S., and Goldthwait, D.A. (1963). J. Biol Chem.
238, 690.
Aposhian, H.V. and Lambooy, J. P. (1955). J. Am. Chem. Soc. 77, 6368.
Aposhian, H.V. and Pointer, N.S. (1958). Federation Proc. 17, 344.
App, A. A. and Meiss, A.N. (1958). Arch. Biochem. Biophys. 77, 181.
Applewhite, T.H., Martin, R.B., and Niemann, C. (1958). J. Am.. Chem. Soc. 80, 1457.
Appleyard, G. and Woods, D.D. (1956). J. Gen. Microbiol. 14, 351.
Araki, K. and Myers, D.K. (1963). Can. J. Biochem. Physiol. 41, 2157.
Arbuthnott, J.P. (1962). Nature 196, 277.
Archibald, R.M. (1944). J. Biol. Chem. 154, 657.
REFERENCES 989
Arkin, A. (1911). J. Pharmacol. Exptl. Therap. 3, 145.
Arkin, A. (1912). J. Infect. Diseases 11, 427.
Armstrong, J. McD., Coates, J.H., and Morton. R.K. (1960). Xafure 186. 1033.
Armstrong, J. McD., Coates, J.H., and Morton, R.K. (1963). Biochem. J. 88, 266.
Arndt, F. (1955). Org. Syn. 20, 26.
Amon, D.I., Allen, M.B., and Whatley, F.R. (1956). Biochim. Biophys. Acta 20, 449.
Asahi, T., Bandurski, R.S., and Wilson, L.G. (1961). J. Biol. Chem. 236, 1830.
Asano, A. and Brodie, A.F. (1963). Biochem. Biophys. Res. Commun. 13, 423.
Asnis, R.E., Vely, V.G., and Glick, M.C. (1956). J. Bacteriol. 11, 314.
Astrup, T. and Thorsell, W. (1954). Acta Chem. Scand. 8, 1859.
Atkinson, D.E. (1956). J. Biol. Chem. 218, 557.
Atkinson, M.R., Jackson, J.F., and Morton, R.K. (1961). Nature 192, 946.
Atkinson, M.R., Morton, R.K., and Murray, A.W. (1963). Biochem. J. 89, 167.
Aubel, E. and Szulmajster, J. (1950). Biochim. Biophys. Acta 5, 255.
Aiibel, E., Rosenberg. A.J., and Szulmajster, J. (1950). Biochim. Biophys. Acta 5, 228.
Auditore, G.V. and Holland, W.C. (1956). Am. J. Physiol. 187, 57.
Augustinsson, K.-B. and Heimbiirger, G. (1955). Acta Chem. Scand. 9, 383.
Austen, K.F. and Brocklehurst, W.E. (1960). Nature 186, 866.
Avi-Dor, Y. and Gonda, O. (1959). Biochem. J. 72, 8.
Avi-Dor, Y.. KuczjTiski, M., Schatzberg, G-, and Mager, J. (1956). J. Gen. Microbiol.
14, 76.
Avi-Dor, Y., Traub, A., and Mager, J. (1958). Biochim. Biophys. Acta 30, 164.
Avron, M. and Biale. J.B. (1957). Plant Physiol. 32, 100.
Avron, M. and Jagendorf, A.T. (1956). Arch. Biochem. Biophys. 65, 475.
Axelrod, A.E., Purvis, S.E., and Hofmann, K. (1948). J. Biol. Chem. 176, 695.
Axelrod, B. and Jang, R. (1954). J. Biol. Chem. 209, 847.
Axelrod, B., Saltman, P., Bandurski, R.S., and Baker, R.S. (1952). J. Biol. Chem.
197, 89.
Axelrod, J. and Cochin, J. (1956). Federation Proc. 15, 395.
Axelrod, J. and Cochin, J. (1957). J. Pharmacol. Exptl. Therap. 121, 107.
Axelrod, J. and Laroche, M.-J. (1959). Science 130, 800.
Ayengar, P. and Roberts, E. (1952). Proc. Soc. Exptl. Biol. Med. 79, 476.
Azoulay, E. and Heydeman, M.T. (1963). Biochim. Biophys. Acta 73, 1.
Azzone, G.F. and Carafoli, E. (1960). Exptl. Cell Res. 21, 447.
Azzone, G.F. and Ernster, L. (1961). J. Biol. Chem. lib, 1518.
Azzone, G.F., Carafoli, E., and Muscatello, U. (1960). Exptl. Cell Res. 21, 456.
Bach, M.K. (1961). Plant Physiol. 36, 317.
Bach, S.J. and Williamson, S. (1942). Nature 150, 575.
Bachmann, H. (1938). Arch. Exptl. Pathol. Pharmakol. 190, 345.
Bacq, Z.M. (1936). Arch. Intern. Physiol. 42, 340.
Bacq, Z.M. (1942). Bull. Acad. Roy. Med. Belg. 7, 108.
Bacq, Z.M., Gosselin, L., Dresse, A., and Renson, J. (1959). Science 130, 453.
Baer, H. (1948). J. Biol. Chem. 173, 211.
Baernstein, H.D. (1936). J. Biol. Chem. 115, 25.
Bahn, R.C. and Longley, J.B. (1956). J. Pharmacol. Exptl. Therap. 118, 365.
Bailey, K. and Marsh, B.B. (1952). Biochim. Biophys. Acta 9, 133.
Bailey, K. and Perry, S.V. (1947). Biochim. Biophys. Acta 1, 506.
990 REFERENCES
Bain, J.A. and Williams, H.L. (1960). In "Inhibition in the Nervous System and y-
Aminobutyric Acid" (E. Roberts, ed.), p. 275. Macmillan (Pergamon), New York
Baker, N. and Wilson, L. (1963). Biochem. Biophys. Res. Commun. 11, 60.
Baker, Z., Fazekas, J.F., and Himwich, H.E. (1938). J. Biol. Chem. 125, 545.
Baldwin, E. (1938). Biochem. J. 32, 1225.
Balinsky, D. and Davies, D.D. (1961a). Biochem. J. 80, 292.
Balinsky, D. and Davies, D.D. (1961b). Biochem. J. 80, 296.
Balis, M.E. and Dancis, J. (19.55). Cancer Res. 15, 603.
Ball, H.A. and Sanders, C.R. (1958). Federation Proc. 17, 427.
Balls, A.K. and Lineweaver, H. (1939 a). Nature 144, 513.
Balls, A.K. and Lineweaver, H. (1939 b). J. Biol. Chem. 130, 669.
Baltscheffsky, H. (1960 a). Biochim,. Biophys. Acta 40, 1.
Baltscheffsky, H. (1960 b). Acta Chem. Scand. 14, 264.
Baltscheffsky, H. and Baltscheffsky, M. (1958). Acta Chem. Scand. 12, 1333.
Bandelin, F.J. (1958). J. Am. Pharm. Assoc, Sci. Ed. 47, 691.
Bandurski, R.S. (1955). J. Biol. Chem. 217, 137.
Banga, I., Ochoa, S., and Peters, R.A. (1939). Biochem. J. 33, 1109.
Baptist, J.N. and Ve.stling, C.S. (1957). Federation Proc. 16, 150.
Barany, M. (1959). In "Sulfur in Proteins" (R. Benesch et al., eds.), p. 317. Academic
Press, New York.
Barany, M. and Barany, K. (1959 a). Biochim. Biophys. Acta 35, 293.
Barany, M. and Barany, K. (1959 b). Biochim. Biophys. Acta 35, 544.
Barany, M., Finkelman, F., and Therattil-Antony, T. (1962). Arch. Biochem. Biophys.
98, 28.
Barban, S. (1961). Biochim. Biophys. Acta 47, 604.
Barban, S. (1962 a). Biochim. Biophys. Acta 65, 376.
Barban, S. (1962 b). J. Biol. Chem. 237, 291.
Barban, S. and Schulze, H.O. (19.56). J. Biol. Chem. Ill, 665.
Barban, S. and Schulze, H.O. (1961). J. Biol. Chem. 236, 1887.
Barbate, L.M. and Abood, L.G. (1963). Biochim. Biophys. Acta 67, 531.
Barber, G.A. (1959). Federation Proc. 18, 185.
Barbour, H.G. and Hjort, A.M. (1921). J. Pharmacol. Exptl. Therap. 18, 201.
Bargoni, N. (1963). Z. Physiol. Chem. 333, 242.
Barker, S.B. (1953). Federation Proc. 12, 9.
Barman, T.E., Parke, D.V., and Williams, R.T. (1961). Biochem. Pharmacol. 8, 46.
Barman, T.E., Parke, D.V., and Williams, R.T. (1963). Toxicol. Appl. Pharmacol. 5,
545.
Barnabas, J. (1955). Anal. Chem. 27, 443.
Barnard, E.A. and Ramel, A. (1962). Biochem. J. 84, 72p.
Barnard, E.A. and Stein, W.D. (1958). Advan. Enzymol. 20, 51.
Barnes, H. and Stanbury, F.A. (1948). J. Exptl. Biol. 25, 270.
Barnes, J.M., Magee, P.N., Boyland, E., Haddow, A., Passey, R.D., Bullough, W.S.,
Cruickshank, C.N.D., Salaman, M.H., and Williams, R.T. (1957). Nature 180, 62.
Barnett, R.C. (1953). Biol. Bull. 104, 263.
Barnett, R.C. and Downey, M. (1955). Federation Proc. 14, 9.
Barone, J.A., Tieckelmann, H., Guthrie, R., and Holland, J.F. (1960). J. Org. Chem.
25, 211.
Barrett, F.R. (1957). Med. J. Australia p. 242.
REFERENCES 991
Barron, E.S.G. (1951). Advan. Enzymol. 11, 201.
Barron, E.S.G. and Dickman, S. (1949). J. Gen. Phijsiol 32, 595.
Barron, E.S.G. and Ghiretti, F. (1953). Biochim. Biophys. Acta 12, 239.
Barron, E.S.G. and Goldinger, J.M. (1941). Proc. Soc. Exptl. Biol. Med. 48, 570.
Barron, E.S.G. and Kalnitsky, G. (1947). Biochem. J. 41, 346.
Barron, E.S.G. and Levine, S. (1952). Arch. Biochem. Biophys. 41, 175.
Barron, E.S.G. and Singer, T.P. (1945). J. Biol. Chem. 157, 221.
Barron, E.S.G. and Tahmisian, T.N. (1948). J. Cellular Comp. Physiol. 32, 57.
Barron, E.S.G., Dick, G.F., and Lyman, CM. (1941). J. Biol. Chem. 137, 267.
Barron, E.S.G., Nelson, L., and Ardao, M.I. (1948). J. Gen. Physiol. 32, 179.
Barron, E.S.G., Ardao, M.I., and Hearon, M. (1950). Arch. Biochem. Biophys. 29, 130.
Barron, E.S.G., Sights, W.P., and Wilder, V. (1953). Arch. Exptl. Pathol. Pharmakol.
219, 338.
Barry, R.J.C., Dikstein, S., Matthews, J., Smyth, D.H., and Wright, E.M. (1964).
J. Physiol. [London) 171, 316.
Bartlett, G.R. (1948). J. Am. Chem. Soc. 70, 1010.
Bartley, W. and Davies, R.E. (1954). Biochem. J. 57, 37.
Bassham, J.A., Benson, A.A., and Calvin, M. (1950). J. Biol. Chem. 185, 781.
Bastarrachea, F., Anderson, D.G., and Goldman, D.S. (1961). J. Bacteriol. 82, 94.
Battaglia, F.C. and Randle, P.J. (1960). Biochem. J. 75, 408.
Bauchop, T. and Dawes, E.A. (1959). Biochim. Biophys. Acta 36, 294.
Bauer, C.W. and La Sala, E.F. (1956). J. Am. Pharm. Assoc, Sci. Ed. 45, 675.
Baugh, C.L., Claus, G.W., and Werkman, C.H. (1960). Arch. Biochem. Biophys. 86,
255.
Baumann, C.A. and Stare, F.J. (1940). J. Biol. Chem. 133, 183.
Baxter, C.F. and Roberts, E. (1958). J. Biol. Chem. 233, 1135.
Baxter, R.M. and Quastel, J.H. (1953). J. Biol. Chem. 201, 751.
Bayne, S. (1952). Biochem. J. 50, xxvii.
Bean, R.C. and Hassid, W.Z. (1956). Science 124, 171.
Beaton, G.H. and McHenry, E.W. (1953). Brit. J. Nutr. 7, 357.
Beaton, J.R. and Goodwin, M.E. (1955). J. Biol. Chem. 212, 195.
Beaton, J.R., Goodwin, M.E., Ozawa, G., and McHenry, E.W. (1954). Arch. Biochem.
Biophys. 51, 94.
Beck, G.E. and Bein, H.J. (1948). Helv. Physiol. Pharmacol. Acta 6, 398.
Becker, B. (1961). Am. J. Physiol. 200, 804.
Becker, B. and Friedenwald, J.S. (1949). Arch. Biochem. 22, 101.
Becker, C.E. (1954). Federation Proc. 13, 180.
Becker, C.E. and Day, H.G. (1953). J. Biol. Chem. 201, 795.
Becker, C.E. and May, C.E. (1949). J. Am. Chem. Soc. 71, 1491.
Becker, V. and Rauschke, J. (1951). Z. Ges. Exptl. Med. 117, 374.
Becker, V. and Rieken, E. (1954). Arch. Pathol. Anat. Physiol. 325, 1.
Beckmann, F. (1934). Arch. Ges. Physiol. 234, 469.
Beechey, R.B. (1961). Nature 192, 975.
Beers, R.F., Jr. (1961). J. Biol. Chem. 236, 2703.
Beevers, H. (1952). Plant Physiol. TJ, 725.
Beevers, H. (1954). Plant Physiol. 29, 265.
Beevers, H. (1956). Plant Physiol. 31, 440.
Beevers, H. and French, R.C. (1954). Arch. Biochem. Biophys. 50, 427.
992 REFERENCES
Beevers, H., Goldschmidt, E.P., and Koffler, H. (1952). Arch. Biochem.. Biophys. 39,
236.
Behal, F.J. (1959). Federation Proc. 18, 188.
Behal, F.J. and Hamilton, R.D. (1962). Arch. Biochem. Biophys. 96, 530.
Beher, W.T. and Anthony, W.L. (1953). J. Biol. Chem. 203, 895.
Beher, W.T., Holliday, W.M., and Gaebler, O.H. (1952). J. Biol. Chem. 198, 573.
Beher, W.T., Baker, G.D., and Madoff, M. (1959). J. Biol. Chem. 234, 2388.
Beinert, H. (1960). In "The Enzymes" (P.D. Boyer, H. Lardy, and K. Myrback, eds.),
Vol. 2, p. 339. Academic Press, New York.
Beinert, H. and Stansly, P.G. (19.53). J. Biol. Chem. 204, 67.
Beling, C.G. and Diczfalusy, E. (1959). Biochem. J. 71, 229.
Belkin, M. and Hardy, W.G. (1961). J. Biophys. Biochem. Cytol. 9, 733.
Bell, F.E. and Mounter, L.A. (1958). J. Biol. Chem. 233, 900.
Bellin, S.A. and Smeby, R.R. (1958). Arch. Biochem. Biophys. 75, 1.
Bello, L.J., van Bibber, M.J., and Bessman, M.J. (1961). Biochim. Biophys. Acta 53,
194.
Benesch, R. and Benesch, R.E. (1952). Arch. Biochem. Biophys. 38, 425.
Benesch, R. and Benesch, R.E. (1962). In "Methods of Biochemical Analysis" (D.
Glick ed.), Vol. 10, p. 43. Wiley (Interscience), New York.
Benesch, R., Benesch, R.E., and Rogers, W.I. (1954). In "Glutathione" (S.P. Colo-
wick, ed.), p. 31. Academic Press, New York.
Benesch, R.E. and Benesch, R. (1953). J. Am. Chem. 8oc. 75, 4367.
Benesch, R.E. and Benesch, R. (1954). Arch. Biochem. Biophys. 48, 38.
Benesch, R.E. and Benesch, R. (19.55). J. Am.. Chem.. Soc. 77, 5877.
Benesch, R.E. and Benesch, R. (1962). Biochemistry 1, 735.
Benigno, P. and Santi, R. (1946). Ric. Sci. 16, 1779.
Bennett, D.R., Andersen, K.S., Andersen, M.V., Jr., Robertson, D.N., and Cheno-
weth, M.B. (1958). J. Pharmacol. Exptl. Therap. '[11, 489.
Bennett, E.D. and Bauerle, R.H. (1960). Australian J. Biol. Sci. 13, 142.
Bennett, H.S. (1948 a). Anat. Record 100, 640.
Bennett, H.S. (1948 b). Anat. Record 100, 732.
Bennett, H.S. (1951). Anat. Record 110, 231.
Bentley, L.E. (1952). Nature 170, 847.
Bentley, R. and Keil, J.G. (1961). Proc. Chem. Soc. p. 111.
Bentley, R. and Thiessen, C.P. (1957). J. Biol. Chem.. lib, 689.
Beppu, T. (1961). J. Biochem. [Tokyo) SO, 386.
Ber, A. and Singer-Altbeker, R. (1961). Biochem. Pharmacol. 7, 169.
Bergel, F. and Todd, A.R. (1937). J. Chem. Soc. p. 1504.
Berger, J. and Avery, G.S., Jr. (1943 a). Am. J. Botany 30, 290.
Berger, J. and Avery, G.S., Jr. (1943 b). Am. J. Botany 30, 297.
Berger, M. and Harman, J.W. (1954). Federation Proc. 13, 183.
Bergersen, F.J. (1960). J. Gen. Microbiol. 11, 671.
Bergersen, F.J. (1962). J. Gen. Microbiol. 29, 113.
Bergkvist, R. (1963). Acta Chem. Scand. 17, 1541.
Bergmann, F., Segal, R., and Rimon, S. (1957). Biochem. J. 67, 481.
Bergmann, F., Levin, G., and Kwietny, H. (1959). Arch. Biochem. Biophys. 80, 318.
Bergmann, F., Kwietny-Govrin, H., Ungar-Waron, H., Kalmus, A., and Tamari, M.
(1963 a). Biochem. J. 86, 567.
REFERENCES 993
Bergmann, F., Ungar-Waron, H., and Kwietny-Govrin, H. (1963 b). Biochem. J. 86,
292.
Bergmeyer, H.-U., Holz, G., Kauder, E.M., Mollering, H., and Wieland, O. (1961).
Biochem. Z. 333, 471.
Bergquist, P.L. (1958). Physiol. Plantarum 11, 760.
Berk, S., Ebert, H., and Teitell, L. (1957). Ind. Eng. Chem. 49, 1115.
Berlin, M. and Gibson, S. (1963). Arch. Environ. Health 6, 617.
Berlin, M. and Ullberg, S. (1963). Arch. Environ. Health 6, 589.
Berman, D.A. (1951). Ph.D. Thesis, University of Southern CaHfornia.
Berman, D.A. and Saunders, P.R. (1955). Circulation Res. 3, 559.
Bernfeld, P. (1963). In "Metabolic Inhibitors" (R.M. Hochster and J.H. Quastal, eds.),
Voh 2, p. 437. Academic Press, New York.
Bernfeld, P. and Kelley, T.F. (1963). J. Biol. Chem. 238, 1236.
Bernfekl, P., Tuttle, L.P., and Hubbard, Pv.W. (1961). Arch. Biochern. Biophys. 92,
232.
Bernhard, S.A. and Gutfreund, H. (1956). Biochem. J. 63, 61.
Bernheim, A. I., Hirschhorn, L., and Muhnos, M.G. (1932). J. Pharmacol. Exptl. Therap.
44, 81.
Bernheim, F. (1928). Biochem. J. 22, 1178.
Bernheim, F. (1963). Proc. Soc. Exptl. Biol. Med. 113, 411.
Bernheim, F. and Dixon, M. (1928). Biochem. J. 22, 113.
Bernheim, F., DeTurk, W.E., and Pope, H. (1953). J. Bacteriol. 65, 544.
Bernstein, D.E. (1944). Arch. Biochem. 3, 445.
Bernstein, G.S. and Black, R.E. (1959). Proc. Soc. Exptl. Biol. Med. 102, 531.
Berry, L.J. (1955). Ann. N. Y. Acad. Sci. 62, 327.
Berry, L.J. and Beuzeville, C. (1960). J. Infect. Diseases 106, 183.
Berry, L.J. and Derbyshire, J.E. (1956). Proc. Soc. Exptl. Biol. Med. 92, 315.
Berry, L.J. and Mitchell, R.B. (1953 a). Science 118, 140.
Berry, L.J. and Mitchell, R.B. (1953 b). J. Infect. Diseases 93, 75.
Berry, L.J. and Mitchell, R.B. (1953 c). J. Infect. Diseases 93, 83.
Berry, L.J. and Mitchell, R.B. (1954). Proc. Soc. Exptl. Biol. Med. 85, 209.
Berry, L.J., Merritt, P., and Mitchell, R.B. (1954 a). J. Infect. Diseases 94, 144.
Berry, L.J., Ehlers, K.H., and Mitchell, R.B. (1954b). J. Infect. Diseases 94, 152.
Bersin, T. and Logemann, W. (1933). Z. Physiol. Chem. 220, 209.
Bertino, J.R., Gabrio, B.W., and Huennekens, F.M. (1960). Biochem. Biophys. Res.
Commun. 3, 461.
Beyer, K.H. and Baer, J.E. (1960). Progr. Drug Res. 2, 9.
Beyer, K.H., Wright, L.D., Skeggs, H.R., Russo, H.F., and Shaner, G.A. (1947). Am.
J. Physiol. 1 51 , 202.
Beyer, R.E. (1960). Biochim. Biophys. Acta 41, 552.
Beyer, R.E., Lamberg, S.L., and Neyman, M.A. (1961). Can. J. Biochem. Physiol. 39,
73.
Bhuvaneswaran, C, Sreenivasan, A., and Rege, D.V. (1961). Enzymologia 23, 185.
Bhuyan, B.K. and Simpson, F.J. (1962). Can. J. Microbiol. 8, 737.
Bickers, J.N., Bresler, E.H., and Weinberger, R. (1960). J. Pharmacol. Exptl. Therap.
128, 283.
Bide, R.W. and Collier, H.B. (1964). Can. J. Biochem. Physiol. 42, 669.
Bieleski, R.L. (1960). Australian J. Biol. Sci. 13, 221.
994 REFEKENCES
Bilinski, E. and Jonas, R.E.E. (1964). Can. J. Biochem. Physiol. 42, 345.
Bilodeau, F. and Elliott, K.A.C. (1963). Can. J. Biochem. Physiol. 41, 779.
Birkenhager, J.C. (1959). Biochim. Biophys. Acta 31, 595.
Birkenhager, J.C. (1960). Biochim. Biophys. Acta 40, 182.
Bivings, L. and Lewis, G. (1948). J. Pediat. 32, 63.
Bjerrum, J. (1941). "Metal Ammine Formation in Aqueous Solution." P. Haase,
Copenhagen.
Bjerrum, J., Schwarzenbach, G., and Sillen, L.G. (1957). "Stability Constants," Part
1. Chemical Soc, London.
Black, M.M. and Kleiner, I.S. (1947). Cancer Res. 7, 717.
Black, M.M., Kleiner, I.S., and Bolker, H. (1947). Cancer Res. 7, 818.
Black, S. (1951). Arch. Biochem. Biophys. 34, 86.
Blair, D.G.R. and Potter, V.R. (1961). J. Biol. Chem. 236, 2503.
Blakley, E.R. and Boyer, P.D. (1955). Biochim. Biophys. Acta 16, 576.
Blakley, E.R. and Ciferri, 0. (1961). Can. J. Microbiol. 7, 61.
Blakley, R.L. (1952). Biochem. J. 52, 269.
Blakley, R.L. (1954). Biochem. J. 58, 448.
Blakley, R.L. (1957). Biochem. J. 65, 342.
Blakley, R.L. and McDougall, B.M. (1961). J. Biol. Chem. 236, 1163.
Blanchard, M., Green, D.E., Nocito, V., and Ratner, S. (1944). J. Biol. Chem. 155,
421.
Blaschko, H. (1935). Biochem. J. 29, 2303.
Blaschko, H. (1942). J. Physiol. (London) 101, 337.
Blaschko, H. and Duthie, R. (1945). Biochem. J. 39, 347.
Blaschko, H. and Hinims, J.M. (1955). Brit. J. Pharmacol. 10, 451.
Blaschko, H., Fastier, F.N., and Wajda, I. (1951). Biochem. J. 49, 250.
Blaylock, B.A. and Nason, A. (1963). J. Biol. Chem. 238, 3453.
Bhckenstaff, D.D. (19.54). Am. J. Physiol. 178, 371.
Blum, J.J. (1955). Arch. Biochem. Biophys. 55, 486.
Blum, J.J. (1960). Arch. Biochem. Biophys. 87, 104.
Blum, J.J. (1962 a). Arch. Biochem. Biophys. 97, 309.
Blum, J.J. (1962 b). Arch. Biochem. Biophys. 97, 321.
Blumenstein, J. and Williams, G.R. (1963). Can. J. Biochem. Physiol. 41, 201.
Blumenthal, H.J. and Pan, Y.-L. (1963). Proc. Soc. Exptl. Biol. Med. 113, 322.
Blumson, N.L. (1957). Biochem. J. 65, 138.
Bock, K.R., Robinson, J.B.D., and Chamberlain, G.T. (1958). Nature 182, 1607.
Bodansky, O. (1961). J. Biol. Chem. 236, 328.
Boeri, E. and Tosi, L. (1954). Arch. Biochem. Biophys. 52, 83.
Boeri, E., Cutolo, E., Luzzati, M., and Tosi, L. (1955). Arch. Biochem. Biophys. 56,
487.
Boeri, E., Cremona, T., and Singer, T.P. (1960). Biochem. Biophys. Res. Commun. 2,
298.
Bone, D.H. and Hochster, R.M. (1960). Can. J. Biochem. Physiol. 38, 193.
Boney, A.D., Corner, E.D.S., and Sparrow, B.W.P. (1959). Biochem. Pharmacol. 2, 37.
Bonner, B.A. and Machlis, L. (1957). Plant Physiol. 32, 291.
Bonner, J. (1948). Arch. Biochem. 17, 311.
Bonner, J. (1949). Am. J. Botany 36, 429.
Bonner, J. and Millerd, A. (1953). Arch. Biochem. Biophys. 42, 135.
REFEKENCES 995
Bonner, J. and Wildman, S.G. (1946). Arch. Biochem. 10, 497
Bonner, J., Bandurski, R.S., and Millerd, A. (1953). Physiol. Plantarum 6, 511.
Bonner, W.D., Jr. (1954). Biochem. J. 56, 274.
Bonsignore, A., Pontremcli, S., Grazi, E., and Horecker, B.L. (1960). J. Biol. Chem.
235, 1888.
Booth, A.N., Taylor, J., Wilson, R.H., and DeEds, F. (1952). J. Biol. Chem. 195, 697.
Borghgraef, R.R.M. and Pitts, R.F. (1956). J. Clin. Invest. 35, 31.
Borghgraef, R.R.M., Kessler, R.H., and Pitts, R.F. (1956). J. Clin. Invest. 35, 1055.
Borsook, H., Fischer, E.H., and Keighley, G. (1957). J. Biol. Chem. 229, 1059.
Borst, P. (1962). Biochim. Biophys. Acta 57, 256.
Bortz, W.M. and Lynen, F. (1963). Biochem. Z. 337, 305.
Boser, H. (1959). Z. Physiol. Chem. 315, 163.
Bosund, I. (1959). Acta Chem. Scand. 13, 803.
Bosund. I. (1960 a). Acta Chem. Scand. 14, 1231.
Bosund, I. (1960 b). Acta Chem. Scand. 14, 1243.
Boucek, R.J. and Paff, G.H. (1961). Am. J. Physiol. 201, 668.
Bovarnick, M.R., Lindsay, A., and Hellerman, L. (1946). J. Biol. Chem. 163, 523.
Bowman, I.B.R., Grant, P.T., Kermack, W.O., and Ogston, D. (1961). Biochem. J.
78, 472.
Bowman, I.B.R., Tobie, E.J., and von Brand, T. (1963). Comp. Biochem.. Physiol. 9, 105.
Boxer, G.E., Shonk, C.E., Stoerk, H.C., Bielinski, T.C., and Budzilovich, T.V. (1955).
J. Biol. Chem. 217, 541.
Boyd, E.S. and Neuman, W.F. (1954). Arch. Biochem. Biophys. 51, 475.
Boyer, P.D. (1954). J. Am. Chem. Soc. 76, 4331.
Boyer, P.D. (1959). In "The Enzymes" (P.D. Boyer, H. Lardy, and K. Myrback, eds.).
Vol. 1, p. 511. Academic Press, New York.
Boyer, P.D. and Segal, H.L. (1954). In "The Mechanism of Enzyme Action" (W.D.
McElroy and H.B. Glass, eds.), p. 520. Johns Hopkins Press, Baltimore, Maryland.
Boyland, E. (1940). Biochem. J. 34, 1196.
Boyland, E. and Boyland, M.E. (1936). Biochem. J. 30, 224.
Boyland, E. and Levi, A.A. (1936). Biochem. J. 30, 2007.
Boyland, E., Wallace, D.M., and Williams, D.C. (1957). Brit. J. Cancer 11, 578.
Boyland, E., Davies, J.H., and Williams, K. (1959). Biochem. J. 72, 28p.
Boylen, J.B. and Quastel, J.H. (1962). Nature 193, 376.
Bradshaw, W.H. and Barker, H.A. (1960). J. Biol. Chem. 235, 3620.
Brady, C.J. (1961). Biochem. J. 78, 631.
Brady, R.O. (1963). Biochim. Biophys. Acta 70, 467.
Brady, R.O. and Gurin, S. (1952). J. Biol. Chem. 199, 421.
Brady, R.O., Spyropoulos, C.S., and Tasaki, I. (1958). Am. J. Physiol. 194, 207.
Brand, E. and Kassell, B. (1940). J. Biol. Chem. 133, 437.
Branster, M.V. and Morton, R.K. (1956). Biochem. J. 63, 640.
Braun, K. (1949). J. Pharmacol. Exptl. Therap. 95, 58.
Braune, W. (1963). Arch. Mikrobiol. 45, 252.
Bramistein, A.E. (1960). In "The Enzymes" (P.D. Boyer, H. Lardy, and K. Myrback,
eds.), Vol. 2, p. 113. Academic Press, New York.
Braunstein, A.E., Azarkh, R.M., and Tin-Sen', S. (1962). Biochemistry (USSR) (Eng-
lish Transl.) 26, 760.
Bray, R.E. and VanArsdel, P.P. (1961). Proc. Soc. Exptl. Biol. Med. 106, 255.
996 REFERENCES
Bregoff, H.M. and Kamen, M.D. (1952). Arch. Biochem. Biophys. 36, 202.
Bresnick, E. (1962). Cancer Res. 22, 1246.
Bresnick, E. (1963). Biochim. Biophys. Acta 67, 425.
Bressler, R. and Wakil, S.J. (1961). J. Biol. Chem. 236, 1643.
Bressler, R. and Wakil, S.J. (1962). J. Biol. Chem. 237, 1441.
Breusch, F.L. (1942). Enzymologia 10, 165.
Breusch, F.L. and Keskin, H. (1944). Enzymologia 11, 243.
Brewer, C.R. and Werkman, C.H. (1939). Enzymologia 6, 273.
Briault, P.G. (1956). Nature 178, 1223.
Bridger, W.A. and Cohen, L.H. (1963). Biochim. Biophys. Acta 73, 514.
Briggs, F.N., Chernick, S., and Chaikoff, I.L. (1949). J. Biol. Chem. 179, 103.
Brink, N.G. (1953 a). Acta Chem. Scand. 7, 1081.
Brink, N.G. (1953 b). Acta Chem. Scand. 7, 1090.
Brock, N., Druckrey, H., and Herken, H. (1939). Arch. Exptl. Pathol. Pharmakol.
193, 679.
Brockman, R.W. and Anderson, E.P. (1963). In "Metabolic Inhibitors" (R.M. Hoch-
ster and J.H. Quastel, eds.), Vol. 1, p. 239. Academic Press, New York.
Brodersen,.R. and Kjaer, A. (1946). Acta Pharmacol. Toxicol. 2, 109.
Brodie, A.F. (1952). J. Biol. Chem. 199, 835.
Brody, T.M. (1956). J. Pharmacol. Exptl. Therap. 117, 39.
Brooks, J.L., Christensen, E., Stewart, C.J., and Wick, A.N. (1961). Proc. Soc. Exptl.
Biol. Med. 107, 395.
Brooks, P. and Davidson, N. (1960). J. Am. Chem. Soc. 82, 2118.
Brooks, S.A., Lawrence, J.C, and Ricketts, C.R. (1959). Biochem. J. 73, 566.
Brooks, S.A., Lawrence, J.C, and Ricketts, C.R. (1960). Nature 187, 1028.
Broquist, H.P., Kohler, A.R., Hutchinson, D.J., and Burchenal, J.H. (1953). J. Biol.
Chem. 202, 59.
Brown, D.D., Tomchick, R., and Axelrod, J. (1959). J. Biol. Chem. 234, 2948.
Brown, E.G. (1958). Biochem. J. 70, 313.
Brown, LA. (1954). A.M.A. Arch. Neurol. Psychiat. 72, 674.
Brown, J. and Bachrach, H.L. (1959). Proc. Soc. Exptl. Biol. Med. 100, 641.
Brown, W.D. and Tappel, A.L. (1959). Arch. Biocheyn. Biophys. 85, 149.
Bruchmann, E.-E. (1961a). Naturivissenschaften 48, 79.
Bruchmann, E.-E. (1961b). Naturwissenschaften 48, 305.
Bruchmann, E.-E. (1961c). Biochem. Z. 335, 199.
Bruemmer, J.H., Wilson, P.W., Glenn, J.L., and Crane, F.L. (1957). J. Bacterial.
73, 113.
Brun, C, Hilden, T., and Raaschou, F. (1947). Acta Pharmacol. Toxicol. 3, 1.
Brunnemann, A., Coper, H., and Herken, H. (1962). Arch. Exptl. Pathol. Pharmakol.
223, 223.
Brunngraber, E.G. (1958). J. Biol. Chem. 233, 472.
Bruns, F.H., Noltman, E., and Vahlhaus, E. (1958). Biochem. Z. 330, 483.
Brust, M. and Barnett, R.C. (1952). Federation Proc. 11, 19.
Bryce, G.F. and Rabin, B.R. (1964). Biochem. J. 90, 513.
Bublitz, C. and York, J.L. (1961). Biochim. Biophys. Acta 48, 56.
Buch, M.L., Montgomery, R., and Porter, W.L. (1952). Anal. Chem. 24, 489.
Buchman, E.R., Heegaard, E., and Bonner, J. (1940). Proc. Natl. Acad. Sci. U.S.
26, 561.
REFERENCES 997
Bueding, E. and MacKinnon, J.A. (1955). J. Biol. Chem. 215, 495.
Bulen, W.A. (1956). Arch. Biochem. Biophys. 62, 173.
Bulen, W.A. (1961). J. Baderiol. 82, 130.
Bullough, W.S. and Johnson, M. (1951). Proc. Boy. Soc. B138, 562.
BuUough, W.S. and Laurence, E.B. (1957). Brit. J. Exptl. Pathol. 38, 278.
Bu'Lock, J.D., Smalley, H.M., and Smith, G.N. (1962). J. Biol. Chem. 237, 1778.
Burch, G., Ray, T., Threefoot, S., Kelly, F.J., and Svedberg, A. (1950). J. Clin.
Invest. 29, 1131.
Burch, H.B., Lowry, O.H., Padilla, A.M., and Combs, A.M. (1956). J. Biol. Chem.
223, 29.
Burdette, W.J. (1952). Am. Heart J. 44, 823.
Burgess, E.A., Kerly, M., and Rahman, M.A. (1960). Biochem. J. 74, 4p.
Burnett, G.H. and Cohen, P.P. (1957). J. Biol. Chem. 229, 337.
Burney, T.E. and Golub, O.J. (1948). J. Immunol. 60, 213.
Burr, M.J. and Finamore, F.J. (1963). Biochim. Biophys. Acta 68, 608.
Burris, R.H. (1956). In "Inorganic Nitrogen MetaboHsm" (W.D. McElroy and B.
Glass, eds.), p. 316. Johns Hopkins Press, Baltimore, Maryland.
Burris, R.H. and Wilson, P.W. (1939). Cold Spring Harbor Symp. Quant. Biol. 7, 349.
Burton, K. (1951a). Biochem. J. 48, 458.
Burton, K. (1951b). Biochem. J. 50, 258.
Busch, H. and Nair, P.V. (1957). J. Biol. Chem. 229, 377.
Busch, H. and Potter, V.R. (1952 a). J. Biol. Chem. 198, 71.
Busch, H. and Potter, V.R. (1952 b). Cancer Res. 12, 660.
Busch, H. and Potter, V.R. (1953). Cancer Res. 13, 168.
Busch, H., Hurlbert, R.B., and Potter, V.R. (1952). J. Biol. Chem. 196, 717.
Butt, W.D. and Lees, H. (1960). Biochem. J. 76, 425.
Buzard, J.A. and Nytch, P.D. (1957). J. Biol. Chem. Til, 225.
Byers, S.O. (1952). J. Am. Pharm. Assoc, Sci. Ed. 41, 611.
Cady, P., Abraham, S., and Chaikoff, I.L. (1963). Bicchim. Biophys. Acta 70, 118.
Caffrey, R.W., Trembley, R., Gabrio, B.W., and Huennekens, F.M. (1956). J. Biol.
Chem. 112, 1.
Cafruny, E.J. (1962). Biochem:. Phmmacol. 9, 15.
Cafruny, E.J. and Farah, A. (1956). J. Pharmacol. Exptl. Therap. 117, 101.
Cafruny, E.J. and Palmer, J.F. (1961). J. Pharmacol. Exptl. Therap. 131, 250.
Cafruny, E.J., di Stefano, H.S., and Farah, A. (1955 a). J. Histochem. Cytochem. 3, 354.
Cafruny, E.J., Farah, A., and di Stefano, H.S. (1955b). J. Pharmacol. Exptl. Therap.
115, 390.
Cain, R.B. (1958). J. Gen. Microbiol. 19, 1.
Calcutt, G. (1950). Nature 166, 443.
Caldwell, E.F. and McHenry, E.W. (1953). Arch. Biochem. Biophys. 45, 466.
Calesnick, B., Wase, A., and Altarelli, V.R. (1960). Proc. Soc. Exptl. Biol. Med. 103,
22.
Callaghan, O.H. and Weber, G. (1959). Biochem. J. 73, 473.
Caltrider, P.G. and Gottlieb, D. (1963). Phytopathology 53, 1021.
Calvin, M. (1954). In "Glutathione" (S.P. Colowick et al., eds.), p. 3. Academic Press,
New York.
Calvin, M., Magel, T.T., and Hurd, CD. (1941). J. Am. Chem. Soc. 63, 2174.
998 REFERENCES
Campbell, D.E.S. (1959). Acta Pharmacol. Toxicol. 16, 151.
Campbell, P.N., Creasey, N.H., and Parr, C.W. (1952). Biochem. J. 52, 448.
Canellakis, Z.N. and Cohen, P.P. (1956 a). J. Biol. Chem. 222, 53.
Canellakis, Z.N. and Cohen, P.P. (1956 b). J. Biol. Chem. 222, 63.
Cannata, J.J.B. and Stoppani, A.O.M. (1963 a). Nature 200, 573.
Cannata, J.J.B. and Stoppani, A.O.M. (1963 b). J. Biol. Chem. 238, 1208.
Cannata, J.J.B. and Stoppani, A.O.M. (1963 c). J. Biol. Chem. 238, 1919.
Caraway, W.T. and Hellerman, L. (1953). J. Am. Chem. Soc. 75, 5334.
Carbon, J.A. (1962). Biochem. Biophys. Res. Commun. 7, 366.
Carney, S.A., Lawrence, J.C, and Ricketts, C.R. (1962). Biochem. J. 83, 533.
Carpenter, A.T. and Scott, J.J. (1961). Biochiyn. Biophys. Acta 52, 195.
Carpenter, T.M. and Benedict, F.G. (1909). Am. J. Physiol. 24, 187.
Carpenter, W.D. and Beevers, H. (1959). Plant Physiol. 34, 403.
Carr, A.J. (1963). Nature 198, 1104.
Cartwrigt, N.J. and Cain, R.B. (1959). Biochem. J. 71, 248.
Cascarano, J. and Zweifach, B.W. (1962). Federation Proc. 21, 151.
Castelfranco, P., Stumpf, P.K., and Contopoulou, C.R. (1955). J. Biol. Chem. 214, 567.
Caughey, W.S., Smiley, J.D., and Hellerman, L. (1957). J. Biol. Chem.. 224, 591.
Cavallito, C.J. and Haskell, T.H. (1945). J. Am. Chem. Soc. 67, 1991.
Cecil, R. and Snow, N.S. (1962). Biochem. J. 82, 247.
Cedrangolo, F. (1941). Atti Accad. Italia, Rend. Classe Sci. Fis., Mat. Nat. 2, 78.
Cedrangolo, F. and Adler, E. (1939). Enzymologia 7, 297.
Cerecedo, L.R. and Eich, S. (19.55). J. Biol. Chem. 213, 893.
Cerecedo, L.R., Soodak, M., and Eusebi, A.J. (1951). J. Biol. Chem. 189, 293.
Cerecedo, L.R., Eich, S., and Bresnick, E. (1954). Biochim. Biophys. Acta 15, 144.
Cerfontain, H. and van Aken, G.M.F.A. (1956). J. Am. Chem. Soc. 78, 2094.
Cha, C.-Y. and Scheraga, H.A. (1961a). Biochem. Biophys. Res. Commun. 5, 67.
Cha, C.-Y. and Scheraga, H.A. (1961b). Biochem. Biophys. Res. Commun. 6, 369.
Chaberek, S. and Martell, A.E. (1959). "Organic Sequestering Agents." Wiley, New
York.
Chaffee, R.R. and Bartlett, W.L. (1960). Biochim. Biophys. Acta 39, 370.
ChaikoflF, I.L. and Brown, G.W., Jr. (1954). In "Chemical Pathways of Metabolism"
(D.M. Greenberg, ed.). Vol. 1, p. 277. Academic Press, New York.
Challenger, F., Subramaniam, V., and Walker, T.K. (1927). J. Chem. Soc. p. 200.
Chambers, R., Cameron, G., and Kopac, M.J. (1943). Cancer Res. 3, 293.
Chance, B. (1954). In "The Mechanism of Enzyme Action" (W.D. McEkoy and B,
Glass, eds.), p. 399. Johns Hopkins Press, Baltimore, Maryland.
Chance, B. and Hollunger, G. (1961a). J. Biol. Chem. 236, 1534.
Chance, B. and Hollunger, G. (1961b). J. Biol. Chem. 236, 1555.
Chance, B. and Hollunger, G. (1961c). J. Biol. Chem. 236, 1562.
Chapman, D.W. and Shaffer, C.F. (1947). Arch. Internal Med. 79, 449.
Chappell, J.B. (1964 a). Biochem. J. 90, 225.
Chappell, J.B. (1964 b). Biochem. J. 90, 237.
Chappell, J.B. and Greville, G.D. (1958). Nature 182, 813.
Chappell, J.B. and Perry, S.V. (1955). Biochim. Biophys. Acta 16, 285.
Chari-Bitron, A. (1961). Biochem. Pharmacol. 6, 169.
Chari-Bitron, A. and Avi-Dor, Y. (1959 a). Biochim. Biophys. Acta 32, 276.
Chari-Bitron, A. and Avi-Dor, Y. (1959b). Biochem. J. 71, 572.
REFERENCES 999
Charmandarjan, M.O. and Tjutjunnikowa, A.W. (1930). Biochem. Z. Ill, 284.
Chattaway, F.W., Thompson, C.C, and Barlow, A.J.E. (1956). Biochem. J. 63, 648.
Chatterjee, G.C. and Haider, D.K. (1960). Naturwissenschaften 47, 86.
Chaudhuri, D.K. (1950 a). Ann. Biochem. Ezptl. Med. (Calcutta) 10, 65.
Chaudhuri, D.K. (1950 b). Ann. Biochem. Exptl. Med. (Calcutta) 10, 71.
Cheesman, D.F., Keeler, M.M., and Sten-Knudsen, O. (1959). Biochem.. J. 72, 357.
Cheftel, R.I., Munier, R., and Macheboeuf, M. (1952). Bull. Soc. Chim. Biol. 34,
380.
Cheldelin, V.H. and Beinert, H. (1952). Biochim.. Biophys. Acta 9, 661.
Chen, K.K., Rose, C.L., and Clowes, G.H.A. (1934). Am. J. Med. Sci. 188, 767.
Chen, R.F. and Plant, G.W.E. (1963). Biochemistry 2, 1023.
Chen, R.F., Brown, D.M., and Plant, G.W.E. (1964). Biochemistry 3, 552.
Cheniae, G. and Evans, H.J. (1959). Biochim. Biophys. Acta 35, 140.
Chiba, H., Kawai, F., and Kondo, K. (1954 a). Bull. Res. Inst. Food Sci., Kyoto Univ.
No. 13, p. 12.
Chiba, H., Kawai, F., and Kondo, K. (1954 b). Bull. Res. Inst. Food Sci., Kyoto Univ.
13, 23.
Chiba, H., Kawai, F., and Kondo, K. (1954 c). Bull. Res. Inst. Food Sci., Kyoto Univ.
13, 53.
Chiba, Y., Inada, A., and Yoshihara, I. (1953). Enzymologia 16, 148.
Chick, H. (1908). J. Hyg. 8, 92.
Chiga, M. and Plant, G.W.E. (1959). J. Biol. Chem. 234, 3059.
Chin, H.P. (1963). Ph.D. thesis. University of Southern California.
Chinard, F.P. (1942). Proc. Soc. Exptl. Biol. Med. 51, 317.
Chinard, F.P. and Hellerman, L. (1954). In "Methods of Biochemical Analysis" (D.
Glick, ed.). Vol. 1, p. 1. Wiley (Interscience), New York.
Chirigos, M.A., Greengard, P., and Udenfriend, S. (1960). J. Biol. Chem. 235, 2075.
Choppin, P.W. and Phihpson, L. (1961). J. Exptl. Med. 113, 713.
Christensen, E. and Wick, A.N. (1963). Proc. Soc. Exptl. Biol. Med. 112, 430.
Christensen, E., Brooke, J.L., Stewart, C.J., and Wick, A.N. (1961). Biochem. Biophys.
Res. Commun. 5, 209.
Christensen, H.N. and Riggs, T.R. (1952). J. Biol. Chem. 194, 57.
Christensen, H.N. and Ross, W.F. (1941). J. Biol. Chem. 137, 101.
Christensen, H.N. and Streicher, J.A. (1949). Arch. Biochem. 23, 96.
Christensen, H.N., Riggs, T.R., and Coyne, B.A. (1954). J. Biol. Chem. 209, 413.
Christie, G.S., Judah, J.D., and Rees, K.R. (1953). Proc. Roy. Soc. B141, 523.
Christman, A.A. and Lewis, H.B. (1921). J. Biol. Chem. 47, 495.
Chung, A.E. and Langdon, R.G. (1963). J. Biol. Chem. 238, 2317.
Cianci, V. (1940). Boll. Soc. Ital. Biol. Sper. 15, 288.
Ciferri, O. (1962). Enzymologia 24, 283.
Ciferri, O. and Blakley, E.R. (1959). Can. J. Microbiol. 5, 547.
Citri, N. and Garber, N. (1960). Biochim. Biophys. Acta 38, 50.
Citri, N. and Garber, N. (1961). Biochem. Biophys. Res. Commun. 4, 143.
Citri, N. and Garber, N. (1963). Biochim. Biophys. Acta 67, 64.
Clagett, CO., Tolbert, N.E., and Burris, R.H. (1949). J. Biol. Chem. 178, 977.
Clark, D.S. and Wallace, R.H. (1958). Can. J. Microbiol. 4, 125.
Clark, W.M. (1960). "Oxidation-reduction Potentials of Organic Systems." Williams
& Wilkins, Baltimore, Maryland.
1000 REFERENCES
Clarke, D.D., Mycek, M.J., Neidle, A., and Waelsch, H. (1959). Arch. Biochem. Bio-
phys. 79, 338.
Clarkson, T.W. and Crpss, A.C. (1961). AEC Research and Development Report (UR-
588), University of Rochester, Rochester, New York.
Clarkson, T.W. and Kench, J.E. (1956). Biochem. J. 62, 361.
Claus, D. (1956). Arch. Mikrohiol. 25, 10.
Clayton, R.A. (1959). Arch. Biochem. Biophys. 79, HI.
Clayton, R.K. (1957). Arch. Mikrohiol. 26, 29.
Clayton, R.K., Dettmer, F.H., and Robertson, A.E., Jr. (1957). Arch. Mikrohiol. 26, 20.
Cleland, K.W. (1949). Proc. Linnean Soc. N. S. Wales 74, 296.
Cleland, K.W. and Rothschild, Lord (1952 a). J. Exptl. Biol. 29, 285.
Cleland, K.W. and Rothschild, Lord (1952 b). J. Exptl. Biol. 29, 416.
Cleland, R. and Bonner, J. (1956). Plant Physiol. 31, 350.
Cleland, W.W. (1964). Biochemistry 3, 480.
Clementi, A. (1929). Boll. Soc. Ital. Biol. Sper. 4, 503.
Clements, A.N. (1959). J. Exptl. Biol. 36, 665.
ClifTe, E.E. and Waley, S.G. (1961). Biochem. J. 79, 475.
Cochrane, V.W. (1952). J. Bacteriol. 63, 459.
Cochrane, V.W. and Peck, H.D., Jr. (1953). J. Bacteriol. 65, 37.
Coggeshall, R.E. and MacLean, P.D. (1958). Proc. Soc. Exptl. Biol. Med. 98, 6871
Cohen, E.M. (1953 a). Acta Physiol. Pharmacol. Neerl. 3, 45.
Cohen, E.M. (1953 b). Acta Physiol. Pharmacol. Neerl. 3, 59.
Cohen, L.H. and Bridger, W.A. (1964). Can. J. Biochem. Physiol. 42, 715.
Cohen, L.H. and Parks, R.E., Jr. (1963). Can. J. Biochem. Physiol. 41, 1495.
Cohen, P.P. and Hayano, M. (1946). J. Biol. Chem. 166, 239.
Cohen, R.A. and Gerard, R.W. (1937). J. Cellular Cornp. Physiol. 10, 223.
Cohen, S. (1963). Exptl. Cell Res. 29, 207.
Cohn, M. and Horibata, K. (1959). J. Bacteriol. 78, 624.
Coleman, G.H., Weed, L.A., and Myers, CD. (1937). J. Am. Chem. Soc. 59, 2703.
Coleman, J.E. and Vallee, B.L. (1961). J. Biol. Chem. 236, 2244.
Coleman, R. and Hiibscher, G. (1962). Biochim. Biophys. Acta 56, 479.
Collander, R. (1951). Acta Chem. Scand. 5, 774.
Collias, E.G., McShan, W.H., and Lilly, J.H. (1952). J. Cellular Comp. Physiol. 40,
507.
Colowick, S.P., Lazarow, A., Packer, E., Schwarz, D.R., Stadtman, E.R., and Waelsch,
H. (eds.) (1954). "Glutathione," Academic Press, New York.
Colter, J.S., Kuhn, J., and Ellem, K.A.O. (1961). Cancer Res. 21, 48.
Commoner, B. and Hollocher, T.C., Jr. (1960). Proc. Natl. Acad. Sci. U. S. 46, 405.
Commoner, B. and Thimann, K.V. (1941). J. Gen. Physiol. 24, 279.
Conchie, J. (1953). Biochem. J. 55, xxi.
Conchie, J. (1954). Biochem. J. 58, 552.
Conchie, J. and Hay, A.J. (1959). Biochem. J. 73, 327.
Conchie, J. and Levvy, G.A. (1957). Biochem. J. 65, 389.
Conn, E.E. and Young, L.C.T. (1957). J. Biol. Chem. 226, 23.
Conover, T.E., Danielson, L., and Ernster, L. (1963a). Biochim. Biophys. Acta 67, 254.
Conover, T.E., Prairie, R.L., and Packer, E. (1963 b). J. Biol. Chem. 238, 2831.
Conrad, (1874). Ber. 7, 688. (Cited by WisUcenus, J.)
Converse, J.L. and Besemer, A.R. (1959). J. Bacteriol. 78, 231.
REFERENCES 1001
Conway, T.W., Lansford, E.M., Jr., and Shive, W. (1964). Arch. Biochem. Biophys.
107, 120.
Cooil, B.J. (1952). Plant Physiol. 27, 49.
Cook, A.M. and Steel, K.J. (1959). J. Pharm. Pharmacol. 11, 666.
Cook, E.S. and Kreke, C.W. (1943). Proc. Sac. Exptl. Biol. Med. 53, 222.
Cook, E.S. and Perisutti, G. (1947). J. Biol. Chem. 167, 827.
Cook, E.S., Kreke, C.W., McDevitt, M., and Bartlett, M.D. (1946). J. Biol. Chem.
162, 43.
Cook, R.P. (1930). Biochem. J. lA, 1538.
Cook, S.F. (1926). J. Gen. Physiol. 9, 575.
Coombs, T.L., Omote, Y., and Vallee, B.L. (1964). Biochemistry 3, 653.
Cooper, C. (1958 a). Biochim. Biophys. Acta 30, 484.
Cooper, C. (1958 b). Biochim. Biophys. Acta 30, 529.
Cooper, C. and Kulka, R.G. (1961). J. Biol. Chem. 236, 2351.
Cooper, C. and Lehninger, A.L. (1957). J. Biol. Chem. 224, 561.
Cooper, E.A. and Goddard, A.E. (1957). J. Appl. Chem. 7, 613.
Cooper, E.A. and Mason, J. (1927). J. Hyg. 26, 118.
Cooper, J.A.D., Smith, W., Bacila, M., and Medina, H. (1959). J. Biol. Chem. 234,
445.
Cooperstein, S.J. (1963). J. Biol. Chem. 238, 3606.
Coore, H.G. and Randle, P.J. (1964). Biochem. J. 91, 56.
Copenhaver, J.H., Jr. and Lardy, H.A. (1952). J. Biol. Chem. 195, 225.
Cori, G.T. and Cori, C.F. (1940). J. Biol. Chem. 135, 733.
Cori, G.T., Colowick, S.P., and Cori, C.F. (1939). J. Biol. Chem. 127, 771.
Corley, R.C. and Rose, W.C. (1926). J. Pharmacol. Exptl. Therap. 27, 165.
Cormier, M.J. (1960). Biochim. Biophys. Acta 42, 333.
Cormier, M.J., Totter, J.R., and Rostorfer, H.H. (1956). Arch. Biochem. Biophys. 63,
414.
Corwin, L.M. (1959). J. Biol. Chem. 234, 1338.
Corwin, L.M. and Schwarz, K. (1963). Arch. Biochem. Biophys. 100, 385.
Cottrell, T.L. (1954). "The Strengths of Chemical Bonds." Academic Press, New York.
Cousins, F.B. (1956). Biochem. J. 64, 297.
Covin, J.M. (1961). Ph.D. thesis, University of Southern California, Los Angeles.
Covin, J.M. and Berman, D.A. (1956). J. Pharmacol. Exptl. Therap. 117, 443.
Cowgill, R.W. (1959). J. Biol. Chem. 234, 3146.
Cowgill, R.W. and Pizer, L.I. (1956). J. Biol. Chem. Ill, 885.
Coxon, R.V. and Robinson, R.J. (1956). Proc. Roy. Soc. B145, 232.
Cramer, F.B. and Woodward, G.E. (1952). J. Franklin Inst. 253, 354.
Crandall, D.I. (1955). J. Biol. Chem. 212, 565.
Crandall, D.I. (1959). Federation Proc. 18, 208.
Crane, R.K. (1960). Biochim. Biophys. Acta 45, 477.
Crane, R.K. and Sols, A. (1953). J. Biol. Chem. 203, 273.
Crane, R.K. and Sols, A. (1954). J. Biol. Chem. 210, 597.
Cravens, W.W. and Snell, E.E. (1949). Proc. Soc. Exptl. Biol. Med. 71, 73.
Crawford, M.A. (1963). Biochem. J. 88, 115.
Crawhall, J.C. and Watts, R.W.E. (1962). Biochem. J. 85, 163.
Creaser, E.H. and Scholefield, P.G. (1960). Cancer Res. 20, 257.
Cramer, J.E. (1962). J. Neurochem. 9, 289.
1002 REFEKENCES
Cremer, J.E. (1964). J. Neurochem. 11, 165.
Cremona, T. (J 964). J. Biol. Chem. 239, 1457.
Creveling, C.R., Daly, J.W., Witkop, B., and Udenfriend, S. (1962). Biochim. Biophys.
Acta 64, 125.
Crewther, W.G. (1953). Australian J. Biol. Sci. 6, 205.
Crewther, W.G. (1956). Biochim. Biophys. Acta 21, 178.
Cross, R.J. and Taggart, J.V. (1950). Am. J. Physiol. 161, 181.
Crout, J.R. (1961). Biochem. Pharmacol. 6, 47.
Cruickshank, D.H. and Isherwood, F.A. (1958). Biochem. J. 69, 189.
Cumings, J.N. (1959). "Heavy Metals and the Brain." Thomas, Springfield, Illinois.
Cummins, J.T., Cheldelin, V.H., and King, T.E. (1957). J. Biol. Chem. 226, 301.
Cunningham, L.W. and Nuenke, B.J. (1959). J. Biol. Chem. 234, 1447.
Cunningham, L.W. and Nuenke, B.J. (1960). Biochim. Biophys. Acta 39, 565.
Cunningham, L.W. and Nuenke, B.J. (1961). J. Biol. Chem. 236, 1716.
Cunningham, L.W., Nuenke, B.J., and Strayhorn, W.D. (1957). J. Biol. Chem. 228,
835.
Czekalowski, J.W. (1952). Brit. J. Exptl. Physiol. 33, 57.
Dahl, J.L., Way, J.L., and Parks, R.E., Jr. (1959). J. Biol. Cheyn. 234, 2998.
Dahlqvist, A. (1959). Acta Chem. Scand. 13, 2156.
Dajani, R.M., Danielski, J., Gamble, W., and Orten, J.M. (1961). Biochem. J. 81,
494.
Dale, R.A. and Sanderson, P.H. (1954). Brit. J. Pharmacol. 9, 210.
Dalgarno, L. and Birt, L.M. (1962). Biochem. J. 83, 202.
Dalziel, K. (1962). Nature 195, 384.
Damaschke, K. and Liibke, M. (1958). Z. Naturforsch. 13b, 172.
Danforth, W. (1953). Arch. Biochem.. Biophys. 46, 164.
Daniel, L.J. and Norris, L.C. (1949). Proc. Soc. Exptl. Biol. Med. 72, 165.
Dann, O.T. and Carter, C.E. (1964). Biochem. Pharmacol. 13, 677.
Darlington, W.A. and Quastel, J.H. (1953). Arch. Biochem. Biophys. 43, 194.
Das, H.K. and Roy, S.C. (1961). Biochim. Biophys. Acta 51, 378.
Das, H.K. and Roy, S.C. (1962). Arch. Biochem. Biophys. 96, 445.
Das, N.B. (1937 a). Biochem. J. 31, 1116.
Das, N.B. (1937 b). Biochem. J. 31, 1124.
Das, N.B. and Roy, A. (1943). Trans. Bose Res. Inst. (Calcutta) 15, 91.
Das, S.K. and Chatterjee, G.C. (1962). J. Bacteriol. 83, 1251.
Das, S.N. and Ives, D.J.G. (1961). Proc. Chem. Soc. p. 373.
Datta, A.G. and Racker, E. (1961). J. Biol. Chem. 236, 617.
Daus, L., Meinke, M., and Calvin, M. (1952). J. Biol. Chem. 196, 77.
DaVanzo, J.P., Greig, M.E., and Cronin, M.A. (1961). Am. J. Physiol. 201, 833.
DaVanzo, J.P., Matthews, R.J., and Stafford, J.E. (1964a). Toxicol. Appl. Pharmacol.
6, 388.
DaVanzo, J. P., Matthews, R.J., Young, G.A., and Wingerson, F. (1964 b). Toxicol.
Appl. Pharmacol. 6, 396.
Davenport, H.W. (1954). Federation Proc. 13, 32.
Davenport, H.W. and Chavre, V.J. (1956). Am. J. Physiol. 187, 227.
Davenport, H.W., Chavre, V.J., and Davenport, V.D. (1954). Am. J. Physiol. 177, 418.
Davenport, H.W., Chavre, V.J., and Davenport, V.D. (1955). Am. J. Physiol. 182, 221.
REFERENCES 1003
Davies, G.E., Driver, G.W., Hoggarth, E., Martin, A.R., Paige, M.F.C., Rose, F.L.,
and Wilson, B.R. (1956). Brit. J. Pharmacol. 11, 351.
Davis, F.F. and Allen, F.W. (1955). Federation Proc. 14, 200.
Davison, D.C. (1949). Proc. Linnean Soc. N. 8. Wales 74, 37.
Dawes, E.A. and Holms, W.H. (1958). J. Bacterial. 75, 390.
Dawson, R.M.C. (1953). Biochem. J. 55, 507.
Dawson, R.M.C. and Thompson, W. (1964). Biochem. J. 91, 244.
Day, M.D. and Rand, M.J. (1963). Brit. J. Pharmacol. 22, 72.
de Caro, L., Rindi, G., and Grana, E. (19.54). Experientia 10, 140.
de Caro, L., Rindi, G., Perri, V., and Ferrari, G. (1956). Expeientia 12, 300.
Decker, R.H., Kang, H.H., Leach, F.R.. and Henderson, L.M. (1961). J. Biol. Chem.
236, 3076.
Decker, R.H.. Brown, R.R., and Price, J.M. (1963). J. Biol. Chem. 238, 1049.
DeGraff, A.C. and Lehman, R.A. (1942). J. Am. Med. Assoc. 119, 998.
DeGroot, C.A., Weber, J.F., and Cohen, E.M. (1955). Arch. Intern. Pharmacodyn. 102,
459.
DeGroot, N. and Cohen, P.P. (1962). Biochim. Biophtjs. Acta 59, 588.
de Issaly, I.S. and Stoppani, A.O.M. (1964). Proc. Soc. Exptl. Biol. Med. 115, 528.
Deitrich, R.A. and Hellerman, L. (1963). J. Biol. Chem. 238, 1683.
Dejung, K. (1963). Z. Ges. Exptl. Med. 137, 196.
Dellert, E.E. and Stahmann, M.A. (1955). Nature 176, 1028.
Delnionte, L. and Jukes, T.H. (1962). Pharmacol. Rev. 14, 91.
. Demis, D.J. and Rothstein, A. (1955). Am. J. Physiol. 180, 566.
DeMoss, J.A. and Swim, H.E. (1957). J. Bacterial. 74, 445.
DeMoss, R.D. (1955). In "Methods in Enzymology" (S.P. Colowiek and N.O. Kaplan,
eds.). Vol. 1, p. 504. Academic Press, New York.
Dempsey, W'.B. and Snell, E.E. (1963). Biochemistry 2, 1414.
Dengler, H. and Reichel, G. (1958). Arch. Exptl. Pathol. Pharmakol. 234, 275.
Denison, F.W^, Jr. and Phares, E.F. (1952). Anal. Chem. 24, 1628.
c^ennis, D. and Kaplan, N.O. (1960). J. Biol. Chem. 235, 810.
De Robertis, E. and Peluifo, C.A. (1951). Proc. Soc. Exptl. Biol. Med. 78, 584.
Dervartanian, D.V. and Veeger, C. (1962). Biochem. J. 84, 65p.
Desnuelle, P. and Antonin, S. (1946). Helv. Chim. Acta 29, 1306.
Desnuelle, P. and Rovery, M. (1949). Biochim. Biophys. Acta 3, 26.
Desnuelle, P., Antonin, S., and Casal. A. (1947). Bull. Soc. Chim. Biol. 29, 694.
DeTurk, W.E. and Bernheim, F. (1960). Arch. Biochem. Biophys. 90, 218.
Dewey, D.L., Hoare, D.S., and Work, E. (1954). Biochem. J. 58, 523.
Deykin, D. and Goodman, D.S. (1962). J. Biol. Chem. 237, 3649.
Dhar, S.C. and Bose, S.M. (1962). Enzymologia 24, 155.
Di Carlo, F.J. and Redfern, S. (1947). Arch. Biochem. 15, 343.
Dickens, F. (1946 a). Biochem. J. 40, 145.
Dickens, F. (1946 b). Biochem. J. 40, 171.
Dickens, F. and Glock, G.E. (1951). Biochem. J. 50, 81.
Dickens, F. and Williamson, D.H. (1956). Biochem. J. 64, 567.
Dickerman, H.W., San Pietro, A., and Kaplan, N.O. (1962). Biochim. Biophys. Acta
62, 230.
Dickman, S.R. (1958). Science 127, 1392.
Dickman, S.R. and W^estcott, W.L. (1953). Federation Proc. 12, 197.
1004 REFERENCES
Diczfalusy, E., Ferno, O., Fex, H., Hogberg, B., Linderot, T., and Rosenberg, T.
(1953). Acta Chem. Scand. 7, 913.
Dieterle, W. and Wenzel, F. (1944). Biochem. Z. 316, 357.
Dietrich, E.V., Fisher, A.L., Long, J.P., and Featherstone, R.M. (1952). Arch. Bio-
chem. Biophys. 41, 118.
Dietrich, L.S. and Borries, E. (1956). Arch. Biochem. Biophys. 64, 512.
Dietrich, L.S. and Shapiro, D.M. (1953 a). Proc. Soc. Exptl. Biol. Med. 84, 555.
Dietrich, L.S. and Shapiro, D.M. (1953 b). J. Biol. Chem. 203, 89.
Dietrich, L.S., Monson, W.J., Williams, J.N., Jr., and Elvehjem, C.A. (1952). J. Biol.
Chem. 1 97, 37.
Dietrich, L.S., Friedland, I.M., and Kaplan, L.A. (1958). J. Biol. Chem.. 233, 964.
Dikstein, S. (1959). Biochim. Biophys. Acta 36, 397.
Dilley, R.A. and Vernon, L.P. (1964). Biochemistry 3, 817.
Dils, R. and Popjak, G. (1962). Biochem. J. 83, 41.
D'lorio, A. (1957). Ca7i. J. Biochem. Physiol. 35, 395.
D'lorio, A. and Mavrides, C. (1963). Can. J. Biochem. Phijsiol. 41. 1779.
Dippy, J.F.J. (1939). Chem. Rev. 25, 151.
Di Sabato, G. and Kaplan, N.O. (1963). Biochemistry 2, 776.
Dische, Z. and Ash well, G. (1955). Biochim. Biophys. Acta 17, 56.
Dittrich, S. (1963). J. Chromatog. 12, 47.
Dixon, G.H., Kornberg, H.L., and Lund, P. (1960). Biochim. Biophys. Acta 41, 217.
Dixon, K.C. and Harrison, K. (1932). Biochem. J. 26, 1954.
Dixon, M. (1925). Biochem. J. 19, 507.
Dixon, M. and Thurlow, S. (1924). Biochem. J. 18, 976.
Doctor, V.M. (1958). J. Biol. Chem. 232, 617.
Doctor, V.M. (1959). Blood 14, 1244.
Dodgson, K.S. (1959). Enzymologia 20, 301.
Dodgson, K.S. and Powell, G.M. (1959). Biochem. J. 73, 672.
Dodgson, K.S., Spencer, B., and Williams, K. (1955). Biochem. J. 61, 374.
Doi, R.H. and Halvorson, H. (1961). J. Bacteriol. 81, 51.
Doisy, R.J., Richert, D.A., and Westerfeld, W.W. (1955). J. Biol. Chem. 217, 307.
Dolin, M.I. (1957). J. Biol. Chem. 225, 557.
Dolin, M.I. (1959). J. Bacteriol. 77, 383.
DoHn, M.I. and Gunsalus, I.C. (1951). J. Bacteriol. 62, 199.
Dolin, M.I. and Wood, N.P. (1960). J. Biol. Chem. 235, 1809.
Dominguez, A.M. and Shideman, F.E. (1953). Federation Proc. 12, 316.
Dominguez, A.M. and Shideman, F.E. (19.55). Proc. Soc. Exptl. Biol. Med. 90, 329.
Domonkos, J. and Latzkovits, L. (1961). Arch. Biochem. Biophys. 95, 144.
Donovan, J.W. (1964). Biochemistry 3, 67.
Doran, D.J. (1957). J. Protozool. 4, 182.
Dorfman, A., Roseman, S., Ludowieg, J., Mayeda, M., Moses, F.E., and Cifonelli,
J.A. (19.55). J. Biol. Chem. 216, 549.
Doudoroff, M., Contopoulou, C.R., and Burns, S. (1958). Proc. Intern. Symp. Enzyme
Chem., Tokyo Kyoto 1957 p. 313. Maruzen, Tokyo.
Dove, W.F. and Yamane, T. (1960). Biochem. Biophys. Res. Commun. 3, 608.
Dowdle, E.B., Schachter, D., and Schenker, H. (1960). Am. J. Physiol. 198, 609.
Doyle, M.L., Katzman, P.A., and Doisy, E.A. (1955). J. Biol. Chem. 217, 921.
Drabikowski, W. and Gergely, J. (1963). J. Biol. Chem. 238, 640.
REFERENCES 1005
Dreser, H. (1893). Arch. Exptl. Pathol. Pharmakol. 32, 456.
Dreyfus, P.M. and Moniz, R. (1962). Biochim. Biophys. Acta 65, 181.
Dube, S.K., Rohult, O., and Pressman, D. (1963). J. Biol. Chem. 238, 613.
du Buy, H.G. and Hesselbach, M.L. (1956). J. Histochem. Cytochem. 4, 363.
Duerkson, J.D. and Halvorson, H. (1958). J. Biol. Chem. 233, 1113.
Duggan, D.E. and Titus, E.G. (1959). J. Biol. Chem. 234, 2100.
Duggan, D.E., Weigert, M.G., and Titus, E.G. (1961). Cancer Res. 21, 1047.
Dumont, J.E. (1961). Biochim. Biophys. Acta 50, 506.
Dumont, J.E. (1962). Biochim. Biophys. Acta 56, 382.
Duperon, R. (1956). Rev. Gen. Botan. 63, 137.
Durham, N.N. and Hubbard, J.S. (1959). Nature 184, 1398.
Durham, N.N. and Hubbard, J.S. (1960). J. Bacteriol. 80, 225.
Dyer, H.M. (1938). J. Biol. Chem.. 124, 519.
Dziirik. R. and Krajci-Lazary, B. (1962). Arch. Intern. Pharmacodyn. 135, 1.
Dzurik. R.. Brixova, E., Krajci-Lazary, B., Hostynova, D., Kolesar, P., and Nieder-
land, T.R. (1963). Arch. Intern. Pharmacodyn. 141, 396.
Eaton, M.D. (1952). Arch. Ges. Virusforsch. 5, 53.
Eaton, M.D. and Scala, A.R. (1957). Arch. Ges. Virusforsch. 7, 274.
Eberson, L. and Wadso, I. (1963). Acta Chem. Scand. 17, 1552.
Eberts, F.S., Burris, R.H., and Riker, A.J. (1951). Am. J. Botany 38, 618.
Edelhoch, H., Katchalski, E., Maybury, R.H., Hughes, W.L., Jr., and Edsall, J.T.
(1953). J. Am. Chem. Soc. 75, 5058.
Edlbacher, S. and Zeller, A. (1936). Z. Physiol. Chem. 242, 253.
Edlbacher, S., Baur, H., and Becker. M. (1940). Z. Physiol. Chem. 265, 61.
Edman, K.A.P. (1958). Acta Physiol. Scand. 43, 275.
Edman, K.A.P. (1959). Acta Physiol. Scarid. 46, 209.
Edsall, J.T. and Wyman, J. (1958). "Biophysical Chemistry," Vol. 1. Academic Press,
New York.
Edsall, J.T., Maybury, R.H., Simpson, R.B., and Straessle, R. (1954). J. Am. Chem.
Soc. 76, 3131.
Edson, N.L. (1936). Biochem. J. 30, 1855.
Edson, N.L. and Hunter, G.J.E. (1947). Biochem. J. 41, 139.
Edson, N.L. and Leloir, L.F. (1936). Biochem. J. 30, 2319.
Edwards, J.G. (1942). Am. J. Pathol. 18, 1011.
Eeg-Larsen, N. and Laland, S.G. (1954). Acta Physiol. Scand. 30, 295.
Egami, F. and Yagi, K. (1956). J. Biochem. (Tokyo) 43, 153.
Egami, F., Naoi, M., Tada, M., and Yagi, K. (1956). J. Biochem. (Tokyo) 43, 669.
Eggerer, H. and Remberger, U. (1963). Biochem. Z. 337, 202.
Ehrmantraut, H. and Rabinowitch, E. (1952). Arch. Biochem. Biophys. 38, 67.
Eich, S. and Cerecedo, L.R. (19.54). J. Biol. Chem. 207, 295.
Eich, S. and Cerecedo, L.R. (1955). Arch. Biochem. Biophys. 57, 285.
Eichel, B. and Wainio, W.W. (1948). J. Biol. Chem. 175, 155.
Eichel, H.J. (1956 a). J. Biol. Chem. Ill, 137.
Eichel, H.J. (1956b). Biochim. Biophys. Acta 11, 571.
Eichel, H.J. (19.59). Biochem. J. 71, 106.
Eichel, H.J. and Rem, L.T. (1959). Biochim. Biophys. Acta 35, 571.
Eichel, H.J. and Rem, L.T. (1962). J. Biol. Chem. 237, 940.
1006 REFERENCES
Eisenstadt, J.M., Grossman, L., and Klein, H.P. (1959). Biochim. Biophys. Acta 36,
292.
Eldjarn, L. and Bremer, J. (1962). Biochem.. J. 84, 286.
Eldjarn, L. and Pihl, A. (1957 a). J. Am. Chem. Soc. 79, 4589.
Eldjarn, L. and Pihl, A. (1957 b). J. Biol. Chem. 225, 499.
El Ha wary, M.F.S. (1955). Biochem. J. 61, 348.
Elion, G.B., Callahan, S., Nathan, H., Bieber, S., Bundles, R.W., and Hitchings, G.H.
(1963). Biochem. Pharmacol. 12, 85.
Elkins-Kaufman, E. and Naurath, H. (1949). J. Biol. Chem. 178, 645.
Ellem, K.A.O. and Colter, J.S. (1961). J. Cellular Comp. Physiol. 58, 267.
Ellfolk, N. (1953). Acta Chem. Scand. 7, 1155.
Ellfolk, N. (1954). Acta Chem. Scand. 8, 151.
ElUott, H.W. and Sutherland, V.C. (1952). J. Cellular Comp. Physiol. 40, 221.
Elliott, K.A.C. and Greig, M.E. (1937). Biochem. J. 31, 1021.
Elliott, W.H. (1955). In "Methods in Enzymology" (S.P. Colowick and N.O. Kaplan,
eds.). Vol. 2, p. 337. Academic Press, New York.
Ellis, R.J. and Davies, D.D. (1961). Biochem. J. 78, 615.
Ellis, S. and Anderson, H.L. (1951a). J. Pharmacol. Exptl. Therap. 101, 10.
Ellis, S. and Anderson, H.L. (1951b). J. Pharmacol. Exptl. Therap. 103, 342.
Ellman, G.L. (1959). Arch. Biochem. Biophys. 82, 70.
Elodi, P. (1960). Biochim. Biophys. Acta 40, 272.
El'tsina, N.V. and Beresotskaya, N.A. (1962). Biochemistry {USSR) (English Transl.)
27, 386.
Eltz, R.W. and Vandemark, P.J. (1960). J. Bacteriol. 79, 763.
Ely, J.O., TuU, F.A., and Hard, J.A. (1952). J. Franklin Inst. 253, 361.
Emerson, G.A. and Southwick, P.L. (1945). J. Biol. Chem. 160, 169.
Emerson, G.A. and Tishler, M. (1944). Proc. Soc. Exptl. Biol. Med. 55, 184.
Emerson, G.A., Wurtz, E., and Johnson, O.H. (1945). J. Biol. Chem. 160, 165.
Emerson, P.M., Wilkinson, J.H., and Withycombe, W.A. (1964). Nature 202, 1337.
Emery, T.F. (1963). Biochemistry 2, 1041.
Endahl, B.R. and Kochakian, CD. (1961). Biochim. Biophys. Acta 54, 15.
Engelhardt, W. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo Kyoto 1957 p. 163.
Maruzen, Tokyo.
Englard, S. and Colowick, S.P. (1957). J. Biol. Chem. lib, 1047.
Englard, S., Sorof, S., and Singer, T.P. (1951). J. Biol. Chem. 189, 217.
Ennor, A.H. and Rosenberg, H. (1954). Biochem. J. 57, 203.
Eny, D.M. (1951). Biochem. J. 50, 559.
Eppley, R.W. (1958). J. Gen. Physiol. 41, 901.
Eppley, R.W. (1960). Plant Physiol. 35, 637.
Erdos, E.G., Debay, C.R., and Westerman, M.P. (1960). Biochem. Pharmacol. 5, 173.
Erlenmeyer, H., Waldi, D., and Sorkin, E. (1948). Helv. Chim. Acta 31, 32.
Ernster, L., Danielson, L., and Ljunggren, M. (1962). Biochim. Biophys. Acta 58,
171.
Estler, C.-J., Heim, F., Moosburger, A., and Herzog, U. (1960). Arch. Exptl. Pathol.
Pharmakol. 238, 301.
Eubank, L.L., Huf, E.G., Campbell, A.D., and Taylor, B.B. (1962). J. Cellular Comp.
Physiol. 59, 129.
Eusebi, A.J. and Cerecedo, L.R. (1949). Science 110, 162.
Biol
. Chem.
130.
13.
102,
125.
REFERENCES 1007
Eusebi, A.J. and Cerecedo, L.R. (1950). Federation Proc. 9, 169.
Evans, E.A., Jr., Vennesland, B., and Slotin, L. (1943). J. Biol. Chem. 147, 771.
Evans, J.I. and Monk, C.B. (1952). Trans. Faraday Soc. 48, 934.
Evert, H.E. (1952). Arch. Biochem. Biophys. 41, 29.
Eyring, H. (1932). Phys. Rev. 39, 746.
Ezaki, S. (1940). J. Biochem. (Tokyo) 32, 107.
Fahrlander, H., Favarger, P., Nielsen, H., and Leuthardt, F. (1947). Helv. Physiol.
Pharmacol. Acta 5, 202.
Fahrlander, H., Favarger, P., and Leuthardt, F. (1948). Helv. Chim. Acta 31, 942.
Fain, J.N. (1964). J. Biol. Chem. 239, 958.
Fairclough, R.A. (1938). J. Chem. Soc. p. 1186.
Falcone, G., Salvatore, G., and CoveUi, I. (1959). Biochim. Biophys. Acta 36, 390.
Fanestil, D.D., Hastings, A.B., and Mahowald, T.A. (1963). J. Biol. Chem. 238, 836.
Farah, A. (1952). Arch. Exptl. Pathol. Pharmakol. 215, 29.
Farah, A. and Kruse, R. (1960). J. Pharmacol. Exptl. Therap.
Farah, A. and Mook, W. (1951). J. Pharmacol. Exptl. Therap.
Farah, A. and Rennick, B.R. (1954). Federation Proc. 13, 353.
Farah, A. and Rennick, B.R. (1956). J. Pharmacol. Exptl. Therap. 117, 478.
Farah, A., Mook, W., and Johnson, R. (1951). Proc. Soc. Exptl. Biol. Med. 76, 403.
Farah, A.E. (1957). In "MetaboHc Aspects of Transport Across Cell Membranes"
(Q.R. Murphy, ed.), p. 257. Univ. of Wisconsin Press, Madison, Wisconsin.
Farah, A.E. and Miller, T.B. (1962). Ann. Rev. Pharmacol. 2, 269.
Farkas, G.L. and Ledingham, G.A. (1959). Can. J. Microbiol. 5, 141.
Farkas, G.L., Konrad, £., and Kiraly, Z. (1957 a). Naturwissenschaften 44, 65.
Farkas, G.L., Konrad, fi., and Kiraly, Z. (1957 b). Physiol. Plantarum 10, 346.
Fasella, P. and Hammes, G.G. (1963). Arch. Biochem. Biophys. 100, 295.
Fasold, H., Groschel-Stewart, U., and Turba, F. (1964). Biochem. Z. 339, 487.
Fastier, F.N. (1962). Pharmacol. Rev. 14. 37.
Favelukes, G. and Stoppani, A.O.M. (1958). Biochim. Biophys. Acta 28, 654.
Fawaz, E.N. and Fawaz, G. (1962). Biochem. J. 83, 438.
Fawaz, E.N., Manoukian, E., and Fawaz, G. (1963). Biochem. Z. 337, 195.
Fawaz, G. and Fawaz, E.N. (1951). Proc. Soc. Exptl. Biol. Med. 77, 239.
Fawaz, G. and Fawaz, E.N. (1954). Proc. Soc. Exptl. Biol. Med. 87, 30.
Fawaz, G., Tutunji, B., and Fawaz, E.N. (1958). Proc. Soc. Exptl. Biol. Med. 97, 770.
Feigelson, P. and Davidson, J.D. (1956 a). Cancer Res. 16, 352.
Feigelson, P. and Davidson, J.D. (1956 b). J. Biol. Chem. 223, 65.
Feigelson, P., Williams, J.N., Jr., and Elvehjem, C.A. (1951). J. Biol. Chem. 189,
361.
Feigelson, P., Davidson, J.D.. and Robins. R.K. (1957). J. Biol. Chem. 116, 993.
Feig], F. (1960). "Spot Tests in Organic Analysis," 6th ed. Elsevier, Amsterdam.
Feinberg, R.H. and Greenberg, D.M. (1959). J. Biol. Chem. 234, 2670.
Feinstein, R.N., Cotter, G.J., and Hampton, M.M. (1954). Am. J. Physiol. 177, 156.
Feist, F. (1890). Ann. 257, 253.
Feldberg, W. and Kellaway, C.H. (1938). Australian J. Exptl. Biol. Med. Sci. 16, 249.
Feldman, W. and O'Kane, D.J. (1960). J. Bacteriol. 80, 218.
Felenbok, B. and Neufeld, E.F. (1962). Arch. Biochem. Bioj)hys. 97, 116.
Feller, D.D. and Feist, E. (1957). Federation Proc. 16, 36.
1008 REFERENCES
Fellig, J. and Wiley, C.E. (1959). Arch. Biochem.. Biophys. 85, 313.
Fellig, J. and Wiley, C.E. (1960). Science 132, 1835.
Fellman, J.H. (1956). Pr^oc. Soc. Exptl. Biol. Med. 93, 413.
Fellman, J.H. (1959). Enzymologia 20, 366.
Felts, J.M., Masoro, E.J., Panagos, S.S., and Rapport, D. (1956). Federation Proc.
15, 61.
Fernley, H.N. (1962). Biochem. J. 82, 500.
Ferno, 0., Fex, H., Hogberg, B., Linderot, T., and Rosenberg, T. (1953). Acta Chem.
Scand. 7, 921.
Ferrari, R.A. and Arnold, A. (1963 a). Biochim. Biophys. Acta 77, 349.
Ferrari, R.A. and Arnold, A. (1963 b). Biochim.. Biophys. Acta 77, 357.
Ferrari, R.A., Mandelstam, P., and Crane, R.K. (1959). Arch. Biochem. Biophys. 80,
372.
Ferreira, R., Ben-Zvi, E., Yamane, T. Vasilevskis, J., and Davidson, N. (1961). In
"Advances in the Chemistry of the Coordination Compounds," 6th Intern. Conf.
on Coordination Chem., p. 457. Macmillan, New York.
Festenstein, G.N. (1958). Biochem. J. 70, 49.
Festenstein, G.N. (1959). Biochem. J. 72, 75.
Fewson, C.A. and Nicholas, D.J.D. (1961). Biochim. Biophys. Acta 49, 335.
Fewster, J.A. (1957). Biochem. J. 66, 9p.
Fildes, P. (1940). Brit. J. Exptl. Pathol. 21, 67.
Findlay, J., Levvy, G.A., and Marsh, C.A. (1958). Biochem. J. 69, 467.
Finkle, B.J. and Smith, E.L. (1958). J. Biol. Chem. 230, 669.
Fischer, A. and Herrmann, H. (1937). Enzymologia 3, 180.
Fischer, F.G. and Eysenbach, H. (1937). Ann. 530, 99.
.^Fishbein, W.N. and Bessman, S.P. (1964). J. Biol. Chem. 239, 357.
'Fisher, F.M., Jr. (1964). Biol. Bull. 126, 220.
Fisher, H.F., Krasna, A.I., and Rittenberg, D. (1954). J. Biol. Chem. 209, 569.
Fishgold, J.T. (1957). Federation Proc. 16, 296.
Fishman, R.A. (1964). Am. J. Physiol. 206, 836.
Fishman, W.H. and Green, S. (1957). J. Biol. Chem. 225, 435.
Fitzgerald, R.J. and Jordan, H.V. (1953). Antihiot. Chemotherapy 3, 231.
Fiume, L. (1960). Nature 187, 792.
Flaks, J.G. and Cohen, S.S. (1959). J. Biol. Chem. 234, 2981.
Flamm, W.G. and Crandall, D.I. (1963). J. Biol. Chem. 238, 389.
Flatow, L. (1928). Biochem. Z. 194, 132.
Flavin, M. and Slaughter, C. (1960). J. Biol. Chem. 235, 1103.
Flavin, M., Ortiz, P. J., and Ochoa, S. (1955). Nature 176, 823.
Flesch, P. and Kun, E. (1950). Proc. Soc. Exptl. Biol. Med. 74, 249.
Fhckinger, R.A., Jr. (1949). J. Exptl. Zool. 112, 465.
Florkin, M. and Duchateua-Bosson, G. (1939). Bull. Soc. Chim. Biol. 21, 1204.
Fluharty, A.L. and Ballou, C.E. (1959). J. Biol. Chem. 234, 2517.
Fluri, R. (1959). Arch. Mikrobiol. 33, 195.
Fodor, P.J. (1952). Arch. Biochem. Biophys. 35, 311.
Folk, J.E. (1956). J. Am. Chem. Soc. 78, 3541.
Folk, J.E. and Gladner, J.A. (1958). J. Biol. Chem. 231, 379.
Fonnum, F., Haavaldsen, R., and Tangen, 0. (1964). J. Neurochem. 11, 109,
Forney, R.B. and Harger, R.N. (1949). Federation Proc. 8, 292.
REFERENCES 1009
Forsen, S. and Nilsson, M. (1961). Arkiv Kenii 17, 523.
Forssman, S. (1941). Acta Physiol. Scand. 2, Suppl. 5.
Forssman, S. and Lindsten, T. (1946). Acta Pharmacol. Toxicol. 2, 329.
Forster, R.P. and Goldstein, L. (1961). J. Cellular Camp. Physiol. 58, 247.
Forster, R.P. and Taggart, J.V. (1950). J. Cellular Comp. Physiol. 36, 251.
Forti, G. (1957). Physiol. Plantarum 10, 898.
Foss, O. (1961). In "Organic Sulfur Compounds" (N. Kharasch, ed.). Vol. 1, p. 83
Macmillan (Pergamon), New York.
Foster, R.J. (1961). J. Biol. Chem. 236, 2461.
Foster, R.J. and Niemann, C. (1955a). J. Am. Chem. Soc. 77, 3365.
Foster, R.J. and Niemann, C. (1955b). J. Am. Chem.. Soc. 77, 3370.
Foster, R.J., Shine, H.J., and Niemann, C. (1955). J. Am. Chem. Soc. 77, 2378.
Foulerton, A.G.R. (1921). J. Pathol. Bacteriol. 24, 257.
Foulkes, F.C. and Paine, CM. (1961). J. Biol. Chem. 236, 1019.
Fountain, J.R., Hutchinson, D.J., Waring, G.B., and Burchenal, J.H. (1953). Proc.
Soc. Exptl. Biol. Med. 83, 369.
Fourneau, E. and Melville, K.I. (1931). J. Pharmacol. Exptl. Therap. 41, 21.
Fournier, P., d'Auzac, J., Pujarniwcle, S., and Cuong, T.C. (1961). Compt. Rend. 252,
796.
Fraenkel-Conrat, H. (1955). J. Biol. Chem. 217, 373.
Fraenkel-Conrat, H. (1959). In "Sulfur in Proteins" (R. Benesch et al., eds.), p. 339.
Thomas, Springfield, Illinois.
Francis, M.J.O., Hughes, D.E., Romberg, H.L., and Phizackerley, P.J.R. (1963).
Biochem. J. 89, 430.
Franke, W. (1944 a). Z. Physiol. Chem. 280, 76.
Franks, W. (1944 b). Z. Physiol. Chem. 281, 162.
Fraser, D.M. and Kermack, W.O. (1957). Brit. J. Pharmacol. 12, 16.
Freedland, R.A., Wadzinski, I.M., and Waisman, H.A. (1961). Biochem. Biophys. Res.
Commun, 6, 227.
Freedlander, B.L., French, F.A., and Furst, A. (1956). Proc. Soc. Exptl. Biol. Med.
92, 533.
French, T.V., Dawid, I.B., Day, R.A., and Buchanan, J.M. (1963 a). J. Biol. Chem.
238, 2171.
French, T.C, Dawid, I.B., and Buchanan, J.M. (1963 b). J. Biol. Chem. 238, 2186.
Freudenberg, K. and Eyer, H. (1932). Z. Physiol. Chem. 213, 226.
Friberg, L. (1956). Acta Pharmacol. Toxicol. 12, 411.
Friberg, L. (1959). A.M.A. Arch. hid. Healt 20, 42.
Friberg, L., Odeblad, E., and Forssman, S. (1957). A.M.A. Arch. Ind. Health 16, 163.
Fridhandler, L. (1959). Federation Proc. 18, 48.
Fridhandler, L. and Quastel, J.H. (1955). Arch. Biochem. Biophys. 56, 424.
Fridhandler, L. Hafez, E.S.E., and Pincus, G. (1957). Exptl. Cell Res. 13, 132.
Fridovich, I. and Handler, P. (1954). Federation Proc. 13, 212.
Fridovich, I. and Handler, P. (1956). J. Biol. Chem. 223, 321.
Fridovich, I. and Handler, P. (1957). J. Biol. Chem. 228, 67.
Fridovich, I. and Handler, P. (1958). J. Biol. Chem. 231, 899.
Fried, G.H. and Antopol, W. (1957). J. Cellular Comp. Physiol 49, 201.
Friedemann, T.E. (1934). Science 80, 34.
Frieden, C (1959). J. Biol. Chem. 234, 809.
1010 REFERENCES
Frieden, C. (1962). Biochim. Biophys. Acta 59, 484.
Frieden, C. (1963). J. Biol. Chem. 238, 3286.
Frieden, E. and Maggiolo, I.W. (1957). Biochim. Biophys. Acta 24, 42.
Frieden, E. and Naile, B. (1954). Arch. Biochem. Biophys. 48, 448.
Frieden, E., Westmark, G.W., and Schor, J.M. (1961). Arch. Biochem. Biophys. 92,
176.
Friedland, I.M., Kaplan, L.A., Dietrich, L.S., and Shapiro, D.M. (1958). Federation
Proc. 17, 224.
Friedmann, H.C. and Vennesland, B. (1958). J. Biol. Chem. 233, 1398.
Friedmann, H.C. and Vennesland, B. (1960). J. Biol. Chem. 235, 1526.
Frimmer, M. (1960). Biochem. Z. 332, 522.
Frimmer, M. (1961). Arch. Exptl. Pathol. Pharmakol. lAl, 96.
Frisell, W.R. and Hellerman, L. (1957). J. Biol. Chem. 225, 53.
Frisell, W.R. and Mackenzie, C.G. (1955). J. Biol. Chem. 217, 275.
Frisell, W.R., Weinbach, E.G., and Hellerman, L. (1949). Federation Proc. 8, 200.
Frisell, W.R., Lowe, H.J., and Hellerman, L. (1956). J. Biol. Chem. Ill, 75.
Friz, C.T., Lazarow, A., and Cooperstein, S.J. (1960). Biol. Bull. 119, 161.
Frohman, C.E. and Day, H.G. (1949). J. Biol. Chem. 180, 93.
Frohman, C.E., Orten, J.M., and Smith, A.H. (1951). J. Biol. Chem. 193, 277.
Fromageot, C. and Grand, R. (1944). Enzymologia 11, 235.
Fromm, H.J. and Nordlie, R.C. (1959). Arch. Biochem. Biophys. 81, 363.
Fromni, H.J. and Zewe, V. (1962). J. Biol. Chem. 237. 3027.
Frommel, E., Herschberg, A.D., and Piquet, J. (1944). Helv. Physiol. Pharmacol. Acta
2, 169.
Fruton, J.S. and Mycek, M.J. (1956). Arch. Biochem. Biophys. 65, 11.
Fuentes, V. and Rubino, R. (1923). Anales Fac. Med. Montevideo 8, 360.
Fujie, Y. (1959). Japan. J. Pharmacol. 9, 6.
Fujimoto, D. and Ishimoto, M. (1961). J. Biochem,. (Tokyo) 50, 533.
Fuld, M. and Paul, M.H. (1952). Proc. Soc. Exptl. Biol. Med. 79, 355.
Fulton, J.D. and Christophers, S.R. (1938). Ann. Trop. Med. Parasitol. 32, 77.
Fulton, J.D. and Spooner, D.F. (1956 a). Exptl. Parasitol. 5, 59.
Fulton, J.D. and Spooner, D.F. (1956 b). Exptl. Parasitol. 5, 154.
Fulton, J.D. and Spooner, D.F. (1959). Exptl. Parasitol. 8, 137.
Fulton, J.D. and Spooner, D.F. (1960). Exptl. Parasitol. 9, 293.
Futterman, S. (1957). J. Biol. Chem. 218, 1031.
Futterman, S. and Silverman, M. (1957). J. Biol. Chem. 114, 31.
Gaebler, O.H. and Beher, W.T. (1951). J. Biol. Chem. 188, 343.
GafFney, T.J., Rosenberg, H., and Ennor, A.H. (1964). Biochem. J. 90, 170.
Gage, J.C. and Swan, A.A.B. (1961). Biochem.. Pharmacol. 8, 77.
Gajdos, A. (1939). Compt. Rend. Soc. Biol. 130, 1566.
Gale, G.R. (1961). Arch. Biochem. Biophys. 94, 236.
Galoyan, A.A. (1959). Tiolovye Soedin. v Med., Ukr. Nauchn.-Issled. Sanit.-Khim.
Inst., Tr. Nauchn. Konf., Kiev, 1957, p. 79.
Galoyan, S.A. and Turpaev, T.M. (1958). Dokl. Akad. Nauk Arm. SSR 27, 59.
Galston, A.W., Siegel, S.M., and Sharpensteen, H.H. (1955). Federation Proc. 14, 217.
Gamble, J.L., Jr. (1957). J. Biol. Chem. 228, 955.
Gamble, J.L., Jr. and Najjar, V.A. (1955). J. Biol. Chem. 217, 595.
REFERENCES 1011
Gamborg, O.L., Wetter, L.R., and Neish, A.C. (1961). Can. J. Biochem. Physiol. 39,
1113.
Gammon, G.D., Gumnit, R., Kanirin, R.P., and Kamrin, A. A. (1960). In "Inhibition in
the Nervous System and y-Aminobutyric Acid" (E. Roberts, ed.), p. 328. Macmillan
(Pergamon), New York.
Gane, R. and Ingold, O.K. (1931). J. Chem. Soc. p. 2153.
Garber, N. and Citri, N. (1962). Biochim. Biophys. Acta 62, 385.
Garcia-Hernandez, M. and Kun, E. (1957). Biochim. Biophys. Acta 24, 78.
Gardier, R.W. and Woodbury, R.A. (1955). J. Pharmacol. Exptl. Therap. 114, 453.
Gardner, E.A. and Farah, A. (1954). J. Pharmacol. Exptl. Therap. 111, 255.
Garen, A. and Levinthal, C. (1960). Biochim. Biophys. Acta 38, 470.
Garfinkel, D. (1957). Arch. Biochem. Biophys. 71, 100.
Gause, G.F. (1933). Protoplasma 17, 548.
Gayer, J. and Partowi, R. (1962). Z. Ges. Exptl. Med. 135, 419.
Gelfant, S. (1960). Exptl. Cell Res. 19, 72.
Gelles, E. (1956). J. Chem. Soc. p. 4736.
Gellhorn, A. and Jones, L.O. (1949). Blood 4, 60.
Gemmill, C.L. and Bowman, E.M. (1950). J. Pharmacol. Exptl. Therap. 100, 244.
Gemmill, C.L. and Hellerman, L. (1937). Am. J. Physiol. 120, 522.
Geppert, J. (1889). Berlin. Klin. Wochschr. 26, 789.
Gerarde, H.W., Jones, M., and Winnick, T. (1952). J. Biol. Chem. 196, 51.
Gerardin. C. and Kayser, F. (1959). Compt. Rend. Soc. Biol. 152. 1780.
Gergely, J. (1955). In "Methods in Enzymology" (S.P. Colowick and N.O. Kaplan,
eds.). Vol. 1, p. 602. Academic Press, New York.
Gergely, J. and Maruyama, K. (1960). J. Biol. Chem. 235, 3174.
Gergely, J., Martonosi, A., and Gouvea, M.A. (1959). In "Sulfur in Proteins" (R.
Benesch et al., eds.), p. 297. Academic Press, New York.
Gerhardt, P., Levine, H.B., and Wilson, J.B. (1950). J. Bacteriol. 60, 459.
Gerhart, J.C. and Pardee, A.B. (1961). Federation Proc. 20, 224.
Gerhart, J.C. and Pardee, A.B. (1962). J. Biol. Chem. 237, 891.
Gerhart, J.C. and Pardee, A.B. (1964). Federation Proc. 23, 727.
Gershanovich, V.N. (1963). Biochemistry (URRS) {English Transl.) 27, 868.
Gershenfeld, L. and Witlin, B. (1950). Ann. N. Y. Acad. Sci. 53, 172.
Gessler, U. and Bass, L. (1960). Z. Ges. Exptl. Med. 133, 18.
Gessler, U. and Kuner, E. (1960). Z. Ges. Exptl. Med. 133, 394.
Gest, H., Judis, J., and Peck, H.D., Jr. (1956). In "Inorganic Nitrogen Metabolism"
(W.D. McElroy and H.B. Glass, eds.), p. 298. Johns Hopkins Press, Baltimore,
Maryland.
Geuther (1866). Z. Chem. 2, 8. (Cited in Dyson, G.M. (1950). "A Manual of Organic
Chemistry," Vol. 1. Longmans, Green, New York.)
Geyer, R.P. and Cunningham, M. (1950). J. Biol. Chem. 184, 641.
Geyer, R.P., Matthews, L.W., and Stare, F.J. (1950a). J. Biol. Chem. 182, 101.
Geyer, R.P., Cunningham, M., and Pendergast, J. (1950 b). J. Biol. Cheyn. 188, 185.
Ghiretti, F., Ghiretti-Magaldi. A., and Tosi, L. (1959). J. Gen. Physiol. Al, 1185.
Ghosh, A.K. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo Kyoto 1957 p. 532.
Maruzen, Tokyo.
Ghosh, S., Blumenthal, H.J., Davidson, E., and Roseman, S. (1960). J. Biol. Chem.
235, 1265.
1012 REFERENCES
Gibbins, L.N. and Simpson, F.J. (1963). Can. J. Microbiol. 9, 769.
Gibbs, M. and Calo, N. (1959 a). Federation Proc. 18, 235.
Gibbs, M. and Calo, N. (1959 b). Plant Physiol. 34, 318.
Gibbs, M. and Calo, N. (1960 a). Federation Proc. 19, 329.
Gibbs, M. and Calo, N. (1960 b). Biochim. Biophys. Acta 44, 341.
Gibbs, M., Earl, J.M., and Ritchie, J.L. (1955). Plant Physiol. 30, 463.
Gibson, F. and Yanofsky, C. (1960). Biochim.. Biophys. Acta 43, 489.
Gibson, K.D., Laver, W.G., and Neuberger, A. (1958). Biochem. J. 70, 71.
Gibson, Q.H. and Roughton, F.J.W. (1955). Proc. Roy. Soc. B143, 310.
Giebel, O. and Passow, H. (1960). Arch. Oes. Physiol. T7^, 378.
Giebisch, G. (1958). J. Cellular Comp. Physiol. 51, 221.
Giebisch, G. (1960). Circulation 21, 879.
Giebisch, G. (1961). J. Oen. Physiol. 44, 6.59.
Giebisch, G. and Dornian, P.J. (1958). Proc. Soc. Exptl. Biol. Med. 98, 50.
Gilbert, D.A. (1963). Biochitn. Biophys. Acta 77, 657.
GilfiUan, R.F., Holtman, D.F., and Ross, R.T. (19.56). J. Bacteriol. 72, 620.
Gillespie, J.M. (1956). Australian J. Biol. Sci. 9, 448.
Gillette, J.R., Brodie, B.B., and La Du, B.N. (1957). J. Pharmacol. Exptl. Therap.
119, 532.
Gilman, A., Philips, F.S., Koelle, E.S., Allen, R.P., and St. John, E. (1946). Am. J.
Physiol. 147, 115.
Gilmour, D. (1960). Nature 186, 295.
Gilmour, D. and Gellert, M. (1961). Arch. Biochem. Biophys. 93, 605.
Gilmour, D. and Griffiths, M. (1957). Arch. Biochem. Biophys. 72, 302.
Gil y Gil, C. (1924). Beitr. Pathol. Anat. Allgem. Pathol. 72, 621.
Giovanelli, J. and Stumpf, P.K. (1957). Plant Physiol. 32, 498.
GiovanelH, J. and Stumpf, P.K. (1958). J. Biol. Chem. 231, 411.
Girerd, R.J., Tenebaum, L.E., Berkowitz, J., Rassaert, C.L., and Green, D.M. (1958).
J. Pharmacol. Exptl. Therap. 1 22, 25 AT
Giri, K.V., Krishnaswamy, P.R., and Rao. N.A. (1958). Biochem. J. 70, 66.
Giuditta, A. and Strecker, H.J. (1960). Biochem. Biophys. Res. Commun. 2, 159.
Glaser, L. and Brown, D.H. (1955). J. Biol. Chem. 216, 67.
Glasziou, K.T. (1956). Australian J. Biol. Sci. 9, 253.
Glazko, A.J. and Ferguson, J.H. (1940). Proc. Soc. Exptl. Biol. Med. 45, 43.
Glock, G.E. and McLean, P. (1953). Biochem. J. 55, 400.
Glynn, I.M. (1963). J. Physiol. (London) 169, 452.
Godeaux, J. (1944). Bull. Soc. Roy. Sci. Liege 13, 219.
Godzeski, C. and Stone, R.W. (1955). Arch. Biochem. Biophys. 59, 132.
Goebel, W.F. and Perlmann, G.E. (1949). J. Exptl. Med. 89, 479.
Goedkoop, J.A. and MacGillavry, C.H. (1957). Acta Crrjst. 10, 125.
Goerz, R.D. and Pengra, R.M. (1961). J. Bacteriol. 81, 568.
Goffart, M. and Fischer, P. (1948). Arch. Intern. Physiol. 55, 258.
Gold, A.H. and Segal, H.L. (1964). Biochemistry 3, 778.
Goldberg, G. (1963). Unpublished work.
Goldberg, L., Martin, L.E., and Leigh, J. (1962). Biochem. J. 85, 56.
Goldberg, L.I., DaCosta, F.M., and Ozaki, M. (1960). Nature 188, 502.
Goldman, D.S. (1956 a). J. Bacteriol. 71, 732.
Goldman, D.S. (1956 b). J. Bacteriol. 72, 401.
REFERENCES 1013
Goldman, D.S. (1959 a). Biochim. Biophys. Acta 32, 80.
Goldman, D.S. (1959 b). Biochim. Biophys. Acta 34, 527.
Goldschmidt, E.P., Yall, I., and Koffler, H. (1956). J. Bacterial. 72, 436.
Goldstein, A. and Doherty, M.E. (1951). Arch. Biochem. Biophys. 33, 35.
Goldstein, L., Van Baerle, R.R., and Dearborn, E.H. (1957). Enzymologia 18, 261.
Goldstein, M. and Contrera, J.F. (1961). Ezperientia 17, 267.
Goldstein, M. and Contrera, J.F. (1962). J. Biol. Chem. 237, 1898.
Goldstein, M.H., Levitt, M.F., Hauser, A.D., and Polimeros, D. (1961). J. Clin. In-
vest. 40, 731.
Goldstone, A. and Adams, E. (1962). J. Biol. Chem. 237, 3476.
Gonda, O., Traub, A., and Avi-Dor, Y. (1957). Biochem. J. 67, 487.
Gonnard, P. (1950). Bull. Soc. Chim. Biol. 32, 535.
Gonnert, R. and Bock, M. (1956). Arzneimittel-Forsch. 6, 522.
Gooder, H. and Happold, F.C. (1954). Biochem. J. 57, 369.
Goodman, D.S. (1961). J. Biol. Chem. 236, 2429.
Goodman, D.S. and Popjak, G. (1960). J. Lipid Res. 1, 286.
Goodman, I. and Hiatt, R.B. (1964). Biochem. Pharmacol. 13, 871.
Goodman, I., Fonts, J.R., Bresnick, E., Menegas, R., and Hitchings, G.H. (1959).
Science 130, 450.
Gopinathan, K.P., Sirsi, M., and Vaidyanathan, C.S. (1964). Biochem. J. 91, 277.
Gordon, J.J. (1953). Biochem. J. 55, 812.
Gordon, J.J. and Quastel, J.H. (1948). Biochem. J. 42, 337.
Gore, M.B.R. (1951). Biochem. J. 50, 18.
Goth, A. (1959). Am. J. Physiol. 197, 1056.
Gots, J.S. and Gollub, E.G. (1959). Proc. Soc. Exptl. Biol. Med. 101, 641.
Gottlieb, D. (1964). Endeavour 23, 85.
Gotto, A.M. and Komberg, H.L. (1961). Biochem. J. 81, 273.
Gourevitch, A., Pursiano, T.A., and Lein, J. (1961). Arch. Biochem. Biophys. 93, 283.
Govorov, N.P. (1936). Tr. Vses. Inst. Eksperim. Med. 1, No. 2, 147.
GraflBin, A.L., Gray, N.M., and Bagley, E.H. (1952). Bull. Johns Hopkins Hosp. 91,
137.
Granick, S. (1958). J. Biol. Chem. Ill, 1101.
Grant, B.R. and Beevers, H. (1964). Plant Physiol. 39, 78.
Grant, P.T. and Sargent, J.R. (1960). Biochem. J. 76, 229.
Grant, P.T. and Sargent, J.R. (1961). Biochem. J. 81, 206.
Grant, W.M. and Kinsey, V.E. (1946). J. Biol. Chem. 165, 485.
Grassmann, W., Klenk, L., and Peters-Mayr, T. (1935). Biochem. Z. 280, 307.
Gray, C.T. (1952). J. Bacteriol. 63, 813.
Gray, S.J. and Felsher, R.Z. (1945). Proc. Soc. Exptl. Biol. Med. 59, 287.
Grazi, E., De Flora, A., and Pontremoli, S. (1960). Biochem. Biophys. Res. Commun.
2, 121.
Green, A.A. and McElroy, W.D. (1955). In "Methods in Enzymology" (S.P. Colowick
and N.O. Kaplan, eds.). Vol. 2, p. 857. Academic Press, New York.
Green, D.E. (1936). Biochem. J. 30, 2095.
Green, D.E. (1960). Ciba Found. Symp., Quinones Electron Transport p. 130.
Green, D.E. and Brosteaux, J. (1936). Biochem. J. 30, 1489.
Green, D.E. and Crane, F.L. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo Kyoto
1957 p. 275. Maruzen, Tokyo.
1014 REFERENCES
Green, D.E. and Fleischer, S. (1960). In "Metabolic Pathways" (D.M. Greenberg, ed.)
Vol. 1, p. 41. Academic Press, New York.
Green, D.E. and Wakil, S.J. (1960). In "Lipide Metabolism" (K. Bloch, ed.), p. 1.
Wiley, New York.
Green, D.E., Dewan, J.G., and Leloir, L.F. (1937 a). Biochem. J. 31, 934.
Green, D.E., Needham, D.M., and Dewan, J.G. (1937 b). Biochem. J. 31, 2327.
Green, D.W., Ingram, V.M., and Perutz, M.F. (1954). Proc. Roy. Soc. A 225, 287.
Green, I. and Mommaerts, W.F.H.M. (1954). J. Biol. Chem. 210, 695.
Green, J.W. and Parpart, A.K. (1953). J. Cellular Camp. Physiol. 42, 191.
Green, N.M. and Neurath, H. (1953). J. Biol. Chem. 204, 379.
Greenawalt, J.W., WeibuU, C., and Low, H. (1962). J. Biol. Chem. 237, 853.
Greenbaum, L.M. and Sherman, R. (1962). J. Biol. Chem. 237, 1082.
Greenberg, D.M. and Winnick, T. (1940). J. Biol. Chem. 135, 761.
Greenberg, D.M., Bagot, A.E., and Roholt, D.A., Jr. (1956). Arch. Biochem. Biophys.
62, 446.
Greengard, O. and Feigelson, P. (1962). J. Biol. Chem. 237, 1903.
Greengard, P. and Straub, R.W. (1962). J. Physiol. (London) 161, 414.
Greenstein, J.P. (1938). J. Biol. Chem. 125, 501.
Greenstein, J.P. and Edsall, J.T. (1940). J. Biol. Chem. 133, 397.
Greenstein, J.P. and Jenrette, W.V. (1942). J. Biol. Chem. 142, 175.
Greenstein, J.P., Thompson, J.W., and Jenrette, W.V. (1940). J. Natl. Cancer Inst.
1, 367.
Gregg, D.C. and Nelson, J.M. (1940). .7. Am. Chem. Soc. 62, 2506.
Gregg, J.R. and Ornstein, N. (1953). Biol. Bull. 105, 466.
Gregolin, C. and Singer, T.P. (1963). Biochim. Biophys. Acta 67, 201.
Greif, R.L. (1960). Federation Proc. 19, 362.
Greif, R.L. and Jacobs, G.S. (1958). Am. J. Physiol. 192, 599.
Greif, R.L., Sullivan, W^J., Jacobs, G.S., and Pitts, R.F. (1956). J. Clin. Invest. 35,
38.
Grein, L. and Pfleiderer, G. (1958). Biochem. Z. 330, 433.
Gremels, H. (1929). Arch. Exptl. Pathol. Pharmakol. 140, 205.
Greville, G.D. (1936). Biochem. J. 30, 877.
Greville, G.D. and Mildvan, A.S. (1962). Biochem. J. 84, 21p.
Greville, G.D. and Needham, D.M. (1955). Biochim. Biophys. Acta 16, 284.
Greville, G.D. and Tapley, D.F. (1960). Biochem. J. 76, 49p.
Grey, E.G. (1924). Proc. Roy. Soc. B96, 156.
Griffith, G.C., Butt, E.M., and Walker, J. (1954). Ann. Internal Med. 41, 501.
Griffith, W.H. (1937). Proc. Soc. Exptl. Biol. Med. 37, 279.
Griffiths, D.E. and Chaplain, R.A. (1962). Biochem. Biophys. Res. Commun. 8, 497.
Griffiths, D.E., Morrison, J.F., and Ennor, A.H. (1957). Biochem. J. 65, 153.
Grisolia, S., Caravaca, J., and Joyce, B.K. (1958). Biochim. Biophys. Acta 29, 432.
Gromet-Elhanen, Z. and Avron, M. (1963). Biochem. Biophys. Res. Commun. 10,
215.
Gros, F. and Naono, S. (1961). I7i "Protein Biosynthesis" (R.J.C. Harris, ed.), p. 195.
Academic Press, New York.
Gross, R.E. (1921). Z. Physiol. Chem. 112, 236.
Grossman, L. and Kaplan, N.O. (1958 a). J. Biol. Chem. 231, 717.
Grossman, L. and Kaplan, N.O. (1958 b). J. Biol. Chem. 231, 727.
REFERENCES 1015
Grossowicz, N. and Halpem, Y.S. (1956 a). Nature 177, 623.
Grossowicz, N. and Halpern, Y.S. (1956 b). Biochim. Biophys. Acta 20, 577.
Groupe, V., Engle, C.G., GafFney, P.E., and Manaker, R.A. (1955). Appl. Microbiol.
3, 333.
Gruber, CM., de Berardinis, C, and Erf, L.A. (1949). Arch. Intern. Pharmacodyn.
79, 461.
Gruber, W., Warzecha, K., Pfleiderer, G., and Wieland, T. (1962). Biochem. Z. 336,
107.
Gruber, W., Hohl, R., and Wieland, T. (1963). Biochem. Biophys. Res. Commun. 12,
242.
Grunert, R.R. (1960). Arch. Biochem.. Biophys. 86, 190.
Grunert, R.R. and Rohdenburg, E.L. (1960). Arch. Biochem. Biophys. 86, 185.
Guarino, A.J. and Sable, H.Z. (1956). Biochim. Biophys. Acta 20, 201.
Gubler, C.J. (1958). Federation Proc. 17, 477.
Gubler, C.J. (1961). J. Biol. Chem. 236, 3112.
Guest, J.R. (1960). Biochem. J. 76, 405.
Guggenheim, K. and Diamant, E.J. (1958). Biochem. J. 69, 56.
Guha, S.R. (1962). Enzymologia lA, 310.
Gulland, J.M. and Farrar, W.V. (1944). Nature 154, 88.
Gumbmann, M. and Tappel, A.L. (1962 a). Arch. Biochem. Biophys. 98, 262.
Gumbmann, M. and Tappel, A.L. (1962 b). Arch. Biochem. Biophys. 98, 502.
Gunja, Z.H., Manners, D.J., and Maung, K. (1961). Biochem. J. 81, 392.
Gurd, F.R.N, and Wilcox, P.E. (1956). Advan. Protein Chem. 11, 311.
Gurtner, H.P. (1961). Helv. Physiol. Pharmacol. Acta Suppl. 9.
Haarniann, W. (1943 a). Biochem. Z. 314, 1.
Haarmann, W. (1943 b). Biochem. Z. 314, 18.
Haas, E. (1944). J. Biol. Chem. 155, 321.
Haas, H.T.A. (1941). Arch. Exptl. Pathol. Pharmakol 197, 284.
Haas, W.J., Sizer, I.W., and Loofbourow, J.R. (1951). Biochim. Biophys. Acta 6,
601.
Habeeb, A.F.S.A. (1960). Can. J. Biochem. Physiol. 38, 269.
Hachisuka, Y., Kato, H., Asano, N., and Kuno, T. (1955). J. Bacterial. 69, 407.
Hackett, D.P. (1958). Plant Physiol. 33, 8.
Hackett, D.P., Rice, B., and Schmid, C. (1960). J. Biol. Chem. 235, 2140.
Hackney, F.M.V. (1948). Proc. Linnean Soc. N. S. Wales 73, 439.
Haft, D.E. and Mirsky, I.A. (1952). J. Pharmacol. Exptl. Therap. 104, 340.
Hagen, U. (1955). Arch. Exptl. Pathol. Pharmakol. 124, 193.
Hagihira, H., Lin, E.C.C., Samiy, A.H., and Wilson, T.H. (1961). Biochem. Biophys.
Res. Commun. 4, 478.
Hagihira, H., Wilson, T.H., and Lin, E.C.C. (1963). Biochim. Biophys. Acta 78, 505.
Hagstrom, B.E. (1963). Biol. Bull. 124, 55.
Hahn, H. and Taeger, H. (1931). Klin. Wochschr. 10, 489.
Hahn, L. and Fekete, J. (1953). Acta Chem. Scand. 7, 798.
Hahn, L. and Frank, E. (1953). Acta Chem. Scand. 7, 806.
Hahn, M. and Remy, E. (1922). Deut. Med. Woschschr. 48, 793.
Haining, J.L., Fukui, T., and Axelrod, B. (1960). J. Biol. Chem. 235, 160.
Hakala, M.T., Glaid, A.J., and Schwert, G.W\ (1953). Federation Proc. 12, 213.
1016 REFERENCES
Halasz, P., Mechler, F., Feher, 0., and Damjanovich, S. (1960). Acta Physiol. Acad.
Sci. Hung. 18, 47.
Halenz, D.R., Feng, J.-Y., Hegre, C.S., and Lane, M.D. (1962). J. Biol. Chem. 237,
2140.
Haley, T.J. (1945). J. Am. Pharm. Assoc, Sci. Ed. 36, 229.
HaU, A.N., Lea, D.J., and Rydon, H.N. (1962). Biochem. J. 84, 12.
Hall, G.A., Jr. (1949). J. Am. Chem. Soc. 71, 2691.
Halliday, S.L., Sloboda, A., Will, L.W., and Oleson, J.J. (1957). Federation Proc. 16,
190.
Hallman, N. (1940). Acta Physiol. Scand. 1, Suppl. 4.
Halvorson, H. and Ellias, L. (1958). Biochim. Biophys. Acta 30, 28.
Halvorson, H., Spiegelman, S., and Hinman, R.L. (1955). Arch. Biochem. Biophys.
55, 512.
Halvorson, H. and Spiegelman, S. (1952). J. Bacteriol. 64, 207.
Hamilton-Miller, J.M.T. and Smith, J.T. (1964). Nature 201, 999.
Hamilton-Miller, J.M.T., Smith, J.T., and Knox, R. (1964). Nature 201, 867.
Hampton, A. (1962). Federation Proc. 21, 370.
Hampton, A. (1963). J. Biol. Chem. 238, 3068.
Hand, W.C, Edwards, B.B., and Caley, E.R. (1943). J. Lab. Clin. Med. 28, 1835.
Handler, P. (1945). J. Biol. Chem. 161, 53.
Handler, P. and Klein, J.R. (1942). J. Biol. Chem. 143, 49.
Handley, C.A. and Lavik, P.S. (1950). J. Pharmacol. Exptl. Therap. 100, 115.
Handley, C.A. and Seibert, R.A. (1956). J. Pharmacol. Exptl. Therap. 117, 253.
Handschumacher, R.E. (1960). J. Biol. Chem. 235, 2917.
Handschumacher, R.E. and Pasternak, C.A. (1958). Biochim,. Biophys. Acta 30, 451.
Hanly, V.F., Rowan, K.S., and Turner, J.S. (1952). Australian J. Sci. Res. Ser. B5, 64.
Hannoun, C. (1952). Compt. Rend. 235, 1445.
Hanson, A. (1958). Naturwissenschaften 45, 423.
Hanzlik, P.J. (1923 a). J. Ind. Hyg. 4, 386.
Hanzlik, P.J. (1923 b). J. Ind. Hyg. 4, 448.
Hanzlik, P.J. (1923 c). J. Pharmacol. Exptl. Therap. 20, 463.
Happold, F.C. (1950). Advan. Enzymol. 10, 51.
Harary, I. (1957). Federation Proc. 16, 192.
Hargreaves, A.B. (1955). Arch. Biochem. Biophys. 57, 41.
Harley, J.L. and Ap Rees, T. (1959). New Phytologist 58, 364.
Harper, E. and Strecker, H.J. (1962). J. Neurochem. 9, 125.
Harpur, R.P. and Quastel, J.H. (1949). Nature 164, 693.
Harris, J. and Hellerman, L. (1956). In "Inorganic Nitrogen Metabolism" (W.D.
McElroy and H.B. Glass, eds.), p. 565. Johns Hopkins Press, Baltimore, Maryland.
Harris, J.E. (1957). Am.. J. Ophthalmol. 44, Suppl. p. 262.
Harris, J.O., Eisenstark, A., and Dragsdorf, R.D. (1954). J. Bacteriol. 68, 745.
Harting, J. (1947). Biol. Bull. 93, 194.
Hartman, S.C. (1963 a). J. Biol. Chem. 238, 3024.
Hartman, S.C. (1963 b). J. Biol. Chem. 238, 3036.
Hartman, W.J., Akawie, R.L, and Clark, W.G. (1955). J. Biol. Chem. 216, 507.
Hartmann, K.-U. and Heidelberger, C. (1961). J. Biol. Chem. 236, 3006.
Hartshorne, D. and Greenberg, D.M. (1964). Arch. Biochem.. Biophys. 105, 173.
Harvey, G.T. and Beck, S.D. (1953). J. Biol. Chem. 201, 765.
REFERENCES 1017
Harvey, S.C. (1949). J. Biol. Chem. 179, 435.
Hashimoto, T., Nakao, M., and Yoshikawa, H. (1960). J. Biochem. (Tokyo) 48, 158.
Haskins, F.A., Tissieres, A., Mitchell, H.K., and Mitchell, M.B. (1953). J. Biol. Chem.
200, 819.
Haslam, R.J. and Krebs, H.A. (1963). Biochem. J. 88, 566.
Hass, L.F. and Byrne, W.L. (1960). J. Am. Chem. Soc. 82, 947.
HassaU, K.A. (1958). Nature 181, 1273.
Hasse, K. and Schleyer, H. (1961). Biochem. Z. 334, 360.
Hasselbach, W. (1953). Z. Naturforsch. 8, 212.
Hatch, M.D. and Stumpf, P.K. (1961). J. Biol. Chem. 236, 2879.
Hatch, M.D. and Turner, J.F. (1960). Biochem. J. 76, 556.
Hatch, M.D., Pearson, J.A., Millerd, A., and Robertson, R.N. (1959). Australian J.
Biol. Sci. 12, 167.
Hatefi, Y., Haavik, A.G., and Griffiths, D.E. (1961a). Biochem. Biophys. Res. Commun.
4, 441.
Hatefi, Y., Haavik, A.G., and Jurtshuk, P. (1961b). Biochim. Biophys. Acta 52, 106.
Haugaard, N. (1946). J. Biol. Chem. 164, 265.
Haugaard, N. and Marsh, J.B. (1952). J. Biol. Chem. 194, 33.
Hauge, J.G. (1956). J. Am. Chem. Soc. 78, 5266.
Hauge, J.G. (1960). Biochim. Biophys. Acta 45, 250.
Haughton, B.G. and King, H.K. (1958). Biochem. J. 70, 660.
Haughton, B.G. and King, H.K. (1961). Biochem. J. 80, 268.
Haiirowitz, F. and Tekman, S. (1947). Biochim. Biophys. Acta 1, 484.
Hawtrey, A.O. (1962). Biochem. J. 85, 293.
Hayaishi, 0. (1953). J. Am. Chem. Soc. 75, 4367.
Hayaishi, O. (1954). Federation Proc. 13, 226.
Hayaishi, O. (1955 a). J. Biol. Chem. 215, 125.
Hayaishi, O. (1955 b). In "Amino Acid Metabolism" (W.D. McElroy and H.B. Glass,
eds.), p. 914. Johns Hopkins Press, Naltimore, Maryland.
Hayaishi, O. and Kornberg, A. (1952). J. Biol. Chem. 197, 717.
Hayaishi, 0., Katagiri, M., and Rothberg, S. (1957). J. Biol. Chem. 229, 905.
Heald, P.J. (1953). Biochem. J. 55, 625.
Heilbrunn, L.V. (1925). Biol. Bull. 49, 241.
Heilbrunn, L.V. and Wilson, W.L. (1955). J. Cellular Comp. Physiol. 46, 361.
Heimann-Hollaender, E. and Lichtenstein, N. (1954). Nature 174, 559.
Heinz, F., Bertelsen, K., and Lamprecht, W. (1962). Z. Physiol. Cheyn. 329, 222.
Heinz, R. (1899). Arch. Pathol. Anat. Physiol. 155, 44.
Hellerman, L. (1937). Physiol. Rev. 17, 454.
Hellerman, L. (1939). Cold Spring Harbor Symp. Quant. Biol. 7, 165.
Hellerman, L. and Chinard, F.P. (1955). In "Methods of Biochemical Analysis" (D.
GUck, ed.). Vol. 1, p. 1. Wiley (Interscience), New York.
Hellerman, L. and Newman, M.D. (1932). J. Am. Chem. Soc. 54, 2859.
Hellerman, L. and Perkins, M.E. (1934). J. Biol. Chem. 107, 241.
Hellerman, L., Chinard, F.P., and Ramsdell, P.A. (1941). J. Am. Chem. Soc. 63,
2551.
Hellerman, L., Chinard, F.P., and Deitz, V.R. (1943). J. Biol. Chem. 147, 443.
Hellerman, L., Lindsay, A., and Bovarnik, M.R. (1946). J. Biol. Chem. 163, 553.
HeUerman, L., Schellenberg, K.A., and Reiss, O.K. (1958). J. Biol. Chem. 233, 1468.
1018 REFERENCES
Hellerman, L., Reiss, O.K., Parmar, S.S., Wein, J., and Lasser, N.L. (1960). J. Biol.
Chem. 235, 2468.
Hellerman, L., Reiss, O.K., and Gey, M.K. (1962). J. Biol. Chem. lil, 997.
Hellig, H. and Popjak, G. (1961). J. Lipid Res. 2, 235.
Helmreich, E. and Eisen, H.N. (1959). J. Biol. Chem. 234, 1958.
Hemker, H.C. and Hiilsmann, W.C. (1960). Biochim. Biophys. Acta 44, 175.
Henderson, J.F. (1962). J. Biol. Chem. 237, 2631.
Henderson, J.H.M. and Stauffer, J.F. (1944). Am. J. Botany 31, 528.
Hendley, D.D. and Beers, R.F., Jr. (1959). Federation Proc. 18, 245.
Hendley, D.D. and Beers, R.F., Jr. (1961). J. Biol. Chem. 236, 2050.
Hendlin, D. and Soars, M.H. (1951). Federation Proc. 10, 384.
Hendlin, D., Koditschek, L.K., and Soars, M.H. (1953). J. Bacterial. 65, 466.
Hepler, O.E. and Simonds, J.P. (1946). Arch. Pathol. 41, 42.
Hepler, O.E., Gurley, H., and Simonds, J.P. (1945). Arch. Pathol. 39, 133.
Hepp, P. (1887). Arch. Exptl. Pathol. Pharmakol. 23, 91.
Heppel, L.A., Harkness, D.R., and Hilmoe, R.J. (1962). J. Biol. Chem. 237, 841.
Herbert, D., Gordon, H., Subrahmanyan, V., and Green, D.E. (1940). Biochem. J.
34, 1108.
Herbert, E. (1956). J. Cellular Comp. Physiol. 47, 11.
Heredia, C.F. and Medina, A. (1960). Biochem. J. 77, 24.
Herken, H. and Neuhoff, V. (1963). Z. Physiol. Chem. 331, 85.
Herner, B. (1944). Acta Physiol. Scand. 8, Suppl. 23.
Herring, P.T. and Hynd, A. (1928). J. Physiol. (London) 66, 267.
Herriott, R.M. (1937). J. Gen. Physiol. 20, 335.
Herriott, R.M. (1947). J. Gen. Physiol. 31, 19.
Herz, R., Jr., Tapley, D.F., and Ross, J.E. (1961). Biochim. Biophys. Acta 53, 273.
Herzog, R.O. and Betzel, R. (1911). Z. Physiol. Chem. 74, 221.
Hess, R. and Pearse, A.G.E. (1963). Biochim. Biophys. Acta 71, 285.
Hess, S.M., Connamacher, R.H., Ozaki, M., and Udenfriend, S. (1961). J. Pharmacol.
Exptl. Therap. 134, 129.
Hestrin, S. and Avigad, G. (1958). Biochem. J. 69, 388.
Heydeman, M.T. and Azoulay, E. (1963). Biochim. Biophys. Acta 77, 545.
Heymann, H., Rogach, Z., and Mayer, R.L. (1954). J. Am. Chem. Soc. 76, 6330.
Heymann, H., Gulick, Z.R., De Boer, C.J., De Stevens, G., and Mayer, R.L. (1958).
Arch. Biochem. Biophys. 73, 366.
Heymann, H., Ginsberg, T., Gulick, Z.R., and Mayer, R.L. (1959). Proc. Soc. Exptl.
Biol. Med. 100, 279.
Heymans, J.F. (1889). Arch. Phijsiol. 13, 168.
Hiai, S., Mori, T., Hino, S., and Mori, T. (1957). J. Biochem. {Tokyo) 44, 839.
Hiatt, A.J. (1961). Plant Physiol. 36, 552.
Hiatt, H.H., Marks, P.A., and Shorr, E. (1953). J. Biol. Chem. 204, 187.
Hicks, S.P. (1953). Arch. Pathol. 55. 302.
Hicks, S.P. (1955). Am. J. Pathol. 31, 189.
Hietanen, S. and Sillen, L.G. (1952). Acta Chem. Scand. 6, 747.
Higgins, E.S. (1961). Enzymolocjia 23, 176.
Higgins, E.S., Richert, D.A., and Westerfeld, W.W. (1956a). Federation Proc. 15, 274.
Higgins, E.S., Richert, D.A., and Westerfeld, W.W. (1956b). Proc. Soc. Exptl. Biol.
Med. 92, 509.
REFERENCES 1019
Hildebrandt, A.C., Riker, A. J., and Watertor, J.L. (1954). Phytopathology 44, 422.
Hill, B.R. (1956). Cancer Res. 16, 460.
Hill, R.L. and Smith, E.L. (1957). J. Biol. Chern. 224, 209.
Hilton, J.L. (1958). Science 128, 1509.
Hilz, H. and Klempien, E.J. (1959). Biochem. Z. 331, 563.
Hirade, J. (1952). J. Biochem. (Tokyo) 39, 185.
Hirade, J. and Hayashi, T. (1953). J. Biochem. (Tokyo) 40, 461.
Hirashinia, K. (1934). Folia Pharmacol. Japon. 19, 219.
Hirashima, K. (1935). Folia Pharmacol. Japon. 19, 333.
Hird, F.J.R. and Yates, J.R. (1961). Biochem. J. 80, 612.
Hitchcock, P. (1946). J. Pharmacol. Exptl. Therap. 87, 55.
Hitchings, G.H. (1960). Clin. Pharmacol. Therap. 1, 570.
Hoadley, L. (1930). Biol. Bull. 58, 123.
Hoare, J.L. and Speakman, J.B. (1963). Mature 199, 1091.
Hoberman, H.D. and Rittenberg, D. (1943). J. Biol. Chem. 147, 211.
Hoch, F.L. and Vallee, B.L. (1959). In ''Sulfur in Proteins" (R. Benesch et al., eds.)
p. 245. Academic Press, New York.
Hoch, F.L. and Vallee, B.L. (1960). Arch. Biochem. Biophys. 91, 1.
Hoch, G.E., Schneider, K.C., and Burris, R.H. (1960). Biochim. Biophys. Acta 37, 273.
Hochachka, P.W. and Teal, J.M. (1964). Biol. Bull. 126, 274.
Hochster, R.M. and Chang, V.M. (1963). Can. J. Biochem. Physiol. 41, 1503.
Hochster, R.M. and Nozzolillo, C.G. (1960). Can. J. Biochem. Physiol. 38, 79.
Hochster, R.M. and Quastel, J.H. (1952). Arch. Biochem. Biophys. 36, 132.
Hochster, R.M. and Quastel, J.H. (eds.) (1963). '"Metabolic Inhibitors," Academic
Press, New York. Vol. I.
Hockenhull, D.J.D., Fantes, K.H., Herbert, M., and Whitehead, B.K. (1954a). J.
Gen. Microbiol. 10, 353.
Hockenhull, D.J.D., Herbert, M., Walker, A.D., Wilkin, G.D., and Winder, F.G.
(1954 b). Biochem. J. 56, 73.
Hockenhull, D.J.D., A.shton, G.C., Fantes, K.H., and Whitehead, B.K. (1954 c). Bio-
chim. J. 57, 93.
Hofling, E., Lieb, H., and Schoniger, W. (1952). Monatsh. Chem. 83, 60.
Hofmann, E., Konig, S., and Behne, I. (1962). Naturwissenschaften 49, 40.
Hofmann, E.C.G. (1955). Biochem. Z. 327, 273.
Hofmann, E.C.G. and Rapoport, S. (1957). Biochem. Z. 329, 437.
Hofstee, B.H.J. (1949). J. Biol. Chem. 179. 633.
Hofstee, B.H.J. (1955). J. Biol. Chern. 216, 235.
Hofstee, B.H.J. (1960). J. Am. Chem. Soc. 82, 5166.
Hofstee, B.H.J. (1961). Biochem. Biophys. Res. Commun. 4, 5.
Hokfelt, B. and Bydgeman, S. (1961). Proc. Soc. Exptl. Biol. Med. 106, 537.
Hokfelt, B. and Hultquist, G. (1961). Proc. Soc. Exptl. Biol. Med. 107, 66.
Holden, M. and Pirie, N.W. (1955). Biochem. J. 60, 53.
Holland, J. and Humphrey, G.F. (1953). Australian J. Exptl. Biol. Med. Sci. 31, 299.
Holland, J.F. (1961). Clin. PJmrmacol. Therap. 2, 374.
HoUocher, T.C., Jr. and Commoner, B. (1960). Proc. Natl. Acad. Sci. U. S. 46, 416.
Holly, F.W., Peel, E.W., Mozingo, R., and Folkers, K. (1950). J. Am. Chem. Soc. 72,
5416.
Holmberg, C.G. (1939). Biochem. J. 33, 1901.
1020 REFERENCES
Holmes, W.L. (1956). J. Biol. Chem. 223, 677.
Holt, A. and Wold, F. (1961). J. Biol. Chem. 236, 3227.
Holtkamp, D.E. and Hill, R.M. (1951). Arch. Biochem. Biophys. 34, 216.
Holton, F.A. and Colpa-Boonstra, J. (1960). Biochem. J. 76, 179.
Holtz, P. and Palm, D. (1964). Pharmacol. Rev. 16, 113.
Holzer, H. (1956). Medizinische No. 15, 576.
Holzer, H. and Frank, S. (1958). Angeiv. Chem. 70, 570.
Holzer, H. and Holldorf, A. (1957). Biochem. Z. 329, 292.
Holzer, H. and Schneider, S. (1961). Biochitn. Biophys. Acta 48, 71.
Holzer, H., Holzer, E., and Schultz, G. (1955a). Biochem. Z. 326, 385.
Holzer, H., Haan, J., and Schneider, S. (1955 b). Biochem. Z. 326, 451.
Holzer, H., Haan, J., and Pette, D. (1955 c). Biochem. R. 327, 195.
Holzer, H., Goedde, H.W., and Schneider, S. (1955 d). Biochem. Z. 327, 245.
Hommes, F.A. (1963). Biochim. Biophys. Acta 77, 183.
Honda, S.I. (1955). Plant Physiol. 30, 402.
Honda, S.I. (1957). Plant Physiol. 32, 23.
Honda, S.I. and Muenster, A.-M. (1961). Plant Physiol. 36, 105.
Hopkins, F.G. and Morgan, E.J. (1938). Biochem. J. 32, 611.
Hopkins, F.G., Morgan, E.J., and Lutwak-Mann, C. (1938). Biochem. J. 32, 1829.
Hopper, S. and Segal, H.L. (1964). Arch. Biochem. Biophys. 105, 501.
Hori, K. (1963). J. Biochem. (Tokyo) 53, 354.
Horio, T. and Kamen, M.D. (1962 a). Biochemistry 1, 144.
Horio, T. and Kamen, M.D. (1962 b). Biochemistry 1, 1141.
Horio, T. and Okunuki, K. (1954). J. Biochem. (Tokyo) 41, 717.
Horning, M.G., Martin, D.B., Karmen, A., and Vagelos, P.R. (1960). Biochem. Bio-
phys. Res. Commun. 3, 101.
Horowitz, M.G. and Klotz, I.M. (1956). Arch. Biochem. Biophys. 63, 77.
Horwitt, M.K. (1940). Science 92, 89.
Horwitz, L. (1957). J. Cellular Comp. Physiol. 49, 437.
Hoshino, M. (1959). J. Biochem. (Tokyo) 46, 825.
Hoskins, D.D., Whiteley, H.R., and Mackler, B. (1962). J. Biol. Chem. 237, 2647.
Hosoya, N. and Kawada, N. (1958). J. Biochem. (Tokyo) 45, 363.
Hosoya, N. and Kawada, N. (1961). J. Biochem. (Tyoko) 49, 728.
Hosoya, N., Kawada, N., and Matsumnra, Y. (1960). J. Biochem. (Tokyo) 48, 262.
Hosoya, T. (1960). J. Biochem. (Tokyo) 48, 375.
Hotchkiss, R.D. (1956). Arch. Biochem. Biophys. 65, 302.
Houck, C.R. (1942). J. Cellular Comp. Physiol. 20, 277.
Houck, J.C. (1957 a). Arch. Biochem. Biophys. 71, 336.
Houck, J.C. (1957 b). Biochim. Biophys. Acta 26, 649.
Howe, W.B., Melson, G.L., Meridith, C.H., Morrison, J.R., Piatt, M.H., and Stro-
minger, J.L. (1964). J. Pharmacol. Exptl. Therap. 143, 282.
Huang, H.T. and Niemann, C. (1952). J. Am. Chem. Soc. 74, 101.
Hubbard, J.S. and Durham, N.N. (1961). J. Bacteriol. 82, 361.
Hudson, M.T. and Woodward, G.E. (1958). Biochim. Biophys. Acta 28, 127.
Huennekens, F.M., Basford, R.E., and Gabrio, B.W. (1955). J. Biol. Chem. 213, 951.
Huennekens, F.M., Caifrey, R.W., Basford, R.E., and Gabrio, B.W. (1^51 a,). J. Biol.
Chem. in, 261.
Huennekens, F.M., Felton, S.P., and Snell, E.E. (1957 b). J. Am. Chem. Soc. 79, 2258.
REFERENCES 1021
Huffaker, R.C. and Wallace, A. (1961). Biochim. Biophys. Acta 46, 403.
Hughes, A.F.W. (1950). Quart. J. Microscop. Sci. 91, 251.
Hughes, C. and Spragg, S.P. (1958). Biochem. J. 70, 205.
Hughes, D.E. and Williamson, D.H. (1952). Biochem. J. 51, 45.
Hughes, W.J., Jr. (1947). J. Am. Ckem. Soc. 69, 1836.
Hughes, W.L., Jr. (1950). Cold Spring Harbor Symp. Quant. Biol. 14, 79.
Hughes, W.L., Jr. (1957). Ann. N. Y. Acad. Sci. 65, 454.
Hughes, W.L., Jr. and Dintzis, H.M. (1964). J. Biol. Ckem. 239, 845.
Hughes, W.L., Jr. and Straessle, R. (1950). J. Am. Chem. Soc. 72, 452.
Huisman, T.H.J. (1959). In "Sulfur in Proteins" (R. Benesch et al., eds.), p. 153.
Academic Press, New York.
Hiilsmann, W.C. (1961). Nature 192, 1153.
Hummel, J.P., Anderson, D.O., and Patel, C. (1958). J. Biol. Chem. 233, 712.
Humphrey, B.A. and Humphrey, G.F. (1948). J. Exptl. Biol. 25, 123.
Humphrey, G.F. (1947). J. Exptl. Biol. 24, 352.
Humphrey. G.F. (1950). Australian J. Exptl. Biol. Med. Sci. 28, 1.
Humphrey, G.F. (1957). Biochem. J. 65, 546.
Humphreys, T.E., Newcomb, E.H., Bokman, A.H., and Stumpf, P.K. (1954). J. Biol.
Chem. 210, 941.
Hunter, A. and Downs, C.E. (1945). J. Biol. Chem. 157, 427.
Hunter, D., Bomford, R.R., and Russell, D.S. (1940). Quart. J. Med. 9, 193.
Hunter, F.E., Jr. and Ford, L. (1955). J. Biol. Chem. 216, 357.
Hunter, F.E., Jr., Fink, J., and Hurwitz, A. (19.59 a). Federation Proc. 18, 251.
Hunter, F.E., Jr., Levy, J.F., Fink, J., Schutz, B., Guerra, F., and Hurwitz, A. (1959b).
J. Biol. Chem. 234, 2176.
Hunter, F.R. (1960). Exptl. Parasitol. 9, 271.
Hunter, G.J.E. (1953). Biochem. J. 55, 320.
Hunter, N.W. (1958). Physiol. Zool. 31, 23.
Hunter, N.W. and Hunter, R. (1957). Science 126, 1246.
Hunter, W.C. (1929). Ann. Internal Med. 2, 796.
Hunter, W.R. (1949). J. Exptl. 'Biol. 26, 113.
Hurwitz, J. (1952). Biochim. Biophys. Acta 9, 496.
Hurwitz, J. (1953). J. Biol. Chem. 205, 935.
Hurwitz, J. (1955 a). J. Biol. Chem. 212, 757.
Hurwitz, J. (1955 b). J. Biol. Chem. 217, 513.
Hurwitz, J., Weissbach, A., Horecker, B.L., and Smyrniotis, P.Z. (1956). J. Biol.
Chem. 218, 769.
Hurwitz, J., Heppel, L.A., and Horecker, B.L. (1957). J. Biol. Chem. 226, 525.
Hurwitz, L. and Chaffee, E. (1954). Federation Proc. 13, 369.
Huszak, S. (1935). Z. Physiol. Chem. 236, 66.
Huszak, S. (1940). Biochem. Z. 303, 349.
Huxley, J.S. (11928). Am. Naturalist 62, 363,
Hylin, J.W. ahd Matsumoto, H. (1964). Toxicol. Appl. Pharmacol. 6, 168.
Hynd, A. (1927). Proc. Roy. Soc. B101, 244.
Ibsen, K.H.,
oe, E.L., and McKee, R.W. (1958). Biochim. Biophys. Acta 30, 384.
Ibsen, K.H., Coe, E.L., and McKee, R.W. (1962). Cancer Res. 22, 182.
Ichihara, A., I
hihara, E.A., and Suda, M. (1960). J. Biochem. (Tokyo) 48, 412.
1022 REFERENCES
Ichihara, K., Yashimatsu, H., and Sakamoto, Y. (1956). J. Biochem. (Tokyo) 43, 803.
Igarasi, H. (1939). J. Agr. Chem. Sac. Japan 15, 225.
Iliffe, J. and Myant, N.B. (1964). Biochem. J. 91, 369.
Illingworth, B., Jansz, H.S., Brown, D.H., and Cori, C.F. (1958). Proc. Natl. Acad.
Sci. U.S. 44, 1180.
Imamoto, F., Iwasa, K., and Okunuki, K. (1959). J. Biochem. {Tokyo) 46, 141.
Imshenetsky, A. A. and Perova, K.Z. (1957). Microbiology {USSR) {English Transl.)
26, 301.
Inagaki, T. (1940). J. Biochem. {Tokyo) 32, 57.
Ingrahara, R.C. and Visscher, M.B. (1936). Am. J. Physiol. 114, 681.
Ingram, V.M. (1955). Biochem. J. 59, 653.
Irvin, J.L. and Irvin, E.M. (1954). J. Biol. Chem. 210, 45.
Isaka, S. and Aikawa, T. (1963). Exptl. Cell Res. 30, 139.
Ishikawa, S. and Lehninger, A.L. (1962). J. Biol. Chem.. 237, 2401.
Ishimoto, M. and Fujimoto, D. (1961). J. Biochem. {Tokyo) 50, 299.
Ishimoto, M. and Yagi, T. (1961). J. Biochem. {Tokyo) 49, 103.
Ishimoto, M., Kondo, Y., Kameyama, T., Yagi, T., and Shiraki, M. (1958). Proc. In-
tern. Syynp. Enzyme Chem., Tokyo Kyoto 1957 p. 229. Maruzen, Tokyo.
Isles, T.E. and Jocelyn, P.C. (1963). Biochem. J. 88, 84.
Ito, E., Kondo, S., and Watanabe, S. (1955). J. Biochem. {Tokyo) 42, 793.
Ives, D.H., Morse, F.A., Jr., and Potter, V.R. (1963). J. Biol. Chem. 238, 1467.
Iwatsubo, M. and Labeyrie, F. (1962). Biochim. Biophys. Acta 59, 614.
Iwatsuka, H., Kuno, M., and Maruyama, M. (1962). Plant Cell Physiol. {Tokyo) 3, 157.
Izar, G. (1909). Biochem. Z. 11, 371.
Jaattela, A.J. and Paasonen, M.K. (1961). Acta Pharmacol. Toxicol. 18, 95.
Jackson, D.E. (1926 a). J. Pharmacol. Exptl. Therap. 27, 244.
Jackson, D.E. (1926 b). J. Pharmacol. Exptl. Therap. 29, 471.
Jackson, P.C., Hendricks, S.B., and Vasta, B.M. (1962). Plant Physiol. 37, 8.
Jacob, H.S. and Jandl, J.H. (1962). J. Clin. Invest. 41, 779.
Jacobs, E.E., Jacob, M., Sanadi, D.R., and Bradley, L.B. (1956). J. Biol. Chem. 113,
147.
Jacobs, F.A. (1958). Federation Proc. 17, 79.
Jacobs, F.A. and Hillman, R.S.L. (1958). J. Biol. Chem. 232, 445.
Jacobs, F.A., Coen, L.J., and Hillman, R.S.L. (1960). J. Biol. Chem. 235, 1372.
Jacobs, H.I. (1950). Ph.D. thesis. University of Southern California.
Jacobsohn, K. (1953). Enzymologia 16, 113.
Jacobson, K.B. (1959). J. Gen. Physiol. 43, 323.
Jacobson, K.B. (1960). J. Biol. Chem. 235, 2713.
Jacobson, K.B. and Kaplan, N.O. (1957 a). J. Biophys. Biochem. Cytol. 3, 31.
Jacobson, K.B. and Kaplan, N.O. (1957 b). J. Biol. Chem. lib, 427.
Jacoby, G.A. and La Du, B.N. (1964). J. Biol. Chem. 239, 419.
Jacoby, M. (1933). Biochem. Z. 161, 181.
Jacquez, J.A. (1961). Am. J. Physiol. 200, 1063.
Jaenicke, L. and Brode, E. (1961). Biochem. Z. 334, 342.
Jagendorf, A.T. and Avron, M. (1959). Arch. Biochem. Biophys. 80, 246.
Jahn, F. (1914). Arch. Exptl. Pathol. Pharmakol. 76, 16.
Jakoby, W.B. (1954). J. Biol. Chem. 207, 657.
REFERENCES 1023
Jakoby, W.B. and Bonner, D.M. (1953). J. Biol. Chem. 205, 709.
James, W.O. and Beavers, H. (1950). New Phijtologist 49, 353.
Janacek, K. (1962). Biochim. Biophys. Acta 56, 42.
Jansen, E.F., Nutting, M.-D.F., Jang, R., and Balls, A.K. (1950). J. Biol. Chem. 185,
209.
Jansen, E.F., Curl, A.L., and Balls, A.K., (1951). J. Biol. Chem. 189, 671.
Jarnefelt, J. (1962). Biochim. Biophys. Acta 59, 643.
Jecsai, G. and Elodi, P. (1963). Acta Physiol. Acad. Sci. Hung. 24, 29.
Jedeikin, L.A. and Weinhouse, S. (1954). Arch. Biochem. Biophys. 50, 134.
Jeffree, G.M. (1956). Biochem.. J. 64, 49p.
Jeffree, G.M. (1957). Biochim. Biophys. Acta 24, 532.
Jencks, W.P. and Lipmann, F. (1957). J. Biol. Chem. 225, 207.
Jenerick, H.P. (1957). J. Cellular Comp. Physiol. 49, 171.
Jenkins, W.T., Yphantis, D.A., and Sizer, I.W. (1959). J. Biol. Chem. 234, 51.
Jensen, E.V., Hospelhorn, V.D., Tapley, D.F., and Huggins, C. (1950). J. Biol. Chem.
185, 411.
Jermstad, A. and Jensen, K.B. (1950). Pharm. Acta Helv. 25, 209.
Jermstad, A. and Jensen, K.B. (1951). Bull. Soc. Chivi. Biol. 33, 258.
Jervis, E.L. and Smyth, D.H. (1960). J. Physiol. (London) 151, 51.
Joachimoglu, G. (1922). Biochem. Z. 130, 239.
Jodrey, L.H. and Wilbur, K.M. (1955). Biol. Bull. 108, 346.
Johnson, G.T. and Moos, C. ( 1956). J. Cellular Comp. Physiol. 48, 243.
Johnson, M.A. and Frear, D.S. (1963). Phrjtochemistry 2, 75.
Johnson, M.P. and Bonner, J. (1956). Physiol. Plantarum 9, 102.
Johnson, W., Corte, G., and Jasmin, R. (1958). Proc. Soc. Exptl. Biol. Med. 99,
677.
Johnson, W.J. (195.5). Can. J. Biochem. Physiol. 33, 107.
Johnson, W.J. and kcCoU. J.D. (1955). Science 122, 834.
Johnson, W.J. and McColl, J.D. (1956). Federation Proc. 15, 284.
Johnston, R.L. (1941). J. Lab. Clin. Med. 27, 303.
Johnstone, J.H. and Mitchell, I.L.S. (1953). Biochem. J. 55, xvii.
Johnstone, R.M. (1958). Federation Proc. 17, 250.
Joklik, W.K. (1950a). Australian J. Sci. Res. Ser. B3, 28.
Joklik, W.K. (1950 b). Australian J. Exptl. Biol. Med. Sci. 28, 331.
Jolley, R.L., CheldeHn, V.H., and Newburgh, R.W. (1958). J. Biol. Chem. 233, 1289.
Jones, E.A. and Gutfreund. H. (1961). Biochem. J. 79, 608.
Jones, E.A. and Gutfreund, H. (1964). Biochem. J. 91, Ic.
Jones, J.D., Hulme, A.C., and Wooltorton, L.S.C. (1964). Phytochemistry 3, 201.
Jones, J.H., Foster, C, and Henle, W. (1948). Proc. Soc. Exptl. Biol. Med. 69, 454.
Jones, J.R. (1946). J. Exptl. Biol. 23, 298.
Jones, O.T.G. (1963). J. Exj)tl. Botany 14, 399.
Joshi, J.G. and Handler, P. (1962). J. Biol. Chem. 237, 929.
Jowett, M. and Brooks, J. (1928). Biochem. J. 22, 720.
Jowett, M. and Quastel, J.H. (1935 a). Biochem. J. 29, 2143.
Jowett, M. and Quastel, J.H. (1935 b). Biochem. J. 29, 2159.
Jowett, M. and Quastel, J.H. (1935 c). Biochem. J. 29, 2181.
Jowett, M. and Quastel, J.H. (1937). Biochem. J. 31, 275.
Joyce, B.K. and Grisolia, S. (1938). J. Biol. Chem. 233, 350.
1024 REFERENCES
Joyce, C.R.B., Moore, H., and Weatherall, M. (1954). Brit. J. Pharmacol, 9, 463.
Jung, F. (1947). Arch. Exptl. Pathol. Pharmakol. 204, 139.
Kagawa, Y., Minakami, S., and Yoneyama, Y. (1959). J. Biochetn. (Tokyo) 46, 771.
Kahana, S.E., Lowry, O.H., Schulz, D.W., Passonneau, J.V., and Crawford, E.J.
(1960). J. Biol. Chem. 235, 2178.
Kahn, J.S. and Jagendorf, A.T. (1961). J. Biol. Chem. 236, 940.
Kaiser, C. (1960). In "Medicinal Chemistry" (A. Burger, ed.). 2nd ed., p. 89. Wiley
(Interscience), New York.
Kaiser, E. (1953). Nature 171, 607.
Kalckar, H.M., Kjeldgaard, N.O., and Klenow, H. (1948). J. Biol. Chem,. 174, 771.
Kalckar, H.M., Kjeldgaard, N.O., and Klenow, H. (1950). Biochim. Biophys. Acta 5,
586.
Kaldor, G. (1960). Proc. Soc. Exptl. Biol. Med. 105, 645.
Kaldor, G. and Gitlin, J. (1963). Arch. Biochem. Biophys. 102, 216.
Kallen, R.G. and Lowenstein, J.M. (1962). Arch. Biochem. Biophys. 96, 188.
Kalnitsky, G. and Tapley, D.F. (1958). Biochem. J. 70, 28.
Kaltenbach, J.P. and Kalnitsky, G. (1951a). J. Biol. Chem. 192, 629.
Kaltenbach, J.P. and Kalnitsky, G. (1951b). J. Biol. Chem. 192, 641.
Kalyankar, G.D., Krishnaswamy, P.R., and Sreenivasaya, M. (1952). Current Set,
(India) 21, 220.
Kamrin, R.P. and Kamrin, A.A. (1961). J. Neurochem. 6, 219.
Kanamori, M. and Wixom, R.L. (1963). J. Biol. Chem. 238, 998.
Kandler, O. (1955). Z. Naturforsch. 10b, 38.
Kann, E.E. and Mills, R.C. (1955). J. Bacterial. 69, 659.
Kantarjian, A.D. (1961). Neurology 11, 639.
Kaper, J.M. and Houwing, C. (1962 a). Arch. Biochem. Biophys. 96, 125.
Kaper, J.M. and Houwing, C. (1962 b). Arch. Biochem. Biophys. 97, 449.
Kaplan, C. (1959). Nature 184, 1074.
Kaplan, E.H., Kennedy, J., and Davis, J. (1954). Arch. Biochem. Biophys. 51, 47.
Kaplan, N.O. (1959). Ciba Found. Study Group 2, 37.
Kaplan, N.O. and Ciotti, M.M. (1954). J. Biol. Chem. 211, 431.
Kaplan, N.O., Colowick, S.P., and Nason, A. (1951). J. Biol. Chem. 191, 473.
Kaplan, N.O., Goldin, A., Humphreys, S.R., Ciotti, M.M., and Venditti, J.M. (1954).
Science 120, 437.
Kaplan, N.O., Ciotti, M.M., and Stolzenbach, F.E. (1956). J. Biol. Chem. 221, 833.
Kaplan, N.O., Ciotti, M.M., Hamolsky, M., and Bieber, R.E. (1960). Science 131, 392.
Kappner, W. (1961). Protoplasma 53, 504.
Karasek, M.A. and Greenberg, D.M. (1957). J. Biol. Chem. Ill, 191.
Karlsson, J.L. (1950). J. Biol. Chem. 183, .549.
Karnofsky, D.A., Stock, C.C, Ridgway, L.P., and Patterson, P.A. (1950). J. Biol.
Chem. 182, 471.
Karunairatnam, M.C. and Levvy, G.A. (1949). Biochem. J. 44, 599.
Kasamaki, A., Sasaki, S., and Usami, S. (1963). J. Biochem. (Tokyo) 53, 89.
Kashket, E.R. and Brodie, A.F. (1963 a). Biochim. Biophys. Acta 78, 52.
Kashket, E.R. and Brodie, A.F. (1963 b). J. Biol. Chem-. 238, 2564.
Kassanis, B. and Kleczkowski, A. (1944). Biochem. J. 38, 20.
Katchalski, E., Berger, A., and Neumann, H. (1954). Nature 173, 998.
REFERENCES 1025
Kato, I., Matsushima, T., and Akabori, S. (1960). J. Biochem. (Tokyo) 48, 199.
Katoh, S. and Takamiya, A. (1963). Arch. Biochem. Biophys. 102, 189.
Katoh, T. (1963). Plant Cell Physiol. (Tokyo) 4, 13.
Katsuya, H., Kinoshita, T., Doi, Y., Hashizume, T., and Motoyama, H. (1963). Na-
ture 198, 497.
Katyal, J.M. and Gorin, G. (1959). Arch. Biochem. Biophys. 82, 319.
Katz, A.M. (1963). Biochim. Biophys. Acta 71, 397.
Katz, A.M. and Mommaerts, W.F.H.M. (1962). Biochim. Biophys. Acta 65, 82.
Katz, J. and Chaikoff, I.L. (1955). J. Am. Chem. Soc. 77, 2659.
Katz, S. (1952). J. Am. Chem. Soc. 74, 2238.
Katz, S. (1962). Nature 194, 569.
Katz, S. and Santilli, V. (1962 a). Federation Proc. 21, 376.
Katz, S. and Santilli, V. (1962 b). Biochim. Biophys. Acta 55, 621.
Kaufman, B. and Kaplan, N.O. (1961). J. Biol. Chem. 236, 2133.
Kaufman, B.T. (1964). J. Biol. Chem. 239, PC669.
Kaufman, B.T. and Kaplan, N.O. (1960). Biochim. Biophys. Acta 39, 332.
Kaufman, S. and Neurath, H. (1949). Arch. Biochem. 21, 245.
Kavanau, J.L. (1963). Nature 198, 525.
Kawano, Y. and Kawabata, S. (1953). J. Ferment. Technol. (Japan) 31, 184.
Kay, CM. and Marsh, M.M. (1959). Biochim.. Biophys. Acta 35, 262.
Kaziro, Y. (1959). J. Biochem. (Tokyo) 46, 1523.
Kaziro, Y., Hass, L.F., Boyer, P.D., and Ochoa, S. (1962). J. Biol. Chem. 237, 1460.
Kearney, E.B. (1952). J. Biol. Chem. 194, 747.
Kearney, E.B. (1955). In "Methods in Enzymology" (S.P. Colowick and N.O. Kaplan,
eds.). Vol. 2, p. 640. Academic Press, New York.
Kearney, E.B. (1957). J. Biol. Chem. 113, 363.
Kearney, E.B. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo Kyoto 1957 p. 340.
Maruzen, Tokyo.
Kearney, E.B. (1960). J. Biol. Chem. 235, 865.
Kearney, E.B. and Singer, T.P. (1956). J. Biol. Chem. 219, 963.
Keech, D.B. and Utter, M.F. (1963). J. Biol. Chem. 238, 2609.
Keilin, D. and Hartree, E.F. (1936). Proc. Roy. Soc. B119, 114.
Keilin, D. and Hartree, E.F. (1940). Proc. Roy. Soc. B129, 277.
Keilin, D. and King, T.E. (1960). Proc. Roy. Soc. B152, 163.
Keister, D.L. and San Pietro, A. (1963). Arch. Biochem. Biophys. 103, 45.
Keister, D.L., San Pietro, A., and Stolzenbach, F.E. (1960). J. Biol. Chem. 235,
2989.
Keleti, T. and Telegdi, M. (1959). Acta Physiol. Acad. Sci. Hung. 16, 235.
Keller, D.M. and Lotspeich, W.D. (19.59 a). J. Biol. Chem. 234, 987.
Keller, D.M. and Lotspeich, W.D. (1959 b). J. Biol. Chem. 234, 991.
Kellerman, G.M. (1959). Biochim. Biophys. Acta 33, 101.
Kelly, S. (1953). Biol. Bull. 104, 138.
Kenney, F.T. (1959). J. Biol. Chem. 234, 2707.
Kenten, R.H. (1958). Biochem. J. 69, 439.
Keodam, J.C, Steyn-Parve, E.P., and van Rheenen, D.L. (1956). Biochim. Biophys,
Acta 19, 181.
Kerby, G.P. and Eadie, G.S. (1953). Proc. Soc. Exptl. Biol. Med. 83, 111.
Kermack, W.O. and Matheson, N.A. (1957). Biochem. J. 65, 48.
1026 REFERENCES
Kernan, R.P. (1963). J. Physiol. {London) 169, 862.
Kessler, D. and Krause, R.M. (1963). Proc. Soc. Exptl. Biol. Med. 114, 822.
Kessler, R.H. (1960). Clin. Pharmacol. Therap. 1, 723.
Kessler, R.H., Lozano, R., and Pitts, R.F. (1957 a). J. Pharmacol. Exptl. Therap. 121,
432.
Kessler, R.H., Lozano, R., and Pitts, R.F. (1957 b). J. Clin. Invest. 36, 656.
Kessler, R.H., Hierholzer, K., Gurd, R.S., and Pitts, R.F. (1958). Am. J. Physiol.
194, 540.
Keston, A.S. (1963). Arch. Biochem. Biophys. 102, 306.
Kharasch, N. (ed.) (1961). "Organic Sulfur Compounds," Vol. 1. Macmillan (Perga-
mon). New York.
Khomutov, R.M., Karpeiskii, M.Y., Severin, E.S., and Gnuchev, N.V. (1961). Dokl.
Akad. Nauk SSSR 140, 2.
Kielley, W.W. and Bradley, L.B. (1956). J. Biol. Chem. 218, 653.
Kielley, W.W. and Kielley, R.K. (1951). J. Biol. Chem. 191, 485.
Kielley, W.W. and KieUey, R.K. (1953). J. Biol. Chem. 200, 213.
Kiesow, L. (1959). Z. Naturforsch. 14b, 492.
Kiesow, L. (1960 a). Z. Naturforsch. 15b, 174.
Kiesow, L. (1960 b). Z. Naturforsch. 15b, 291.
Kiesow, L. (1960 c). Z. Naturforsch. 15b, 293.
Kiessling, K.-H. (1956). Biochim. Biophys. Acta 20, 293.
Kihara, H. and Snell, E.E. (1957). J. Biol. Chem. 226, 485.
Kilbourne, E.D. (1959). Nature 183, 271.
Kilsheimer, G.S. and Axelrod, B. (1957). J. Biol. Chem. 227, 879.
Kilsheimer, G.S. and Axeb-od, B. (1958). Nature 182, 1733.
Kimmel, J.R. and Smith, E.L. (1954). J. Biol. Chem. 207, 515.
Kimmel, J.R. and Smith, E.L. (1957). Advan. Enzymol. 19, 267.
Kimura, H. and Kummerow, F.A. (1963). Arch. Biochem.. Biophys. 102, 86.
Kimura, T. and Tobari, J. (1963). Biochim. Biophys. Acta 73, 399.
Kimura, Y. and Niwa, T. (1953). Nature 171, 881.
King, T.E. and Cheldelin, V.H. (1954). J. Biol. Chem. 208, 821.
King, T.E. and Cheldelin, V.H. (1956). J. Biol. Chem. 220, 177.
King, T.E. and Howard, R.L. (1962). J. Biol. Chem. 237, 1686.
King, T.E., Kawasaki, E.H., and Cheldelin, V.H. (1956). J. Bacteriol. 72, 418.
Kini, M.M. and Quastel, J.H. (1959). Nature 184, 252.
Kinsky, S.C. and McElroy, W.D. (1958). Arch. Biochem. Biophys. 73, 466.
Kipnis, D.M. (1958). Federation Proc. 17, 254.
Kipnis, D.M. and Cori, C.F. (1959). J. Biol. Chem. 234, 171.
Kipnis, D.M. and Cori, C.F. (1960). J. Biol. Chem. 235, 3070.
Kirkham, D.S. and Flood, A.E. (1963). J. Gen. Microbiol. 32, 123.
Kirkman, H.N. (1962). J. Biol. Chem. 237, 2364.
Kishimoto, U. (1961). Comp. Biochem. Physiol. 2, 81.
Kistiakowsky, G.B. and Shaw, W.H.R. (1953). J. Am. Chem. Soc. 75, 866.
Kistiakowsky, G.B., Mangelsdorf, P.C, Rosenberg, A.J., and Shaw, W.H.R. (1952).
J. Am. Chem. Soc. 74, 5015.
Kistner, S. (1960). Acta Chem. Scand. 14, 1389.
Kit, S. and Greenberg, D.M. (1951). Cancer Pes. 11, 495.
Klein, A.O. and Vishniac, W. (1961). J. Biol. Chem. 236, 2544.
REFERENCES 1027
Klein, J.R. (1940). J. Biol. Chem. 134, 43.
Klein, J.R. (1953). J. Biol. Chem. 205, 725.
Klein, J.R. (1956). Federation Proc. 15, 290.
Klein, J.R. (1957). Arch. Biochem. Biophys. 67, 423.
Klein, J.R. (1960). Biochim. Biophys. Acta 37, 534.
Klein, J.R. and Kamin, H. (1941). J. Biol. Chem. 138, 507.
Klein, J.R. and Olsen, N.S. (1947). J. Biol. Chem. 170, 151.
Klein, M., Brewer, J.H., Perez, J.E., and Day, B. (1948). J. Immunol. 59, 135.
Klein, W. (1957). Z. Physiol. Chem. 307, 254.
Kleinfeld, M., Stein, E., and Murphy, B. (1964). Am. J. Physiol. 206, 975.
Kleinzeller, A. (1961). In "Membrane Transport and Metabolism" (A. Kleinzeller and
A. Kotyk, eds.), p. 527. Academic Press, New York.
Kleinzeller, A., Kotyk, A., and Kovac, L. (1959). Nature 183, 1402.
Klemperer, H.G. (1955). Biochem. J. 60, 122.
Klemperer, H.G. (1957). Biochem. J. 67, 381.
Klimek, J.W., Cavallito, C.J., and Bailey, J.H. (1948). J. Bacteriol. 55, 139.
Kline, D.L. and DeLuca, H.A. (1956). Can. J. Biochem. Physiol. 34, 429.
Klingenberg, M. and Biicher, T. (1961). Biochem. Z. 334, 1.
Klingenberg, M. and Schollmeyer, P. (1960). Biochem. Z. 333, 335.
Klingman, J.D. and Handler, P. (1958). J. Biol. Chem. I'il, 369.
Klotz, I.M. and Klotz, T.A. (1959). In "Sulfur in Proteins" (R. Benesch e< aZ., eds.),
p. 127. Academic Press, New York.
Klotz, I.M. and Tietze, F. (1947). J. Biol. Chem. 168, 399.
Klotz, I.M., Ayers, J., Ho, J.Y.C., Horowitz, M.G., and Heiney, R.E. (1958). J. Am.
Chem. Soc. 80, 2132.
Knopfmacher, H.P. and Salle, A.J. (1941). J. Oen. Physiol. 24, 377.
Kobayashi, Y. (1957). Arch. Biochem. Biophys. 71, 352.
Koch, R. (1881). Mitt. Gesundh. 1, 234.
Kocholaty, W. and Krejci, L.E. (1948). Arch. Biochem. 18, 1.
Kocourek, J., Ticha, M., Jiracek, V., and Kostif, J. (1963). Biochim. Biophys. Acta
71, 497.
Koedam, J.C. (1958). Biochim. Biophys. Acta 29, 333.
Koedam, J.C, Steyn-Parve, E.P., and van Rheenen, D.L. (1956). Biochim. Biophys.
Acta 19, 181.
Koeppe, O.J., Boyer, P.D., and Stulberg, M.P. (19.56). J. Biol. Chem. 219, 569.
Koike, K. and Okui, S. (1964). J. Biochem. [Tokyo) 55, 195.
Koishi, T. (1959 a). Japan. J. Pharmacol. 8, 101.
Koishi, T. (1959 b). Japan. J. Pharmacol. 8, 124.
Koivusalo, M. and Luukkainen, T. (1959). Acta Physiol. Scand. 45, 283.
Kolthoff, I.M. and Anastasi, A. (1958). J. Am. Chem. Soc. 80, 4248.
KolthofF, I.M., Stricks, W., and Morren, L. (1954). Anal. Chem. 26, 366.
Kominz, D.R. (1961). Biochim. Biophys. Acta 51, 456.
Kono, M. and Quastel, J.H. (1962). Biochem. J. 85, 24.
Kono, T. and Colowick, S.P. (1961). Arch. Biochem. Biophys. 93, 514.
Korey, S. (1950). Biochim. Biophys. Acta 4, 58.
Korn, E.D. (1962). J. Biol. Chem. 237, 3423.
Kornberg, H.L. and Krebs, H.A. (1957). Nature 179, 988.
Kornguth, S.E. and Stahmann, M.A. (1960). Arch. Biochem. Biophys. 91, 32.
1028 REFERENCES
Kostytschew, S. and Berg, V. (1930). Z. Physiol. Chem. 188, 133.
Kotaka, S. (1963). J. Gen. Physiol. 46, 1087.
Koukol, J. and Conn, E.E. (1961). J. Biol. Chem. 236, 2692.
Kovac, L. (1960). Enzymologia 11, 27.
Kovachevich, R. and Wood, W.A. (1955). J. Biol. Chem. 213, 745.
Krahe, E. (1924). Klin. Wochschr. 3, 70.
Krahl, M.E. and Clowes, G.H.A. (1940). J. Gen. Physiol. 23, 413.
Kramer, M. and Arrigoni-Martelli, E. (1960). Arch. Ezptl. Pathol. Pharmalcol. 238, 43.
Kramer, P.J. (1951). Plant Physiol. 26, 30.
Krane, S.M. and Crane, R.K. (1959). J. Biol. Chem. 234, 211.
Krasna, A.I. (1961). J. Biol. Chem. 236, 749.
Krasna, A.I. and Rittenberg, D. (1954). Proc. Natl. Acad. Sci. U.S. 40, 225.
Krasna, A.I., Riklis, E., and Rittenberg, D. (1960). J. Biol. Chem. 235, 2717.
Krebs, E.G. and Norris, E.R. (1949). Arch. Biochem. 24, 49.
Krebs, E.G., Graves, D.J., and Fischer, E.H. (1959). J. Biol. Chem. 234, 2867.
Krebs, H.A. (1930). Biochem. Z. 220, 289.
Krebs, H.A. (1935). Biochem. J. 29, 1951.
Krebs, H.A. and Eggleston, L.V. (1940). Biochem. J. 34, 442.
Krebs, H.A. and Eggleston, L.V. (1944). Biochem. J. 38, 426.
Krebs, H.A. and Eggleston, L.V. (1948). Biochim. Biophys. Acta 1, 319.
Krebs, H.A. and Johnson, W.A. (1948). Tabulae Biol. 19, 100.
Krebs, H.A. and Lowenstein, J.M. (1960). In "Metabolic Pathways" (D.M. Greenberg
ed.). Vol. 1, p. 129. Academic Press, New York.
Krebs, H.A., Salvin, E., and Johnson, W.A. (1938). Biochem. J. 32, 113.
Krebs, H.A., Gurin, S., and Eggleston, L.V. (1952). Biochem. J. 51, 614.
Kreke, C.W., Kroger, M.H., and Cook, E.S. (1949). J. Biol. Chem. 180, 565.
Kreke, C.W., Schaefer, M.A., Seibert, M.A., and Cook, E.S. (1950). J. Biol. Chem.
185, 469.
Krisch, K. (1963). Biochem. Z. 337, 546.
Kriszat, G. and Runnstrom, J. (1952). Exptl. Cell Res. 3, 500.
Krueger, A.P. and Baldwin, D.M. (1933). J. Gen. Physiol. 17, 129.
Krueger, A.P. and Baldwin, D.M. (1934). J. Gen. Physiol. 17, 499.
Krueger, R. (1950). Experientia 6, 106.
Krueger, R.C. (1955). Arch. Biochem. Biophys. 57, 52.
Krusius, F.-E. (1940). Acta Physiol. Scand. Suppl. 3.
Kubowitz, F. and Ott, P. (1941). Naturwissenschaften 39, 590.
Kuby, S.A., Noda, L., and Lardy, H.A. (1954). J. Biol. Chem. 210, 65.
Kuhn, R. (1923). Z. Physiol. Chem. 129, 57.
Kuhn, R. and Beinert, H. (1943). Ber. 76, 904.
Kuhn, R. and Beinert, H. (1947). Ber. 80, 101.
Kuhn, R. and Desnuelle, P. (1938). Z. Physiol. Chem. 251, 14.
Kuhn, R. and Franke, W. (1935). Ber. 68, 1528.
Kuhn, R., Katz, H., and Franke, W. (1934). Naturwissenschaften 11, 808.
Kull, F.C., Grimm, M.R., and Mayer, R.L. (1954). Proc. Soc. Exptl. Biol. Med. 86,
330.
Kun, E., Dechary, J.M., and Pitot, H.C. (1954). J. Biol. Chem. 210, 269.
Kun, E., Grassetti, D.R., Fanshier, D.W., and Featherstone, R.M. (1958). Biochem.
Pharmacol. 1, 207.
REFERENCES 1029
Kun, E., Fanshier, D.W., and Grassetti, D.R. (1960). J. Biol. Chem. 235, 416.
Kunz, H.A. (1956). Helv. Physiol. Pharmacol. Acta 14, 411.
Kunz, W. (1963). Z. Physiol. Chem. 334, 128.
Kuo, M.-H. and Blumenthal, H.J. (1961). Biochim. Biophys. Acta 52, 13.
Kupiecki, F.P. and Coon, M.J. (1959). J. Biol. Chem. 234, 2428.
Kupiecki, F.P. and Virtanen, A.I. (1960). Acta Chem. Scand. 14, 1913.
Kuratomi, K. (1959). J. Biochem. (Tokyo) 46, 453.
Kuratomi, K. and Fukunaga, K. (1963). Biochim. Biophys. Acta 78, 617.
Kurtz, A.N. and Niemann, C. (1961). Biochim. Biophys. Acta 50, 520.
Kurup, C.K.R. and Vaidyanathan, C.S. (1963). Biochem. J. 88, 239.
Kuschinsky, G. and Liillmann, H. (1954). Arch. Exptl. Pathol. Pharmakol. 11^, 299.
Kuschinsky, G. and Turba, F. (1950). Naturwissenschaften 37, 425.
Kuschinsky, G. and Turba, F. (1951). Biochim. Biophys. Acta 6, 426.
Kuschinsky, G., Mercker, H., Westermann, E., and Liillmann, H. (1953). Arch. Exptl.
Pathol. Pharmakol. 217, 401.
Kusunose, M., Nagai, S., Kusunose, E., and Yamamura, Y. (1956). J. Bacteriol. 72,
754.
Kusunose, E., Kusunose, M., Kowa, Y., and Yamamura, Y. (1960). J. Biochem. (To-
kyo) 47, 689.
Kutscha, W. (1961). Arch. Ges. Physiol. 273, 409.
Kutscher, W. and Hasenfuss, M. (1940). Z. Physiol. Chem. 265, 181.
Kutscher, W. and Sarreither, W. (1940). Z. Physiol. Chem. 265, 152.
Kuttner, R. and Wagreich, H. (1953). Arch. Biochem. Biophys. 43, 80.
Kvam, D.C. and Parks, R.E., Jr. (1960). J. Biol. Chem. 235, 2893.
Kvamme, E. (1957). Acta Chem. Scand. 11, 1091.
Kvamme, E. (1958 a). Acta Physiol. Scand. 42, 204.
Kvamme, E. (1958 b). Acta Physiol. Scand. 42, 219.
Kvamme, E. (1958 c). Acta Physiol. Scand. A2, 231.
Kvamme, E. (1958d). Acta Physiol. Scand. 42, 239.
Labbe, R.F. and Hubbard, N. (1961). Biochim. Biophys. Acta 52, 130.
Labeyrie, F. and Stachiewicz, E. (1961). Biochim. Biophys. Acta 52, 136.
Ladd, J.N. (1956). Australian J. Biol. Sci. 9, 92.
Ladd, J.N. and Nossal, P.M. (1954). Australian J. Exptl. Biol. Med. Sci. 32, 523.
La Du, B.N. and Zannoni, V.G. (1955). J. Biol. Chem. 217, 777.
Lagnado, J.R. and Sourkes, T.L. (1956). Can. J. Biochem. Physiol. 34, 1185.
Laki, K. (1935). Z. Physiol. Chem. 236, 31.
Lakshmanan, M.R., Vaidyanathan, C.S., and Cama, H.R. (1964). Biochem. J. 90, 569.
Lambie, A.T. and Robson, J.S. (1960). Clin. Sci. 20, 123.
Lambooy, J.P. (1950). J. Am. Chem. Soc. 72, 5225.
Lambooy, J.P. (1955). Natl. Vitamin Found. Nutr. Symp. Sec. 11, p. 12.
Landau, B.R., Laszlo, J., Stengle, J., and Burk, D. (1958). J. Natl. Cancer Inst. 21,
485.
Landau, J.V., Marsland, D., and Zimmerman, A.M. (1954). Anat. Record. 120, 789.
Landon, E.J. and Carter, C.E. (1960). J. Biol. Chem. 235, 819.
Landon, E.J. and Norris, J.L. (1963). Biochim. Biophys. Acta 71, 266.
Lands, W.E.M. and Niemann, C. (1959). J. Am. Chem. Soc. 81, 2204.
Lane, M. and Brindley, CO. (1964). Proc. Soc. Exptl. Biol. Med. 116, 57.
1030 REFERENCES
Lane, M., Fahey, J.L., Sullivan, R.D., and Zubrod, C.G. (1958). J. Pharmacol. Ezptl.
Therap. 122, 315.
Lane, M., Petering, H.G., and Brindley, CO. (1959). J. Natl. Cancer Inst. 22, 349.
Lang, C.A. and Nason, A. (1959). J. Biol. Chem. 234, 1874.
Lang, K. and Bassler, K.H. (1953). Biochem. Z. 323, 456.
Langdon, R.G. and Weakley, D.R. (1957). Federation Proc. 16, 208.
Lange, C.F., Jr. and Kohn, P. (1961 a). J. Biol. Chem. 236, 1.
Lange, C.F., Jr. and Kohn, P. (1961b). Cancer Pes. 21, 1055.
Langer, L.J. and Engel, L.L. (1958). J. Biol. Chem. 233, 583.
Lara, F.J.S. (1952). J. Bacteriol. 64, 279.
Lardy, H.A. and Philipps, P.H. (1943 a). J. Biol. Chem. 148, 333.
Lardy, H.A. and Phillips, P.H. (1943 b). J. Biol. Chem. 148, 343.
Lardy, H.A. and Phillips, P.H. (1945). Arch. Biochem. 6, 53.
Lardy, H.A. and Wellman, H. (1953). J. Biol. Chem. 201, 357.
Lardy, H.A., Wiebelhaus, V.D., and Mann, K.M. (1950). J. Biol. Chem. 187, 325.
Laris, P.O., Ewers, A., and Novinger, G. (1962). J. Cellular Comp. Physiol. 59, 145.
Lamer, J. and Gillespie, R.E. (1956). J. Biol. Chem. 223, 709.
Lamer, J. and Schliselfeld, L.H. (1956). Biochim. Biophys. Acta 20, 53.
Larner, J., Jackson, W.T., Graves, D.J., and Stamer, J.R. (1956). Arch. Biochem.
Biophys. 60, 352.
Larrabee, M.G. and Bronk, D.W. (1952). Cold Spring Harbor Symp. Quant. Biol. 17,
245.
Larson, F.C. and Albright, E.G. (1961). J. Clin. Invest. 40, 1132.
Lascelles, J. (1956). Biochem. J. 62, 78.
Lassen, U.V. and Overgaard-Hansen, K. (1962). Biochim. Biophys. Acta 57, 111.
Laszio, J. Landau, B.R., Wight, K., and Burk, D. (1958). J. Natl. Cancer Inst. 21, 475.
Laszlo, J., Humphreys, S.R., and Goldin, A. (1960). J. Natl. Cancer Inst. 24, 267.
Laszio, J., Harlan, W.R., Klein, R.F., Kirshner, N., Estes, E.H., Jr., and BogdonofF,
M.D. (1961). J. Clin. Invest. 40, 171.
Lathe, G.H. and Walker, M. (1958). Biochem. J. 70, 705.
Laties, G.G. (1949a). Arch. Biochem. 20, 284.
Laties, G.G. (1949 b). Arch. Biochem. 22, 8.
Laties, G.G. (1953). Phtjsiol. Plantarum 6, 215.
Laties, G.G. (1959 a). Arch. Biochem. Biophys. 79, 364.
Laties, G.G. (1959 b). Arch. Biochem. Biophys. 79, 378.
Laties, G.G. (1964). Plant Physiol. 39, 391.
Latuasan, H.E. and Berends, W. (1961). Biochim. Biophys. Acta 52, 502.
Launoy, L. and Levaditi, C. (1913). Compt. Rend. Soc. Biol. 72, 653.
Lazdunski, M. and Ouellet, L. (1962). Can. J. Biochem. Physiol. 40, 1619.
Lazzarini, R.A. and Atkinson, D.E. (1961). J. Biol. Chem. 236, 3330.
Lazzarini, R.A. and San Pietro, A. (1964). Arch. Biochem. Biophys. 106, 6.
Leach, S.J. (1960). Australian J. Chem. 13, 520.
Lebrun, J. and Delaunay, A. (1951). Ann. Inst. Pasteur 80, 524.
Leder, I.G. (1957). J. Biol. Chem. 225, 125.
Ledoux, L. (1953). Biochim. Biophys. Acta 11, 517.
Ledoux, L. (1954). Biochim.. Biophys. Acta 13, 121.
Lee, C.-P., Azzone, G.F., and Ernster, L. (1964). Nature 201, 152.
Lee, H.A., Jr. and Abeles, R.H. (1963). J. Biol. Chem. 238, 2367.
REFERENCES 1031
Lee, J.S. and Lifson, N. (1951). J. Biol. Chem. 193, 253.
Lee, Y.-P. (1957). J. Biol. Chem. 227, 999.
Lee, Y.-P. (1960 a). Biochim. Biophys. Acta 43, 18.
Lee, Y.-P. (1960 b). Biochim. Biophys. Acta 43, 25.
Lees, H. and Simpson, J.R. (1955). Biochem. J. 59, i.
Lees, H. and Simpson, J.R. (1957). Biochem. J. 65, 297.
LeFevre, C.G. and LeFevre, R.J.W. (1937). J. Chem. Soc. p. 1088.
LeFevre, P.G. (1947). Biol. Bull. 93, 224.
LeFevre, P.G. (1948). J. Gen. Physiol. 31, 505.
LeFevre, P.G. (1954). Symp. Soc. Exptl. Biol. 8, 118.
LeFevre, P.G. and Davies, R.I. (1951). J. Gen. Physiol. 34, 515.
LeFevre, P.G. and Marshall, J.K. (1958). Am. J. Physiol. 194, 333.
Legge, J.W. and Turner, A.W. (1954). Australian J. Biol. Sci. 7, 496.
Lehman, I.R. and Nason, A. (1956). J. Biol. Chem. Ill, 497.
Lehman, I.R., Roussos, G.G., and Pratt, E.A. (1962 a). J. Biol. Chem. 237, 819.
Lehman, I.R., Roussos, G.G., and Pratt, E.A. (1962 b). J. Biol. Chem. 237. 829.
Lehman, R.A., King, E.E., and Taube, H. (1950). J. Pharmacol. Exptl. Therap. 99,
149.
Lehmann, J. (1938). Skand. Arch. Physiol. 80, 237.
Lehninger, A.L. (1946 a). J. Biol. Chem. 164, 291.
Lehninger, A.L. (1946 b). J. Biol. Chem. 165, 131.
Lehninger, A.L. (1949). J. Biol. Chem. 178, 625.
Lehninger, A.L. (1951). In "Phosphorus MetaboHsm" (W.D. McElroy and H.B. Glass,
eds.). Vol. 1, p. 344. Johns Hopkins Press, Baltimore, Maryland.
Lehninger, A.L. and Kennedy, E.P. (1948). J. Biol. Chem. 173, 753.
Lehinger, A.L., Wadkins, C.L., Cooper, C, Devlin, T.M., and Gamble, J.L., Jr. (1958).
Science 128, 450.
Leloir, L.F. and Goldemberg, S.H. (1960). J. Biol. Chem. 235, 919.
Leloir, L.F., Olavarria, J.M., Goldemberg, S.H., and Carminatti, H. (1959). Arch.
Biochem. Biophys. 81, 508.
Lembeck, F. and Resch, H. (1960). Arch. Exptl. Pathol. Pharmakol. 240, 210.
Lenney, J.F. (1956). J. Biol. Chem. ll^, 919.
Lenti, C. and Grillo, M.A. (1955). Z. Physiol. Chem. 302, 253.
Lenz, G.R. and Martell, A.E. (1964). Biochemistry 3, 745.
Lenzi, F. and Caniggia, A. (1953). Folia Cardiol. 12, No. 3 and 4.
Lerner, A.B., Fitzpatrick, T.B., Calkins, E., and Summerson, W.H. (1951). J. Biol.
Chem. 191, 799.
Leshe, J., Butler, L.G., and Gorin, G. (1962 a). Arch. Biochem. Biophys. 99, 86.
Leshe, J., WilUams, D.L., and Gorin, G. (1962 b). Anal. Biochem. 3, 257.
Lester, G. and Bonner, D.M. (1952). J. Bacteriol. 63, 759.
Letnansky, K. and Seehch, F. (1960). Z. Krehsforsch. 64, 1.
Leuschner, F., Heim, F., and Poertzel, H. (1957). Arch. Intern. Pharmacodyn. 110, 177.
Levenberg, B., Melnick, I., and Buchanan, J.M. (1957). J. Biol. Chem. 225, 163.
Levey, H.A. and Szego, CM. (1955). Am. J. Physiol. 182, 507.
Levey, S. and Sheinfeld, S. (1954). Gastroenterology T7, 625.
Levvy, G.A. (1952). Biochem. J. 52, 464.
Levvy, G.A. (1954). Biochem. J. 58, 462.
Levvy, G.A. and Marsh, C.A. (1952). Biochem. J. 52, 690.
1032 REFERENCES
Levvy, G.A. and Marsh, C.A. (1960). In "The Enzymes" (P.D. Boyer, H. Lardy, and
K. Myrback, eds.). Vol. 4, p. 397. Academic Press, New York.
Levvy, G.A. and Worgan, J.T. (1955). Biochem. J. 59, 451.
Levvy, G.A., Hay, A. J., and Marsh, C.A. (1957). Biochem. J. 65, 203.
Levvy, G.A., McAllan, A., and Hay, A.J. (1962). Biochem. J. 82, 225.
Levy, H.M., Sharon, N., Ryan, E.M., and Koshland, D.E., Jr. (1962). Biochim. Bio-
phys. Acta 56, 118.
Levy, H.R. and Talalay, P. (1959). J. Biol. Chem. 234, 2014.
Levy, R.I., Weiner, I.M., and Mudge, G.H. (1958). J. Clin. Invest. 37, 1016.
Lewin, J. (1951). J. Am. Chem. Soc. 73, 3906.
Lewis, S.E. and Slater, E.G. (1954). Biochem. J. 58, 207.
Li, C.H. (1945). J. Am. Chem. Soc. 67, 1065.
Li, T.-K., Ulmer, D.D., and Vallee, B.L. (1962). Biochemistry 1, 114.
Li, Y., Li, S.C., and Shetlar, M.R. (1963). Arch. Biochem. Biophys. 103, 436.
Lichtenstein, N., Ross, H.E., and Cohen, P.P. (1953). Nature 171, 45.
Liebecq, C. and Peters, R.A. (1949). Biochim. Biophys. Acta 3, 215.
Lieben, F. and Bauminger, B. (1933a). Biochem. Z. 261, 374.
Lieben, F. and Bauminger, B. (1933 b). Biochem. Z. 261, 387.
Lieber, E. and Smith, G.B.L. (1939). Chem. Rev. 25, 213.
Lieberman, I. (1956). J. Biol. Chem. 223, 327.
Lieberman, M. (1961). Plant Physiol. 36, 804.
Lieberman, M. and Biale, J.B. (19.56 a). Plant Physiol. 31, 420.
Lieberman, M. and Biale, J.B. (1956 b). Plant Physiol. 31, 425.
Liener, I.E. (1961). Biochim.. Biophys. Acta 53, 332.
Lifson, N. and Stolen, J.A. (1950). Proc. Soc. Exptl. Biol. Med. 74, 451.
LindeU, S.E., Nilsson, K., Rocs, B.-E., and Westling, H. (1960). Brit. J. Pharmacol.
15, 351.
Linderholm, H. (1952). Acta Physiol. Scand. 27, Suppl. 97.
Linderstrom-Lang, K. and Jacobsen, C.F. (1941). J. Biol. Chem. 137, 443.
Lindner, R.C., Kirkpatrick, H.C., and Weeks, T.E. (1959). Phytopathology 49, 802.
Lindstrom, E.S., Burris, R.H., and Wilson, P.W. (1949). J. Bacterial. 58, 313.
Lineweaver, H. (1933). J. Biol. Chem. 99, 575.
Ling, G. and Gerard, R.W. (1949). J. Cellular Comp. Physiol. 34, 413.
Link, G.K.K., Klein, R.M., and Barron, E.S.G. (1952). J. Exptl. Botany 3, 216.
Lipke, H. and Kearns, C.W^ (1959). J. Biol. Chem. 234, 2129.
Lipsett, M.N. (1964). J. Biol. Chem. 239, 1250.
Lister, A.J. (1956). J. Gen. Microbiol. 14, 478.
Litchfield, J.H. and Ordal, Z.J. (1958). Can. J. Microbiol. 4, 205.
Livermore, A.H. and Sealock, R.R. (1947). J. Biol. Chem. 167, 699.
Lloyd, D.R. and Prince, R.H. (1961). Proc. Chem. Soc. p. 464.
Lockwood, W.H. and Benson, A. (1960). Biochem. J. 75, 372.
Lodin, Z., Janacek, K., and Miiller, J. (1963). J. Cellular Comp. Physiol. 62, 215.
Loefer, J.B. (1951). Cancer Res. 11, 481.
Loevenhart, A.S. and Grove, W.E. (1909). J. Pharmacol. Exptl. Therap. 1, 289.
Loevenhart, A.S. and Grove, W.E. (1911). J. Pharmacol. Exptl. Therap. 3, 101.
London, M., McHugh, R., and Hudson, P.B. (1958). Arch. Biochem. Biophys. 73, 72.
Long, W.K. and Farah, A. (1946). J. Pharmacol. Exptl. Therap. 88, 388.
Loomis, W.F. and Lipmann, F. (1948). J. Biol. Chem. 173, 807.
REFERENCES 1033
Lorand, L. and Rule, N.G. (1961). Nature 190, 722.
Lord, C.F. and Husa, W.J. (1954). J. Am. Pharm. Assoc, Sci. Ed. 43, 438.
Lorenz, B., Lorenz, R., and Zollner, N. (1960). Z. Naturforsch. 15b, 62.
Lotspeich, W.D. and Peters, R.A. (1951). Biochem. J. 49, 704.
Lotspeich, W.D. and Woronkow, S. (1964). Am. J. Physiol. 206, 391.
Loureiro, J.A. de and Lito, E. (1946). J. Hyg. 44, 463.
Loveless, L.E., Spoeri, E., and Weisman, T.H. (1954). J. Bacteriol. 68, 637.
Lovenberg, W., Barchas, J., Weissbach, H., and Udenfriend, S. (1963). Arch. Bio-
chem. Biophys. 103, 9.
Levtrup, S. and Svennerholm, L. (1963). Exptl. Cell. Res. 29, 298.
Low, B.W. (1952). J. Am. Chem. Soc. 74, 4830.
Low, B.W. and Weichel, E.J. (1951). J. Am. Chem. Soc. 73, 3911.
Low, H. (1959 a). Biochim. Biophys. Acta 32, 1.
Low, H. (1959 b). Exptl. Cell. Res. 16, 456.
Low, H., Siekevitz, P., Ernster, L., and Lindberg, O. (1958). Biochim. Biophys. Acta
29, 392.
Low, H., Lindberg, O., and Ernster, L. (1959). Biochim. Biophys. Acta 35, 284.
Lowry, O.H. (1943). Biol. Symp. 10, 233.
Lowry, O.K., Bessey, O.A., and Crawford, E.J. (1949 a). J. Biol. Chem. 180, 389.
Lowry, O.H., Bessey, O.A., and Crawford, E.J. (1949 b). J. Biol. Chem. 180, 399.
Liick, H. (1957). Biochem. Z. 328, 411.
Ludwig, B.J. and Nelson, J.M. (1939). J. Am. Chem. Soc. 61, 2601.
Luh, B.S. and Phaff, H.J. (1954). Arch. Biochem. Biophys. 48, 23.
Lukomskaia, I.S. and Rozenfel'd, E.L. (1958). Biochemistry (USSfi) (English Transl.)
23, 244.
Luteraan, P.J. (1953). Compt. Rend. 236, 2531.
Lutwak-Mann, C. (1947). Biochem. J. 41, 19.
Luzzato, R. and Satta, G. (1910). Arch. Farmacol. Sper. 8, 554.
Lwoff, A. and Audureau, A. (1947). An7i. Inst. Pasteur 73, 517.
Lwoff, A., Audereau, A., and Cailleau, R. (1947). Compt. Rend. 11^, 303.
Lynen, F. (1943). Ann. 554, 40.
Lynen, F., Reichert, E., and Rueff, L. (1951). Ann. 574, 1.
Lyon, I. and Geyer, R.P. (1954). J. Biol. Chem. 208, 529.
McCaUa, T.M. and Foltz, V.D. (1941). Trans. Kansas Acad. Sci. 44, 46.
McComb, R.B. and Yushok, W.D. (1959). Biochim. Biophys. Acta 34, 515.
McComb, R.B. and Yushok, W.D. (1964 a). Cancer Res. 24, 193.
McComb, R.B. and Yushok, W.D. (1964 b). Cancer Res. 24, 198.
McCormick, D.B. (1962). J. Biol. Chem. 237, 959.
McCormick, D.B. (1964). Nature 201, 295.
McCormick, D.B. and Snell, E.E. (1961). J. Biol. Chem. 236, 2085.
McCormick, D.B., Gregory, M.E., and Snell, E.E. (1961). J. Biol. Chem. 236, 2076.
McCormick, N.G., Ordal, E.J., and Whiteley, H.R. (1962). J. Bacteriol. 83, 887.
McCrea, F.D. and Meek, W.J. (1929). J. Pharmacol. Exptl. Therap. 36, 295.
McCurdy, H.D., Jr. and Cantino, E.G. (1960). Plant Physiol. 35, 463.
McDaniel, E.G., Hundley, J.M., and Sebrell, W.H. (1955). J. Nutr. 55, 623.
McDonald, J.K., Anderson, J.A., Cheldelin, V.H., and King, T.E. (1963). Biochim.
Biophys. Acta 73, 533.
1034 REFERENCES
Macdonald, K. (1961). Biochem. J. 80, 154.
MacDonnell, L.R., Silva, R.B., Feeney, R.E. (1951). Arch. Biochem.. Biophys. 32, 288.
Macdowall, F.D.H. (1949). Plant Physiol. 24, 462.
McEwen, B.S., Allfrey, V.G., and Mirsky, A.E. (1963 a). J. Biol. Chem. 238, 2571.
McEwen, B.S., Allfrey, V.G., and Mirsky, A.E. (1963 b). J. Biol. Chem. 238, 2579.
McFadden, B.A. and Atkinson, D.E. (1957). Arch. Biochem. Biophys. 66, 16.
McFadden, B.A. and Howes, W.V. (1963). J. Biol. Chem. 238, 1737.
Macfarlane, E.W.E. (1951). Growth 15, 241.
Macfarlane, E.W.E. (1953). Ezptl. Cell Ees. 5, 375.
Macfarlane, E.W.E. and Nadeau, L.V. (1948). Proc. Soc. Exptl. Biol. Med. 67, 118.
Macfarlane, W.V. and Meares, J.D. (1958). J. Physiol. {London) 142, 78.
McGarrahan, J.F. and Maley, F. (1962). J. Biol. Chem. 237, 2458.
McGowan, J.C, Brian, P.W., and Hemming, H.G. (1948). Ann. Appl. Biol. 35, 25.
McGuire, J.S., Jr., Hollis, V.W., Jr., and Tomkins, G.M. (1960). J. Biol. Chem. 235,
3112.
Machlis, L. (1944). Am. J. Botany 31, 183.
Macht, D.I. (1931 a). Am. J. Botany 18, 598.
Macht, D.I. (1931b). Proc. Soc. Exptl. Biol. Med. 28, 687.
Mcllwain, H. (1940). Brit. J. Exptl. Physiol. 21, 136.
Mcllwain, H. (1941). Brit. J. Exptl. Physiol. 11, 148.
Mcllwain, H. (1945). Biochem. J. 39, 329.
Mcllwain, H. (1950). Biochem. J. 46, 612.
Mcllwain, H. (1959). Biochem.. J. 71, 281.
Mcllwain, H. and Rodnight, R. (1949). Biochem. J. 44, 470.
Mackay, D. and Shepherd, D.M. (1962). Biochim. Biophys. Acta 59, 553.
McKinney, G.R. and Gemma, F.E. (1958). J. Pharmacol. Exptl. Therap. 122, 51A.
McKinney, G.R., Smith, L.C., and Martin, S.P. (1955). J. Pharmacol. Exptl. Therap.
113, 37.
Mackler, B. (1961). Biochim. Biophys. Acta 50, 141.
Mackler, B., Mahler, H.R., and Green, D.E. (1954). J. Biol. Chem. 210, 149.
MacLennan, D.H. and Beevers, H. (1964). Phytochemistry 3, 109.
MacLeod, J. (1951). J. Gen. Physiol. 34, 705.
MacLeod, R.A. and Snell, E.E. (1950). Ann. N. Y. Acad. Sci. 52, 1249.
MacLeod, R.M., Farkas, W., Fridovich, I., and Handler, P. (1961). J. Biol. Chem.
236, 1841.
McMahon, R.E. and Mills, J. (1961). J. Med. Pharm. Chem. 4, 211.
MacNider, W. deB. (1918 a). J. Pharmacol. Exptl. Therap. 11, 186.
MacNider, W. deB. (1918 b). Proc. Soc. Exptl. Biol. Med. 16, 82.
MacNider, W. deB. (1941). J. Pharmacol. Exptl. Therap. 73, 186.
McShan, W.H. and Meyer, R.K. (1946). Arch. Biochem. 9, 165.
McShan, W.H., Kramer, S., and Schlegel, V. (1954). Biol. Bull. 106, 341.
Madinaveitia, J. (1946). Biochem. J. 40, 373.
Madsen, N.B. (1956). J. Biol. Chem. 113, 1067.
Madsen, N.B. (1960). Can. J. Biochem. Physiol. 38, 481.
Madsen, N.B. and Cori, C.F. (1955). Biochim. Biophys. Acta 18, 156.
Madsen, N.B. and Cori, C.F. (1956). J. Biol. Chem. 223, 1055.
Madsen, N.B. and Gurd, F.R.N. (1956). J. Biol. Chem. 223, 1075.
Maengwyn-Davies, G.D. and Friedenwald, J.S. (1954). Arch. Biochem. Biophys. 53,29.
REFERENCES 1035
Mager, J. and Magasanik, B. (1960). J. Biol. Chem. 235, 1474.
Mahadevan, S., Murthy, S.K., and Ganguly, J. (1962). Biochem. J. 85, 326.
Mahler, H.R. (1954). J. Biol. Chem. 206, 13.
Mahler, H.R. (1955). In "Methods in Enzymology" (S.P. Colowick and N.O. Kaplan,
eds.). Vol. 2, p. 688. Academic Press, New York.
Mahler, H.R. (1956). Advan. Enzyi7wl. 17, 233.
Mahler, H.R., Mackler, B., Green, D.E., and Bock, R.M. (1954). J. Biol. Chem. 210,
465.
Mahler, H.R., Fairhurst, A.S., and Mackler, B. (1955a). J. Am. Chem. Soc. 77, 1514.
Mahler, H.R., Hiibscher, G., and Baum, H.M. (1955 b). J. Biol. Chem. 216, 625.
Mahler, H.R., Baum, H.M., and Hiibscher, G. (1956). Science 124, 705.
Mahler, H.R., Raw, I., Molinari, R., and do Amaral, D.F. (1958). J. Biol. Chem. 233,
230.
Maio, J.J. and Rickenberg, H.V. (1962). Exptl. Cell. Res. 11. 31.
Maitra, P.K. and Estabrook, R.W. (1962). Biochem. Biophys. Res. Commun. 9, 1.
Maitra, P.K. and Roy, S.C. (1959). J. Biol. Chem. 234, 2497.
Maitra, P.K. and Roy, S.C. (1961). Biochem. J. 79, 446.
Maitre, L. and Staehelin, M. (1963). Experientia 19, 573.
Maizels, M. (1951). J. Physiol. [London) 112, 59.
Maizels, M. and Remington, M. (1958). J. Physiol. {London) 143, 283.
Maizels, M., Remington, M., and Truscoe, R. (19.58). J. Physiol. (London) 140, 80.
Makino, K. and Koike, M. (1954 a). Nature 174, 1056.
Makino, K. and Koike, M. (1954 b). Enzymologia 17, 157.
Malachowski, R. and Wanczura, T. (1933). Bull. Intern. Acad. Polon. Sci., Classe Sci.
Math. Nat. p. 547.
Maley, F. (1958). Federation Proc. 17, 267.
Maley, F. and Lardy, H.A. (1955). J. Biol. Chem. 214, 765.
Maley, F. and Ochoa, S. (1958). J. Biol. Chem. 233, 1538.
Maley, G.F. and Maley, F. (1959). J. Biol. Chem. 234, 2975.
Maley, G.F. and Maley, F. (1964). J. Biol. Chem. 239, 1168.
Malkin, A. and Denstedt, O.F. (1956). Can. J. Biochem. Physiol. 34, 141.
Mallus, E. (1931). Clin. Pediat. 13, 265.
Malmstrom, B.G. (1962). Arch. Biochem. Biophys. Suppl. 1, p. 247.
Malvin, R.L. (1956). Federation Proc. 15, 125.
Mancilla, R. and Agosin, M. (1960). Exptl. Parsitol. 10, 43.
Mandel, H.G. (1955). In "Antimetabolites and Cancer" (C.P. Rhoads, ed.), p. 199,
Am. Assoc. Advance. Sci., Washington, D.C.
Mandelstam, J. (1961). Biochem. J. 79, 489.
Mann, F.D. and Mann, F.C. (1939). Am. J. Digest. Diseases 6, 322.
Mann, P.J.G. and Quastel, J.H. (1941). Biochem. J. 35, 502.
Manning, D.T. and Niemann. C. (1958). J. Am. Chem. Soc. 80, 1478.
Manning, G.B. and Campbell, L.L., Jr. (1957). Can. J. Microbiol. 3, 1001.
Manning, G.B. and Campbell, L.L., Jr. (1961). J. Biol. Chem. 236, 2952.
Mano, Y. and Tanaka, R. (1960). J. Biochem. (Tokyo) 47, 401.
Mansour, T.E. (1963). J. Biol. Chem. 238, 2285.
Mapson, L.W. (1946). Biochem. J. 40, 240.
Mapson, L.W. and Breslow, E. (1958). Biochem. J. 68, 395.
Mapson, L.W. and Isherwood, F.A. (1963). Biochem. J. 86, 173.
1036 REFERENCES
Marcus, G.J. and Manery, J.F. (1963). Cab. J. Biochem. Physiol. 41, 2529.
Marcus, P.I. and Talalay, P. (1955). Proc. Roy. Soc. B144, 116.
Mardashev, S.R. and Semina, L.A. (1961). Biochemistry (USSR) (English Transl.) 26,
27.
Marinetti, G.V., Erbland, J., Albrecht, M., and Stotz, E. (1957). Biochim. Biophys.
Acta 26, 130.
Marini, M. and Levey, S. (1955). Proc. Soc. Ezptl. Biol. Med. 88, 611.
Marks, P.A., Szeinberg, A., and Banks, J. (1961). J. Biol. Chem. 236, 10.
Markwardt, F. (1953 a). Naturwissenschaften 40, 341.
Markwardt, F. (1953 b). Z. Physiol. Chem. 295, 264.
Marre, E. and Servettaz, 0. (1958). Arch. Biochem. Biophys. 75, 309.
Marre, E., Forti, G., and Pece, G. (1956). Atti Accad. Naz. Lincei Rend., Classe Sci.
Fis., Mat. Nat. [8] 20, 646.
Marsh, C.A. and Levvy, G.A. (1957). Biochem. J. 66, 4p.
Marsh, C.A. and Levvy, G.A. (1958). Biochem. J. 68, 610.
Marshall, M., Metzenberg, R.L., and Cohen, P.P. (1961). J. Biol. Chem. 236, 2229.
Marshall, P.B. (1948). Brit. J. Pharmacol. 3, 1.
Martell, A.E. and Calvin, M. (1952). "Chemistry of the Metal Chelating Compounds."
Prentice-Hall, Englewood Cliffs, New Jersey.
Martin, G.J. (1951). "Biological Antagonism." McGraw-Hill (Blakiston), New York.
Martin, G.J. and Beiler, J.M. (1947). Arch. Biochem. 15, 201.
Martin, G.J. and Beiler, J.M. (1948). J. Am. Pharm. Assoc, Sci. Ed. 37, 32.
Martin, G.J., Tolman, L., and Brendel, R. (1947). Arch. Biochem. 15, 324.
Martin, G.J., Brendel, R., Graff, M., and Beiler, J.M. (1950). J. Am. Pharm. Assoc,
Sci. Ed. 39, 338.
Martin, R.G. (1963). J. Biol. Chem. 238, 257.
Martin, S.M. (1954). Can. J. Microbiol. 1, 6.
Martin, S.M. (1955). Can. J. Microbiol. 1, 644.
Martonosi, A. and Gouvea, M.A. (1961). J. Biol. Chem. 236, 1345.
Martonosi, A. and Meyer, H. (1964). J. Biol. Chem. 239, 640.
Maschmann, E. (1937 a). Biochem. Z. 294, 1.
Maschmann, E. (1937 b). Biochem. Z. 295, 1.
Maschmann, E. and Helmert, E. (1933 a). Z. Physiol. Chem. 219, 99.
Maschmann, E. and Helmert, E. (1933 b). Z. Physiol. Chem. 220, 199.
Mason, H.L. (1930). J. Biol. Chem. 86, 623.
Mason, M. (1959). J. Biol. Chem. 234, 2770.
Mason, S.F. (1957). Ciba Found. Symp., Chem. Biol. Purines p. 60.
Massart, L., Vandendriessche, L., and Dufait, R. (1940). Enzymologia 8, 204.
Massey, V. (1953 a). Biochem. J. 53, 67.
Massey, V. (1953 b). Biochem. J. 55, 172.
Massey, V. (1958). Biochim. Biophys. Acta 30, 500.
Massey, V. and Rogers, W.P. (1950). Australian J. Sci. Res. Ser. B3, 251.
Massey, V. and Singer, T.P. (1957). J. Biol. Chem. 228, 263.
Massini, P. (1957). Acta Botan. Neerl. 6, 434.
Masuda, T. (1955). Phnrm. Bull. (Tokyo) 3, 434.
Masuoka, D.T., Berman, D.A., and Saunders, P.R. (1952). Am. J. Physiol. 170, 301.
Mathews, A. P. (1904). Am. J. Physiol. 10, 290.
Mathews, C.K. and Cohen, S.S. (1963). J. Biol. Chem. 238, 367.
REFERENCES 1037
Matkovics, B., Foldeak, S., and Konsanszky, A. (1961). Enzymologia 23, 314.
Matsuo, Y. and Greenberg, D.M. (1959). J. Biol. Chem. 234, 507.
Matthew, M. and Neuberger, A. (1963). Biochem. J. 87, 601.
Matthews, R.E.F. (1958). Pharmacol. Rev. 10, 359.
Matthews, R.E.F. and Proctor, C.H. (1956). J. Gen. Microbiol. 14, 366.
Mauzerall, D. and Granick, S. (1958). J. Biol. Chem. 232, 1141.
Maver, M.E. and Thompson, J.W. (1946). J. Natl. Cancer Inst. 6, 355.
Maw, G.A. (1959). Biochem. J. 72, 602.
Maxwell, E.S. (1957). J. Biol. Chem. 229, 139.
Maxwell, E.S., Kalckar, H.M., and Strominger, J.L. (1956). Arch. Biochem. Biophys.
65, 2.
Mayer, A.M. (1959). Enzymologia 20, 313.
Mazelis, M. and Crevehng, R.K. (1963). J. Biol. Chem. 238, 3358.
Mazehs, M. and Stumpf, P.K. (1955). Plant Physiol. 30, 237.
Mazia, D. (1959). Harvey Lectures 53, 130.
Mazur, A. (1955). In "Methods in Enzymology" (S.P. Colowick and N.O. Kaplan,
eds.). Vol. 1, p. 651. Academic Press, New York.
Medina, A. and Nicholas, D.J.D. (1957 a). Biochem. J. 66, 573.
Medina, A. and Nicholas, D.J.D. (1957 b). Biochim. Biophys. Acta 25, 138.
Mehl, J.W. (1944). Science 99, 518.
Meier, R. (1926). Biochem. Z. 174, 384.
Meier, R. (1933). Z. Ges. Exptl. Med. 87, 283.
Meister, A. (1950). J. Biol. Chem. 184, 117.
Mello Ayres, G.C. and Lara, F.J.S. (1962). Biochim. Biophys. Acta 62, 435.
Mendel, B. (1937). Am. J. Cancer 30, 549.
Mendel, B. and Strelitz, F. (1937). Nature 140, 771.
Mendelsohn, M.L. (1955). Am. J. Physiol. 180, 599.
Mendez, R. (1946). Science 104, 5.
Mendez, R. and Peralta, B. (1947). J. Pharmacol. Exptl. Therap. 90, 128.
Mendez, R. and Pisanty, J. (1949). Arch. Inst. Cardiol. Mex. 19, 709.
Mendicino, J. and Vasarhely, F. (1963). J. Biol. Chem. 238, 3528.
Meneghetti, E. (1922)'. Biochem. Z. 131, 38.
Menon, G.K.K. and Stern, J.R. (1960). J. Biol. Chem. 235, 3393.
Menon, G.K.K. , Friedman, D.L., and Stern, J.R. (1960). Biochim. Biophys. Acta AA,
375.
Merrett, M.J. and Syrett, P.J. (1960). Physiol. Plantarum 13, 237.
Mescon, H. and Flesch, P. (1952). J. Invest. Dermatol. 18, 261.
Messing, R.A. (1961). Enzymologia 23, 52.
Metzenberg, R.L. (1963). Arch. Biochem. Biophys. 100, 503.
Metzger, R.P., Wilcox, S.S., and Wick, A.N. (1964). J. Biol. Chem. 239, 1769.
Meyer, D.K. (1952). Federation Proc. 11, 107.
Meyer, H. (1960). Arch. Mikrohiol. 37, 18.
Meyer, H., Preiss, B., and Bauer, S. (1960). Biochem. J. 76, 27.
Meyer, J.R. (1948). Science 108, 188.
Meyer, V. and Wachter, W. (1892). Ber. 25, 2632.
Meyerhof, O. and Burk, D. (1928). Z. Physik. Chem. A139, 117.
Michaelis, L. and Rona, P. (1914). Biochem. Z. 60, 62.
Michaelis, L. and Pechstein, H. (1914). Biochem. Z. 60, 79.
1038 REFERENCES
Mickelson, M.N. and Schuler, M.N. (1953). J. Bacteriol. 65, 297.
Middlebrook, M. and Szent-Gyorgyi, A. (1955). Biochim. Biophys. Acta 18, 407.
Mii, S. and Ochoa, S. (1957). Biochim. Biophys. Acta 26, 445.
Mildvan, A.S. and Greville, G.D. (1962). Biochem. J. 82, 22p.
Millar, D.B.S. and Schwert, G.W. (1963). J. Biol. Chem. 238, 3249.
Millbank, J.W. (1957). J. Exptl. Botany 8, 96.
Miller, A. and Waelsch, B. (1957). J. Biol. Chem. 228, 365.
Miller, J.J. and Halpern, C. (1956). Can. J. Microbiol. 2, 519.
Miller, O.N. and Olson, R.E. (1954). Arch. Biochem. Biophys. 50, 257.
Miller, T.B. and Farah, A.E. (1962 a). J. Pharmacol. Exptl. Therap. 135, 102.
Miller, T.B. and Farah, A.E. (1962 b). J. Pharmacol. Exptl. Therap. 136, 10.
Miller, V.L., Klavano, P.A., and Csonka, E. (1960). Toxicol. Appl. Pharmacol. 2, 344.
Miller, V.L., Klavano, P.A., Jerstad, A.C., and Csonka, E. (1961). Toxicol. Appl.
Pharmacol. 3, 459.
Millerd, A. (1951). Proc. Linnean Soc. N. S. Wales 76, 123.
Millington, P.F. and Finean, J.B. (1958). J. Ultrastruct. Res. 2, 215.
Millington, P.F. and Finean, J.B. (1961). J. Ultrastruct. Res. 5, 470.
Mills, G.T., Paul, J., and Smith, E.E.B. (1953). Biochem. J. 53, 232.
Mills, R.R. and Cochran, D.G. (1963). Biochim. Biophys. Acta 73, 213.
Milstein, C. (1961). Biochem. J. 79, 591.
Minakami, S., Cremona, T., Ringler, R.L., and Singer, T.P. (1963). J. Biol. Chem.
238, 1529.
Minatoya, H., Luduena, P.P., and von Euler, L. (1960). Federation Proc. 19, 65.
Mirsky, A.E. (1941). J. Gen. Physiol. 24, 709.
Mirsky, A.E. and Anson, M.L. (1935). J. Gen. Physiol. 18, 307.
Mirsky, A.E. and Anson, M.L. (1936 a). J. Gen. Physiol. 19, 439.
Mirsky, A.E. and Anson, M.L. (1936 b). J. Gen. Physiol. 19, 451.
Misaka, E. and Nakanishi, K. (1963). J. Biochem. {Tokyo) 53, 465.
Misra, U.K. and French, D. (1960). Biochem. J. 77, Ip.
Mitchell, I.L.S. and Bayne, S. (1952). Biochem. J. 50, xxvii.
Mitchell, P. (1953). J. Gen. Microbiol. 9, 273.
Mitchell, P. (1954). Symp. Soc. Exptl. Biol. 8, 254.
Mitchell, R.A., Kang, H.H., and Henderson, L.M. (1963). J. Biol. Chem. 238, 1151.
Mittermayer, C. and Breuer, H. (1963). Biochim. Biophys. Acta 77, 191.
Miura, N. (1934). Japan. J. Med. Sci. IV. Pharmacol. 8, No. 1.
Miyachi, S. (1960). Plant Cell Physiol. (Tokyo) 1, 117.
Mize, C.E. and Langdon, R.G. (1962). J. Biol. Chem. 237, 1589.
Mizushima, S. and Izaki, K. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo Kyoto
1957 p. 307. Maruzen, Tokyo.
Modell, W. and Krop, S. (1944). Proc. Soc. Exptl. Biol. Med. 55, 80.
Modignani, R.L. and Foa, P.P. (1963). Proc. Soc. Exptl. Biol. Med. 112, 106.
Mokrasch, L.C. and McGilvery, R.W. (1956). J. Biol. Chem. 221, 909.
Mohnari, R. and Lara, F.J.S. (1960). Biochem. J. 75, 57.
Molnar, D.M., Burris, R.H., and Wilson, P.W. (1948). J. Am. Chem. Soc. 70, 1713.
Momose, G. (1914). Biochem. Z. 61, 312.
Momose, G. (1925). J. Biochem. (Tokyo) 4, 441.
Monroy, A. and Runnstrom, J. (1952). Exptl. Cell. Res. 3, 10.
Montgomery, CM. and Bamberger, J.W. (1955). Science 122, 967.
REFERENCES 1039
Montgomery, CM. and Webb, J.L. (1956 a). J. Biol. Chem. 221, 347.
Montgomery, CM. and Webb, J.L. (1956 b). Unpublished work.
Mookerjea, S. and Sadhu, D.P. (1955). Arch. Biochem. Biophys. 58, 232.
Moore, B.W. (1959). Arch. Biochem. Biophys. 80, 458.
Moos, C, Alpert, N.R., and Myers, T.C (1960). Arch. Biochem. Biophys. 88, 183.
Mora, P.T. (1962). J. Biol. Chem. 237, 3210.
Mora, P.T. and Young, E.G. (1959). Arch. Biochem. Biophys. 82, 6.
Morawetz, H. and Sage, H. (1955). Arch. Biochem. Biophys. 56, 103.
Morgan, E.J. and Friedmann, E. (1938 a). Biochem. J. 32, 733.
Morgan, E.J. and Friedmann, E. (1938 b). Biochem. J. 32, 862.
Morgan, E.J. and Friedmann, E. (1938 c). Biochem. J. 32, 2296.
Morgan, H.E. and Park, CR. (1958). Federation Proc. 17, 278.
Morgan, H.E., Henderson, M.J., Regen, D.M., and Park, CR. (1961 a). J. Biol. Chem.
236, 253.
Morgan, H.E., Cadenas, E., Regen, D.M., and Park, CR. (1961b). J. Biol. Chem. 236,
262.
Morgan, H.R. (1954). J. Exptl. Med. 99, 451.
Morihara, K. (1963). Biochim. Biophys. Acta 73, 113.
Morino, Y., Wada, H., Morisue, T., Sakamoto, Y., and Ichihara, K. (1960). J. Bio-
chem. {Tokyo) 48, 18.
Morisue, T., Morino, Y., Sakamoto, Y., and Ichihara, K. (1960). J. Biochem. {Tokyo)
48, 28.
Moriyama, H. and Ohashi, S. (1941). Arch. Ges. Virusforsch. 2, 205.
Morris, M.P., Pagan, C, and Warmke, H.E. (1954). Science 119, 322.
Morrison, J.F. (1950). Australian J. Exptl. Biol. Med. Sci. 28, 311.
Morrison, J.F., Griffiths, D.E., and Ennor, A.H. (1957). Biochem. J. 65, 143.
Morton, H.E., Kocholaty, W., Junowicz-Kocholaty, R., and Kelner, A. (1945). J.
Bacterial. 50, 579.
Morton, R.K. (1955). Biochem. J. 61, 232.
Moses, V. (1955). J. Gen. Microbiol. 13, 235.
Moulder, J.W., McCormack, B.R.S., and Itatani, M.K. (1953). J. Infect. Diseases 93,
140.
Moulder, J.W., Novosel, D.L., and Officer, J.E. (1963). J. Bacteriol. 85, 707.
Mounter, L.A. (1954). J. Biol. Chem. 290, 813.
Mounter, L.A. and Chanutin, A. (1953). J. Biol. Chem. 204, 837.
Mounter, L.A. and Whittaker, V.P. (1953). Biochem. J. 53, 167.
Moussatche, H. and Prouvost-Danon, A. (1957). Naturwissenschaften 44, 637.
Moussatche, H. and Prouvost-Danon, A. (1958). Arch. Biochem. Biophys. 77, 108.
Moyed, H.S. (1960). J. Biol. Chem. 235, 1098.
Moyed, H.S. and O'Kane, D.J. (1956). J. Biol. Chem. 218, 831.
Moyer, J.H., Handley, C.A., and Seibert, R.A. (1957). Am. N. Y. Acad. Sci. 65, 511.
Mozen, M.M. (1955). Ph. D. Thesis, University of Wisconsin, Madison.
Mozen, M.M., Burris, R.H., Lundbom, S., and Virtanen, A.I. (1955). Acta Chem.
Scand. 9, 1232.
Mudd, J.B. and McManus, T.T. (1962). J. Biol. Chem. 237, 2057.
Mudd, J.B. and McManus, T.T. (1964). Plant Physiol. 39, 115.
Mudd, J.B. and Stumpf, P.K. (1961). J. Biol. Chem. 236, 2602.
Mudge, G.H. (1951). Am. J. Physiol. 167, 206.
1040 REFERENCES
Mudge, G.H. and Weiner, I.M. (1958). Ann. N. Y. Acad. Sci. 71, 344.
Mueller, J.F. and Vilter, R.W. (1950). J. Clin. Invest. 29, 193.
Mueller, J.F., lacono, J., Will. J.J., and Repasky, W. (1959). J. Lab. Clin. Med. 54,
929.
Muir, C, de Kock, L.L., de Kock, P.C., and Inkson, R.H.E. (1959). Experientia 15,
354.
Miiller, A.F. and Leuthardt, F. (1950). Helv. Chim. Acta 33, 268.
Muller, F., Schoeller, W., and Schrauth, W. (1911). Biochem. Z. 33, 381.
Miiller, G., Iwainsky, H., and Taufel, K. (1960). Z. Naturforsch. 15b, 180.
Miiller, E. and Miiller-Rodloff, I. (1935). Ann. Chem. 521, 81.
Mullins, L.J. (1960). J. Gen. Physiol. 43, Suppl. 1, 105.
Murachi, T. and Neurath, H. (1960). J. Biol. Chem. 235, 99.
Murachi, T. and Tashiro, M. (1958). Biochim.. Biophys. Acta 29, 645.
Murayaraa, M. (1958). J. Biol. Chem. 230, 163.
Murray, D.R.P. (1929). Biochem. J. 23, 292.
Murray, D.R.P. and King, C.G. (1930). Biochem. J. lA, 190.
Muscholl, E. (1958). Arch. Exptl. Pathol. Pharmakol. 235, 23.
Muscholl, E. and Maitre, L. (1963). Experientia 19, 658.
Mushett, C.W., Stebbins, R.B., and Barton, M.N. (1947). Am. J. Med. Sci. 213, 509.
Mustakallio, K.K. and Telkka, A. (1953). Science 118, 320.
Muus, J., Brockett, F.P., and Connelly, C.C. (1956). Arch. Biochem. Biophys. 65, 268.
Myers, D.K. and Slater, E.G. (1957 a). Nature 179, 363.
Myers, D.K. and Slater, E.G. (1957 b). Biochem. J. 67, 572.
Myrback, K. (1926). Z. Physiol. Chem. 158, 160.
Myrback, K. (1957 a). Arch. Biochem. Biophys. 69, 138.
Myrback, K. (1957 b). Festschr. Arthur Stoll p. 551.
Myrback, K. and Persson, B. (1953 a). Arkiv Kemi 5, 177.
Myrback, K. and Persson, B. (1953 b). Arkiv Kemi 5, 477.
Naber, E.G., Cravens, W.W., Baumann, C.A., and Bird, H.R. (1954). J. Nutr. 54, 579.
Nachlas, M.M., Margulies, S.I., and Seligman, A.M. (1960). J. Biol. Chem. 235, 499.
Nachmansohn, D. and Lederer, E. (1939). Bull. Soc. Chim. Biol. 21, 797.
Nachmansohn, D. and Machado, A.L. (1943). J. Neurophysiol. 6, 397.
Nadai, Y. (1958). J. Biochem. (Tokyo) 45, 1011.
Nadkarni, S.R. and Sohonie, K. (1963). Enzymologia 25, 337.
Naganna, B. and Menon, V.K.N. (1948). J. Biol. Chem.. 174, 501.
Naik, K.G. and Patel, R.P. (1932). J. Indian Chem. Soc. 9, 533.
Nair, P.M. and Vining, L.C. (1964). Arch. Biochem. Biophys. 106, 422.
Nakada, H.I. (1964). J. Biol. Chem. 239, 468.
Nakada, H.I. and Weinhouse, S. (1950). J. Biol. Chem. 187, 663.
Nakada, H.I. and Wick, A.N. (1956). J. Biol. Chem. Ill, 671.
Nakada, H.I., Wolfe, J.B., and Wick, A.N. (1957). J. Biol. Chem. lib, 145.
Nakamura, H. and Greenberg, D.M. (1961). Arch. Biochem. Biophys. 93, 153.
Nakamura, K. (1960). Biochim. Biophys. Acta 45, 554.
Nakamura, K. and Bernheim, F. (1961). Biochim.. Biophys. Acta 50, 147.
Nakamura, M. (1954). J. Biochem. [Tokyo) 41, 67.
Nakamura, M. and Baker, E.E. (1956). Am. J. Hyg. 64, 12.
Nakata, H.M. and Halvorson, H.O. (1960). J. Bacteriol. 80, 801.
REFERENCES 1041
Nakayama, T. (1959). J. Biochem. {Tokyo) 46, 1217.
Nakayama, T. (1961a). J. Biochem. (Tokyo) 49, 158.
Nakayama, T. (1961b). J. Biochem. (Tokyo) 49, 240.
Nanninga, L.B. (1958). Pacific Slope Biochem. Conf. (Univ. Calif., Los Angeles) 48.
Naoi-Tada, M., Sato-Asano, K., and Egami, F. (1959). J. Biochem. (Tokyo) 46, 757.
Narrod, S.A. and Jakoby, W.B. (1964). J. Biol. Chem. 239, 2189.
Nath, R. and Greenberg, D.M. (1962). Biochemistry 1, 435.
Nathans, D., Tapley, D.F., and Ross, J.E. (1960). Biochim. Biophys. Acta 41, 271.
Navazio, F., Siliprandi, N., and Rossi-Fanelli, A. (1956). Biochim. Biophys. Acta 19,
274.
Neame, K.D. (1961). Nature 192, 173.
Neame, K.D. (1964). J. Neurochem. 11, 67.
Neilands, J.B. (1952). J. Biol. Chem. 199, 373.
Neilands, J.B. (1954). J. Biol. Chem. 208, 225.
Neims, A.H. and Hellerman, L. (1962). J. Biol. Chem. 237, PC976.
Nelson, C.A. and Hummel, J.P. (1961). J. Biol. Chem. 236, 3173.
Nelson, E.K. and Hasselbring, H. (1931). J. Am. Chem. Soc. 53, 1040.
Nelson, L. (1959). Biol. Bull. 117, 327.
Nelson, M. (1964). Comp. Biochem. Physiol. 12, 37.
Nelson, M.M. (1955). In "Antimetabolites and Cancer" (C.P. Rhoads, ed.), p. 107.
Am. Assoc. Advance. Sci., Washington, D.C.
Neufeld, E.F., Kaplan, N.O., and Colowick, S.P. (1955). Biochim. Biophys. Acta 17,
525.
Neuhaus, F.C. and Byrne, W.L. (1959). J. Biol. Chem. 234, 113.
Neuhaus, F.C. and Byrne, W.L. (1960). J. Biol. Chem. 235, 2019.
Neuhaus, F.C. and Lynch, J.L. (1964). Biochemistry 3, 471.
Neuhaus, H. (1910). Arch. Intern. Pharmacodyn. 20, 393.
Neurath, H. and Gladner, J.A. (1951). J. Biol. Chem. 188, 407.
Neurath, H. and Schwert, G.W. (1950). Chem. Rev. 46, 69.
Newey, H. and Smyth, D.H. (1964). J. Physiol. (London) 170, 328.
Newmark, M.Z. and Wenger, B.S. (1960). Arch. Biochem. Biophys. 89, 110.
Ngai, S.H., Ginsburg, S., and Katz, R.L. (1961). Biochem. Pharmacol. 7, 256.
Nichol. C.A. (1954). Cancer Res. 14, 522.
Nichol, C.A. and Welch, A.D. (1950). Proc. Soc. Exptl. Biol. Med. 74, 403.
Nicholas, D.J.D., Medina, A., and Jones, O.T.G. (1960). Biochim. Biophys. Acta 37,
468.
Nickerson, M. and Nomaguchi, G.M. (1950). Am. J. Physiol. 163, 484.
Nickerson, W.J. and Chadwick, J.B. (1946). Arch. Biochem. 10, 81.
Niedergang-Kamien, E. and Leopold, A.C. (1957). Physiol. Plantarum 10, 29.
Niederpruem, D.J. and Hackett, D.P. (1961). Plant Physiol. 36, 79.
Niemer, H. and Kohler, A. (1957). Z. Physiol. Chem. 308, 58.
Nigam, V.N., Davidson, H.M., and Fishman, W.H. (1959). J. Biol. Chem. 234, 1550.
Nihei, T., Noda, L., and Morales, M.F. (1961). J. Biol. Chem. 236, 3203.
Nimmo-Smith, R.H. and Standen, O.D., (1963). Exptl. Parasitol. 13, 305.
Ninomiya, H. and Suzuoki, Z. (1952). J. Biochem. (Tokyo) 39, 321.
Nirenberg, M.W. and Hogg, J.F. (1957). Federation Proc. 16, 227.
Nu-enberg, M.W. and Hogg, J.F. (1958). Cancer Res. 18, 518.
Nishi, A. (1958). J. Biochem. (Tokyo) 45, 991.
1042 REFERENCES
Nishi, A. (1960). J. Biochem. (Tokyo) 47, 47.
Nishimura, J.S. and Greenberg, D.M. (1961). J. Biol. Chem. 236, 2684.
Nishimura, S. (1960). Biochim. Biophys. Acta 45, 15.
Nistratova, S.N. and Turpaev, T.M. (1959). Biochemistry (USSR) (English Transl.)
lA, 155.
Noda, H. and Maruyama, K. (1960). Biochim. Biophys. Acta 41, 393.
Noda, L. (1958). J. Biol. Chem. 1^1, 237.
Noda, L., Nihei, T., and Morales, M.F. (1960). J. Biol. Chem. 235, 2830.
Noe, F.E. (1960). Ind. Med. Surg. 29, 559.
Noguchi, H. and Akatsu, S. (1917). J. Pharmacol. Exptl. Therap. 9, 363.
Nohara, H. and Ogata, K. (1959). Biochim. Biophys. Acta 31, 142.
Noltmann, E. and Bruns, F.H. (1959). Biochem. Z. 331, 436.
Nordlie, R.C. and Lardy, H.A. (1961). Biochim. Biophys. Acta 53, 309.
Nosoh, Y. (1961). J. Biochem. (Tokyo) 50, 450.
Nossal, P.M. (1952). Biochem. J. 50, 349.
Novikoff, A.B., Hecht, L., Podber, E., and Ryan, J. (1952). J. Biol. Chem. 194, 153.
Novoa, W.B. and Schwert, G.W. (1961). J. Biol. Chem. 236, 2150.
'^fovoa, W.B., Winer, A.D., Glaid, A. J., and Schwert, G.W. (1959). J. Biol. Chem. 234,
1143.
Nukada, T., Matsuoka, M., and Imaizumi, R. (1962). Japan. J. Pharmacol. 12, 57.
Nygaard, A.P. (1955). Acta Chem. Scand. 9, 1048.
Nygaard, A.P. (1956). Acta Chem. Scand. 10, 397.
Nygaard, A.P. (1961a). Ann. N. Y. Acad. Sci. 94, 774.
Nygaard, A.P. (1961b). J. Biol. Chem. 236, 920.
Nygaard, A.P. (1961c). J. Biol. Chem. 236, 2779.
Nyhan, W.L. and Busch, H. (1957). Cancer Pes. 17, 227.
Gates, J.A., Gillespie, L., Udenfriend, S., and Sjoerdsma, A. (1961). Science 131, 1890.
Ochoa, S. (1944). J. Biol. Chem. 155, 87.
Ochoa, S. and Weisz-Tabori, E. (1948). J. Biol. Chem. 174, 123.
O'Connor, R.J. and Halvorson, H.O. (1960). Arch. Biochem. Biophys. 91, 290.
O'Connor, R.J. and Halvorson, H.O. (1961a). Biochim. Biophys. Acta 48, 47.
O'Connor, R.J. and Halvorson, H.O. (1961b). J. Bacterial. 82, 706.
Ogata, K., Ogata,"" M., Mochizuki, Y., and Nishiyama, T. (1956). J. Biochem. (Tokyo)
43, 653.
Ogilvie, R.F. (1932). J. Pathol. Bacteriol. 35, 743.
Oginsky, E.L. and Gehrig, R.F. (1952). J. Biol. Chem. 198, 799.
Ohno, M. and Okunuki, K. (1962). J. Biochem. (Tokyo) 51, 313.
Ohr, E.A. (1960). Federation Proc. 19, 248.
Ohta, J. (1954). J. Biochem. (Tokyo) 41, 489.
Okazaki, R. and Romberg, A. (1964). J. Biol. Chem. 239, 275.
Okunuki, K. (1939). Acta Phijtochim. (Japan) 11, 27.
Okunuki, K. and Sekuzu, I. (1955). J. Biochem. (Tokyo) 42, 397.
Oliphant, J.F. (1942). Physiol. Zool. 15, 443.
Olivard, J. and Snell, E.E. (1955). J. Biol. Chem. 213, 203.
OHver, I.T. (1961). Biochim. Biophys. Acta 52, 75.
Olson, E.J. and Park, J.H. (1964). J. Biol. Chem. 239, 2316.
Olson, J. A. (1959). J. Biol. Chem. 234, 5.
REFERENCES 1043
Olson, J. A. and Anfinsen, C.B. (1953). J. Biol. Chem. 202, 841.
Olson, J.M., Duysens, L.N.M., and Kronenberg, G.H.M. (1959). Biochim. Biophys.
Acta 36, 125.
Onrust, H., Van der Linden, A.C., and Jansen, B.C.P. (1952). Enzymologia 15, 351.
Onrust, H., Van der Linden, A.C., and Jansen, B.C.P. (1954). Enzymologia 16, 289.
Ordin, L. and Jacobson, L. (1955). Plant Physiol. 30, 21.
Oren, R., Farnham, A.E., Saito, K., Milofsky, E., and Karnovsky, M.L. (1963). J.
Cell Biol. 17, 487.
Orloff, J. and Berliner, R.W. (1961). Ann. Rev. Pharmacol. 1, 287.
Ornstein, N. and Gregg, J.R. (1952). Biol. Bull. 103, 407.
Orten, J.M. and Smith, A.H. (1937). J. Biol. Chem. 117, 555.
Osborn, M.J., Freeman, M., and Huennekens, F.M. (1958). Proc. Soc. Exptl. Biol. Med.
97, 429.
Ota, S., Wachi, T., Uchida, M., Sakamoto, Y., and Ichihara, K. (1956). J. Biochem,.
{Tokyo) 43, 611.
Ota, S., Fu, T.-H., and Hirohata, R. (1961). J. Biochem. (Tokyo) 49, 532.
Otsuka, S. (1961). J. Biochem. {Tokyo) 50, 85.
Ott, J.L. (1960). J. Bacteriol. 80, 355.
Ott, W.H. (1946). Proc. Soc. Exptl. Biol. Med. 61, 125.
Ott, W.H. (1947). Proc. Soc. Exptl. Biol. Med. 66, 215.
Ottolenghi, P. and Denstedt, O.F. (1958). Can. J. Biochem. Physiol. 36, 1075.
Overell, B.T. (1952). Australian J. Sci. 15, 28.
Owens, H.S., Goodban, A.E.,'and Stark, J.B. (1953). Anal. Chem. 25, 1507.
Owens, R.G. (1953 a). Contrib. Boyce Thompson Inst. 17, 221.
Owens, R.G. (1953 b). Contrib. Boyce Thompson Inst. 17, 273.
Oxender, D.L. and Christensen, H.N. (1963). J. Biol. Chem. 238, 3686.
Ozaki, K. and Wetter, L.R. (1960). Can. J. Biochem. Physiol. 38, 125.
Ozaki, K. and Wetter, L.R. (1961). Can. J. Biochem. Physiol. 39, 843.
Packer, L. (1958). Exptl. Cell. Res. 15, 551.
Packer, L., Papa, M.J., Rust, J.H., Jr., and Ajl, S.J. (1959). Biochim. Biophys. Acta
35, 354.
Padykula, H.A. and Herman, E. (1955). J. Histochem. Cytochem. 3, 170.
PaflF, G.H., Sugiura, H.T., Bocher, C.A., and Roth, J.S. (1952). Anat. Record 114, 499.
Palmer, G. (1962). Biochim. Biophys. Acta 64, 135.
Palmer, G. and Massey, V. (1962). Biochim. Biophys. Acta 58, 349.
Pantlitschko, M. and Kaiser, E. (1951). Biochem. Z. 322, 137.
Pantlitschko, M. and Seelich, F. (1960). Biochem. Z. 333, 78.
/)Papaconstantinou, J. and Colowick, S.P. (1957). Federation Proc. 16, 230.
^vPapaconstantinou, J. and Colowick, S.P. (1961a). J. Biol. Chem. 236, 278.
Papaconstantinou, J. and Colowick, S.P. (1961b). J. Biol. Chem. 236, 285.
Pardee, A.B. and Potter, V.R. (1948). J. Biol. Chem. 176, 1085.
Pardee, A.B. and Potter, V.R. (1949). J. Biol. Chem. 178, 241.
Pardee, A.B. and Prestidge, L.S. (1958). Biochim. Biophys. Acta TJ, 330.
Park, R.B., Pon, N.G., Louwrier, K.P., and Calvin, M. (1960). Biochim. Biophys. Acta
Al, 27.
Parker, A.J. and Kharasch, N. (1959). Chem. Rev. 59, 583.
Parker, C.A. (1954). Nature 173, 780.
1044 REFERENCES
Parker, C.A. and Dilworth, M.J. (1963). Biochim. Biophys. Acta 69, 152.
Parker, C.A. and Scutt, P.B. (1958). Biochim. Biophys. Acta 29, 662.
Parker, C.A. and Scutt, P.B. (1960). Biochim. Biophys. Acta 38, 230.
Parks, R.E., Jr., Ben-Gershom, R., and Lardy, H.A. (1957). J. Biol. Chem. 227, 231.
Parr, C.W. (1956). Nature 178, 1401.
Parr, C.W. (1957). Biochem. J. 65, 34p.
Partridge, A.D. and Rich, A.E. (1962). Phytopathology 52, 1000.
Passonneau, J.V. and Lowtv, O.H. (1963). Biochem. Biophys. Res. Commun. 13, 372.
Passow, H. and Rothstein, A. (1960). J. Gen. Physiol. 43, 621.
Pasternak, C.A. and Handschumacher, R.E. (1959). J. Biol. Chem. 234, 2992.
Pattabiraman, T.N. and Bachhawat, B.K. (1961). Biochim. Biophys. Acta 54, 273.
Patterson, W.B. and Stetten, D., Jr. (1949). Science 109, 256.
Pauling, L. (1946). Chem. Eng. News 24, 1375.
Pauling, L. (1948). Nature 161, 707.
Payes, B. and Laties, G.G. (1963). Biochem. Biophys. Res. Commun. 10, 460.
Pazur, J.H. and Kleppe, K. (1964). Biochemistry 3, 578.
Pearson, D.E. and Levine, M. (1952). J. Org. Chem. 17, 1351.
Pearson, R.G. and Mills, J.M. (1950). J. Am. Chem. Sac. 72, 1692.
Pease, D.C. (1941). J. Exptl. Zool. 86, 381.
Peck, H.D., Jr., Smith, O.H., and Gest, H. (1957). Biochim. Biophys. Acta 25, 142.
Pedersen, T.A. (1963). Physiol. Plantarum 16, 167.
Penefsky, H.S., Pullman, M.E., Datta, A., and Racker, E. (1960). J. Biol. Chem. 235,
3330.
Penn, N. and Mackler, B. (1958). Biochim. Biophys. Acta 27, 539.
Pennington, R.J. (1956). Biocheyn. J. 64, 6p.
Pennington, R.J. (1957). Biochem. J. 65, 534.
Pennington, R.J. (1961). Biochem. J. 80, 649.
Pennington, R.J. and Appleton, J.M. (1958). Biochem. J. 69, 119.
Pentschew, A. and Kassowitz, H. (1932). Arch. Exptl. Pathol. Pharmakol. 164, 667.
Pereira, A.S.R. and Linskens, H.F. (1963), Acta Botan. Neerl. 12, 302.
Perez, J.E., Baralt-Perez, J., and Klein, M. (1949). J. Immunol. 62, 405.
Perkins, D.J. (1952). Biochem. J. 51, 487.
Perkins, D.J. (1953). Biochem. J. 55, 649.
Perkins, D.J. (1958). Biochem. J. 69, 35p.
Perkins, D.J. (1961). Biochem. J. 80, 668.
Perry, J.J. and Evans, J.B. (1960). J. BacteHol. 79, 113.
Pershin, G.N. and Shcherbakova, L.I. (1958). Biochemistry {USSR) {English Transl.)
21, 155.
Person, P. and Fine, A. (1959). Arch. Biochem. Biophys. 84, 123.
Petering, H.G. and Schmitt, J. A. (1950). J. Am. Chem. Soc. 72, 2995.
Peters, J.M. and Greenberg, D.M. (1959). Biochim. Biophys. Acta 32, 273.
Peters, R.A. and Wakelin, R.W. (1946). Biochem. J. 40, 513.
Peterson, E.A. and Greenberg, D.M. (1952). J. Biol. Chem. 194, 359.
Petrucci, D. (1954 a). Riv. Biol. {Perugia) 46, 241.
Petrucci, D. (1954 b). Boll. Soc. Ital. Biol. Sper. 30, 282.
Pfleiderer, G., Hohnholz-Merz, E., and Gerlach, D. (1962). Biochem. Z. 336, 371.
Phares, E.F., Mosbach, E.H., Denison, F.W., Jr., Carson, S.F., Long, M.V,, and Gwin,
B.A. (1952). Anal. Chem. 24, 660.
REFERENCES 1045
Philips, F.S., Oilman, A., Koelle, E.S., and Allen, R.P. (1947). J. Biol. Chem. 167,
209.
Phillips, A.H. and Langdon, R.G. (1962). J. Biol. Chem. 237, 2652.
Philpot, J.St.L. and Small, P.A. (1938). Biochem. J. 32, 542.
Pierpoint, W.S. (1959). Biochem. J. 71, 518.
Pietschmann, H., Lachnit, V., and Reinhardt, F. (1962). Z. Ges. Exptl. Med. 136,
108.
Pihl, A. and Lange, R. (1962). J. Biol. Chem. 237, 1356.
Pihl, A., Lange, R., and Evang, A. (1961). Acta Chem. Scand. 15, 1271.
Piloty, 0. and Schwerin, E.G. (1901a). Ber. 34, 1863.
Piloty, O. and Schwerin, E.G. (1901b). Ber. 34, 1870.
Piloty, O. and Schwerin, E.G. (1901c). Ber. 34, 2354.
Piloty, O. and Vogel, W. (1903). Ber. 36, 1283.
Pine, M.J. (1960). J. Bacteriol. 79, 827.
Pisanty, J. (1948). Arch. Inst. Cardiol. Mex. 18, 111.
Pitts, R.F. (1958). Am. J. Med. 24, 745.
Pitts, R.F. (1959). "The Physiological Easis of Diuretic Therapy." Thomas, Spring-
field, Illinois.
Plant, G.W.E. (1957). Arch. Biochem. Biophys. 69, 320.
Plant, G.W.E. and Lardy, H.A. (1949). J. Biol. Chem. 180, 13.
Pletscher, A., Burkard, W.P., and Gey, K.F. (1964). Biochem. Pharmacol. 13, 385.
^Plummer, D.T. and Wilkinson, J.H. (1963). Biochem. J. 87, 423.
Pogell, E.M. (1958). J. Biol. Chem. 131, 761.
Pogrund, R.S. and Clark, W.G. (1956). Federation Proc. 15, 145.
Pohl, J. (1896). Arch. Exptl. Pathol. Pharmakol. 37, 413.
Pohle, W. and Matthies, H. (1959). Arch. Exptl. Pathol. Phermakol. 236, 253.
Polatnick, J. and Bachrach, H.L. (1960). Proc. Soc. Exptl. Biol. Med. 105, 601.
Polls, E.D. and Meyerhof, O. (1947). J. Biol. Chem. 169, 389.
Pollock, M.R. and Knox, R. (1943). Biochem. J. 37, 476.
Polonovski, M., Schapira, G., and Gonnard, P. (1946). Bull. Soc. Chim. Biol. 28,
735.
Polyanovskii, O.L. (1962). Biochemistry (USSR) (English Transl.) 11, 623.
Ponz, F. and Llinas, J.M. (1963). Nature 197, 696.
Popjak, G. and Tietz, A. (1955). Biochem. J. 60, 147.
Popjak, G., Gosselin, L., Gore, I.Y., and Gould, R.G. (1958). Biochem. J. 69, 238.
Porter, C.C. and Hellerman, L. (1939). J. Am. Chem. Soc. 61, 754.
Porter, C.C. and Hellerman, L. (1944). J. Am. Chem. Soc. 66, 1652.
Porter, C.C, Clark, I., and Silber, R.H. (1947). J. Biol. Chem. 167, 573.
Porter, C.C, Totaro, J.A., and Leiby, CM. (1961). J. Pharmacol. Exptl. Therap. 134,
139.
Portzehl, H. (1952). Z. Naturforsch. 7b, 1.
Post, R.L., Morgan, H.E., and Park, C.R. (1961). J. Biol. Chem. 236, 269.
Postgate, J.R. (1956). J. Gen. Microbiol. 14, 545.
Potter, V.R. (1940). J. Biol. Chem. 134, 417.
Potter, V.R. (1951). Proc. Soc. Exptl. Biol. Med. 76, 41.
Potter, V.R. and DuEois, K.P. (1943). J. Gen. Physiol. 26, 391.
Potter, V.R. and Elvehjem, CA. (1937). J. Biol. Chem. 117, 341.
Pott«r, V.R. and Schneider, W.C (1942). J. Biol. Chem. 142, 543.
1046 REFERENCES
Potter, V.R., Busch, H., and Reif, A.E. (1952). Proc. 2nd Natl. Cancer Conf. p. 1294.
Price, C.A. (1953). Arch. Biochem. Biophys. 47, 314.
Pubols, M.H., Zhanley, J.C, and Axelrod, B. (1963). Plant Physiol. 38, 457.
Pugh, D., Leaback, D.H., and Walker, P.G. (1957). Biochem. J. 65, 464.
PuUman, M.E., Penefsky, H.S., Datta, A., and Racker, E. (1960). J. Biol. Chem. 235,
3322.
Pupilli, G. (1930). Biochim. Terap. Sper. 17, 113.
Purpura, D.P., Berl, S., Gonzalez-Monteagudo, O., and Wyatt, A. (1960). In "Inhi-
bition in the Nervous System and y-Aminobutyric Acid" (E. Roberts, ed.), p. 331.
Macmillan (Pergamon), New York.
Putnam, F.W. (1953). In "The Proteins" (H. Neurath and K. Bailey, eds.), Vol. 1,
Pt. B, p. 807. Academic Press, New York.
Putter, J. (1961). Z. Krehsforsch. 64, 101.
Pyefinch, K.A. and Mott, J.C. (1948). J. Exptl. Biol. 25, 276.
Quastel, J.H. and Bickis, I.J. (1959). Nature 183, 281.
Quastel, J.H. and Cantero, A. (1953). Nature 171, 252.
Quastel, J.H. and Scholefield, P.G. (1955). J. Biol. Chem. 214, 245.
Quastel, J.H. and Wheatley, A.H.M. (1931). Biochem. J. 25, 117.
Quastel, J.H. and Wheatley, A.H.M. (1935). Biochem. J. 29, 2773.
Quastel, J.H. and Whetham, M.D. (1925). Biochem. J. 19, 520.
Quastel, J.H. and Wooldridge, W.R. (1928). Biochem. J. 11, 689.
Quesnel, V.C.J. (1956). Exptl. Cell. Res. 10, 575.
Quinn, J.R. and Pearson, A.M. (1964). Nature 201, 928.
Raaflaub, J. (1953). Helv. Physiol. Pharmacol. Acta 11, 142.
Rabin, R.S. and Klein, R.M. (1957). Arch. Biochem. Biophys. 70, 11.
Rabinovitch, M. and Barron, E.S.G. (1955). Biochim. Biophys. Acta 18, 316.
Rabinovitch, M. and Dohi, S.R. (1957). Arch. Biochem. Biophys. 70, 239.
Rabinovitz, M. and McGrath, H. (1959). J. Biol. Chem. 234, 2091.
Rabinovitz, M., Olson, M.E., and Greenberg, D.M. (1954). J. Biol. Chem. 210, 837.
Rabinovitz, M., Olson, M.E., and Greenberg, D.M. (1957). Cancer Res. 17, 885.
Rabinowitz, J.C. and Snell, E.E. (1953 a). Arch. Biochem. Biophys. 43, 399.
Rabinowitz, J.C. and Snell, E.E. (1953 b). Arch. Biochem. Biophys. 43, 408.
Racker, E. and Schroeder, E.A.R. (1958). Arch. Biochem. Biophys. 74, 326.
Racker, E., Klybas, V., and Schramm, M. (1959). J. Biol. Chem. 234, 2510.
Rafter, G.W. (1957). Arch. Biochem. Biophys. 67, 267.
Rafter, G.W. (1958). J. Biol. Chem. 230, 643.
Rafter, G.W. and Colowick, S.P. (1957). Arch. Biochem. Biophys. 66, 190.
Raghupathy, E., Moudgal, N.R., and Sarma, P.S. (1958). Enzymologia 19, 303.
Ragland, J.B. and Liverman, J.L. (1958). Arch. Biochem. Biophys. 76, 496.
Rahatekar, H.I. and Rao, M.R.R. (1963). Enzymologia 25, 292.
Rahn, O. and Barnes, M.N. (1933). J. Gen. Physiol. 16, 579.
Raiziss, G.W., Severac, M., and Moetsch, J.C. (1926). J. Pharmacol. Exptl. Therap.
26, 447.
Rajagopalan, K.V. and Handler, P. (1964). J. Biol. Chem. 239, 2027.
Rajagopalan, K.V., Sundaram, T.K., and Sarma, P.S. (1960). Biochem.. J. 74, 355.
Rajagopalan, K.V., Fridovich, I., and Handler, P. (1962). J. Biol. Chem. 237, 922.
REFERENCES 1047
Rail, J.E., Roche, J., Michel, R., Michel, O., and Varrone, S. (1962). Biochem. Biophys.
Res. Commun. 7, 111.
Ram, D., Kalner, H.S., and Bloch-Frankenthal, L. (1963). Cancer Res. 23, 600.
Ramachandran, S. and Gottlieb, D. (1963). Biochim. Biophys. Acta 69, 74.
Ramakrishnan, C.V. Steel, R., and Lentz, C.P. (1955). Arch. Biochem. Biophys. 55,
270.
Ramasarma, T. and Giri, K.V. (1956). Arch. Biochem. Biophys. 62, 91.
Ranzi, S. (1955). "Symposium on the Fine Structure of Cells," p. 229. Wiley (Inter-
science), New York.
Rao, D.R. and Oesper, P. (1961). Biochem. J. 81, 405.
Rao, M.R.R. and Altekar, W.W. (1961). Biochem. Biophys. Res. Commun. 4, 101.
Rao, M.R.R. and Gunsalus, I.C. (1955). Federation Proc. 14, 267.
Rao, N.A., Cama, H.R., Kumar, S.A., and Vaidyanathan, C.S. (1960). J. Biol. Chem.
235, 3353.
Rapkine, L. (1938). Biochem. J. 32, 1729.
Rapoport, S. and Luebering, J. (1951). J. Biol. Chem. 189, 683.
Rapoport, S., Luebering, J., and Wagner, R.H. (1955). Z. Physiol. Chem. 302, 105.
Rapp, G.W. and Sliwinski, R.A. (1956). Arch. Biochem. Biophys. 60, 379.
Raska, S.B. (1946). J. Am. Med. Assoc. 131, 1093.
Rassweiler, C.F. and Adams, R. (1924). J. Am. Chem. Sac. 46, 2758. \
Rathbun, R.C. and Shideman, F.E. (1951). Federation Proc. 10, 330.
Ratner, S. (1955). In "Amino Acid Metabolism" (E.D. McElroy and H.B. Glass, eds.),
p. 231. Johns Hopkins Press, Baltimore, Maryland.
Ratner, S. and Pappas, A. (1949). J. Biol. Chem. 179, 1199.
Raw, I., Nogueira, O.C, and Filho, J.B.M. (1961). Enzymologia 23, 123.
Razzell, W.E. (1961). J. Biol. Chem. 236, 3031.
Razzell, W.E. and Khorana, H.G. (1959). J. Biol. Chem. 234, 2105.
Read, C.P. (1952). Exptl. Parasitol 1, 353.
Read, C.P. (1956). Exptl. Parasitol. 5, 325.
Recknagel, R.O. and Potter, V.R. (1951). J. Biol. Chem. 191, 263.
Redetzki, H.M. and Alvarez-O'Bourke, F. (1962). J. Pharmacol. Exptl. Therap. 1 37, 173.
Redfearn, E.R. (1960). Ciha Found. Symp., Quinones Electron Transport p. 346.
Rees, K.R. (1953). Biochem. J. 55, 478.
Rees, K.R. (1954). Biochem. J. 58, 196.
Reichard, P. (1957). Acta Chem. Scand. 11, 523.
Reichard, P. and HanshofF, G. (1956). Acta Chem.. Scand. 10, 548.
Reichmann, M.E. and Hatt, D.L. (1961). Biochim. Biophys. Acta 49, 153.
Reinbothe, I.H. (1957). Pharmazie 12, 732.
Reiner, L. and Gellhorn, A. (1956). J. Pharmacol. Exptl. Therap. 117, 52.
Reisberg, R.B. (1954). Biochim. Biophys. Acta 14, 442.
Reiss, O.K. and Hellerman, L. (1958). J. Biol. Chem. 231, 557.
Remy, C.N. (1961). J. Biol. Chem. 236, 2999.
Rendina, G. and Mills, R.C. (1957). J. Bacteriol. 74, 456.
Rennels, E.G. and Ruskin, A. (1954). Proc. Soc. Exptl. Biol. Med. 85, 309.
Repaske, R. (1954). J. Bacteriol. 68, 555.
Repaske, R. and Josten, J.J. (1958). J. Biol. Chem. 233, 466.
Repaske, R. and Wilson, P.W. (1952). J. Am. Chem. Soc. 74, 3101.
Resnick, H., Carter, J.R., and Kalnitsky, G. (1959). J. Biol. Chem. 234, 1711.
1048 REFERENCES
Revel, H.R. and Racker, E. (1960). Biochim. Biophys. Acta 43, 465.
Reznikoff, P. (1926). J. Gen. Physiol. 10, 9.
Rhoads, C.P. (ed.) (1955). "Antimetabolites and Cancer" Am. Assoc. Advance. Sci.,
Washington, D.C.
Rhoads, W.A. and Wallace, A. (1957). Plant Physiol. 32, xxii.
Rice, L.I. and Berman, D.A. (1961). Am. J. Physiol. 200, 727.
Richards, O.C. and Rutter, W.J. (1961). J. Biol. Chem. 236, 3185.
Richter, D. (1934). Biochem. J. 28, 901.
Rieser, P. (1961). Exptl. Cell. Res. 24, 165.
Rigbi, M. and Sela, M. (1964). Biochemistry 3, 629.
Riggs, A. (1959). In "Sulfur in Proteins" (R. Benesch et al., eds.), p. 173. Academic
Press, New York.
Riggs, A.F. (1952). J. Gen. Physiol. 36, 1.
Riggs, A.F. and Wolbach, R.A. (1956). J. Gen. Physiol. 39, 585.
Riggs, T.R. and Walker, L.M. (1963). J. Biol. Chem. 238, 2663.
Riggs, T.R., Walker, L.M., and Christensen, H.N. (1958). J. Biol. Chem. 233, 1479.
Riklis, E. and Quastel, J.H. (1958). Can. J. Biochem. Physiol. 36, 363.
Rimington, C. and Tooth, B.E. (1961). J. Biochem. (Tokyo) 49, 456.
Rindi, G. and Ferrari, G. (1959). Nature 183, 608.
Rindi, G. and Perri, V. (1961). Biochem. J. 80, 214.
Rindi, G., Perri, V., and Ventura, U. (1959). Nature 183, 1126.
Rindi, G., Perri, V., and De Caro, L. (1961). Experientia 17, 546.
Robbins, W.J. (1941). Proc. Natl. Acad. Sci. U. S. 27, 419.
Robert, B., Robert, M., and Robert, L. (1952). Bull. Soc. Chim.. Biol. 34, 183.
Roberts, E. (1953). J. Biol. Chem. 202, 359.
Roberts, E. and Simonsen, D.G. (1963). Biochem. Pharmacol 12, 113.
Roberts, E., Younger, F., and Frankel, S. (1951). J. Biol. Chem. 191, 277.
Robertson, M.E. (1943). Nature 151, 365.
Robertson, W. van B., Ropes, M.W., and Bauer, W. (1940). J. Biol. Chem. 133, 261.
Robinson, B. and Shepherd, D.M. (1961). Biochim. Biophys. Acta 53, 431.
Robinson, J.C., Keay, L., Molinari, R., and Sizer, I.W. (1962). J. Biol. Chem. 237,
2001.
Robinson, J.D., Brady, R.O., and Bradley, R.M. (1963). J. Lipid Res. 4, 144.
Robinson, J.R. (1956). J. Physiol. {London) 134, 216.
Robinson, W.G. and Coon, M.J. (1957). J. Biol. Chem. 225, 511.
Robson, J. and Fenn, P. (1961). Nature 189, 501.
Rocca, E. and Ghiretti, F. (1958). Arch. Biochem. Biophys. 77, 336.
Roche, J., Glahn, P.-E., Manchol, P., and Thoai, N. (1959). Biochim. Biophys. Acta
35, 111.
Rogers, E.F. (1962). Ann. N. Y. Acad. Sci. 98, 412.
Rogers, H.J. and Spensley, P.C. (1954). Biochim. Biophys. Acta 13, 293.
Rogers, K.S., Thompson, T.E., and Hellerman, L. (1962). Biochim. Biophys. Acta 64,
202.
Rogers, K.S., Geiger, P.J., Thompson, T.E., and Hellerman, L. (1963). J. Biol. Chem.
238, 481.
Rogers, L.L. and Shive, W. (1947). J. Biol. Chem. 169, 57.
Rohnson, G.N. (1954). J. Gen. Microbiol. 11, 412.
Romberger, J.A. and Norton, G. (1961). Plant Physiol. 36, 20.
REFERENCES 1049
Rosa, N. and Zalik, S. (1963). Can. J. Biochem. Physiol. 41, 533.
Eoscoe, H.E. and Nelson, W.L. (1964). J. Biol. Ckem. 239, 8.
Rose, I.A., Grunberg-Manago, M., Korey, S.R., and Ochoa, S. (1954). J. Biol. Chem.
211, 737.
Rose, W.C. (1924). J. Pharmacol. Exptl. Therap. 24, 147.
RoseU-Perez, M. and Lamer, J. (1964). Biochemistry 3, 773.
Rosen, F., Milholland, R.J., and Nichol, C.A. (1960). In "Inhibition in the Nervous
System and y-Aminobutyric Acid" (E. Roberts, ed.), p. 338. Macmillan (Pergamon),
New York.
Rosen, S.W. and Klotz, I.M. (1957). Arch. Biochem. Biophys. 67, 161.
Rosenberg, A.J. (1948). Compt. Bend. Soc. Biol. 142, 443.
Rosenberg, L.E., Downing, S.J., and Segal, S. (1962). J. Biol. Chem. 237, 2265.
Rosenberg, T. and Wilbrandt, W. (1962). Exptl. Cell Res. T7 , 100.
Rosenthal, A. and Geyer, R.P. (1959). Federation Proc. 18, 544.
Ross, W.F. and Green, L.S. (1941). J. Biol. Chem. 137, 105.
Roth, J.S. (1953 a). Nature 171, 127.
Roth, J.S. (1953 b). Arch. Biochem. Biophys. 44, 265.
Roth, J.S. (1956). Biochim. Biophys. Acta 21, 34.
Roth, J.S. (1958). J. Biol. Chem. 231, 1097.
Rothschild, A.M. and Schayer, R.W. (1959). Biochim. Biophys. Acta 34, 392.
Rothschild, H.A. and Barron, E.S.G. (1954). J. Biol. Chem. 209, 511.
Rothschild, H.A., Cori, O., and Barron, E.S.G. (1954). J. Biol. Chem. 208, 41.
Rothstein, A. (1959). Federation Proc. 18, 1026.
Rothstein, A. (1964). Personal communication.
Rothstein, A. and Bruce, M. (1958). J. Cellular Comp. Physiol. 51, 439.
Rothstein, A. and Hayes, A.D. (1960). J. Pharmacol. Exptl. Therap. 130, 166.
Roussos, G.G. and Nason, A. (1960). J. Biol. Chem. 235, 2997.
Rowe, A.W. and Weill, C.E. (1962). Biochim. Biophys. Acta 65, 245.
Rowland, R.L. (1952). J. Am. Chem. Soc. 74, 5482.
Roy, A.B. (1953). Biochem. J. 55, 653.
Roy, A.B. (1954a). Biochim. Biophys. Acta 15, 300.
Roy, A.B. (1954 b). Biochem. J. 57, 465.
Roy, A.B. (1956). Biochem. J. 64, 651.
Rubin, B.A. and Ladygina, M.E. (1960). Biochemistry (USSR) (English Transl.) 25,
617.
Rubin, R.J., Reynard, A., and Handschumacher, R.E. (1964). Cancer Res. 24, 1002.
Rubino, R. and Varela (1923). Klin. Wochschr. 2, 484.
Rubinstein, D., Ottolenghi, P., and Denstedt, O.F. (1956). Can. J. Biochem. Physiol.
34, 222.
RufFo, A. and Adinolfi, A. (1963). Biochem. J. 89, 50p.
Ruffo, A. and D'Abramo, F. (1952). Boll. Soc. Ital. Biol. Sper. 28, 850.
RufFo, A., Testa, E., Adinolfi, A., and Pelizza, G. (1962 a). Biochem. J. 85, 588.
Ruffo, A., Adinolfi, A., Budillon, G., and Capobianco, G. (1962 b). Biochem. J. 85, 593.
Rulon, O. (1948). Physiol. Zool. 21, 100.
Rumpf, P., d'Incan, E., and Schaal, R. (1955). Bull. Soc. Chim. France p. 122.
Runnstrom, J. and Kriszat, G. (1952). Exptl. Cell. Res. 3, 497.
Ruska, H. (1947). Arch. Exptl. Pathol. Pharmakol. 204, 576.
Ruskin, A. and Jolmson, J.E. (1949). Proc. Soc. Exptl. Biol. Med. 72, 572.
1050 REFERENCES
Ruskin, A. and Ruskin, B. (1953). J. Pharmacol. Exptl. Therap. 107, 325.
Russell, P. (1955). Nature 176, 1123.
Rutter, W.J. and Lardy, H.A. (1958). J. Biol. Chem. 233, 374.
Ryan, C.A. and King, T.E. (1962 a). Biochim. Biophys. Acta 62, 269.
Ryan, C.A. and King, T.E. (1962b). Arch. Biochem. Biophys. 85, 450.
Ryan, K.J. and Engel, L.L. (1957). J. Biol. Chem. 225, 103.
Ryley, J.F. (1955 a). Biochem. J. 59, 353.
Ryley, J.F. (1955 b). Biochem. J. 59, 361.
Ryzhkov, V.L. and Marchenko, N.K. (1954). DoH. Akad. Nauk SSSR 98, 1033.
Ryzhkov, V.L. and Marchenko, N.K. (1955). Bev. Appl. Mycol. 34, 617.
Sabato, G. di and Fonnesu, A. (1959). Biochim. Biophys. Acta 35, 358.
Sachs, G. (1921). Ber. 54, 1849.
Sachs, H.W. and Dulskas, A. (1956). Arch. Pathol. Anat. Physiol. 329, 466.
Sacktor, B. (1953). J. Gen. Physiol. 36, 371.
Sacktor, B. and Cochran, D.G. (1957). J. Biol. Cher^. 226, 241.
Sacktor, B., Thomas, G.M., Moser, J.C, and Bloch, D.I. (1953). Biol. Bull. 105, 166.
Sacktor, B., Cummins, J.I., and Packer, L. (1959). Federation Proc. 18, 314.
SafFran, M. and Prado, J.L. (1949). J. Biol. Chem. 180, 1301.
Sahasrabudhe, M.B., Nerurkar, M.K., Narurkar, M.V., Tilak, B.D., and Bhavsar,
M.D. (1960). Brit. J. Cancer 14, 457.
Sakai, H. (1962). J. Gen. Physiol. 45, 411.
Sakami, W. (1955). In "Amino Acid Metabolism" (W.D. McElroy and H.B. Glass,
eds.), p. 658. Johns Hopkins Press, Baltimore, Maryland.
Sakata, K., Hayano, S., and Sloviter, H.A. (1963). Am. J. Physiol. 204, 1127.
Salant, W. and Brodman, K. (1929 a). J. Pharmacol. Exptl. Therap. 36, 195.
Salant, W. and Brodman, K. (1929 b). J. Pharmacol. Exptl. Therap. 37, 55.
Salant, W. and Brodman, K. (1929 c). J. Pharmacol. Exptl. Therap. 37, 121.
Salant, W. and Kleitman, N. (1922). J. Pharmacol. Exptl. Therap. 19, 315.
Salant, W. and Nadler, J.E. (1927). J. Lab. Clin. Med. 13, 117.
Salant, W. and Nagler, H. (1930). Proc. Soc. Exptl. Biol. Med. 27, 859.
Salant, W. and Nagler, H. (1931). J. Pharmacol. Exptl. Therap. 41, 407.
Salle, A.J. (1943). J. Pharmacol. Exptl. Therap. 79, 271.
Salle, A.J. and Ginoza, Y.W. (1943). Proc. Soc. Exptl. Biol. Med. 54, 85.
Salle, A.J. and Lazarus, A.S. (1936). Proc. Soc. Exptl. Biol. Med. 34, 371.
Salles, J.B.V. and Ochoa, S. (1950). J. Biol. Chem. 187, 849.
Saltman, P. (1953). J. Biol. Chem. 200, 145.
Saltman, P., Fiskin, R.D., and Bellinger, S.B. (1955). Federation Proc. 14, 274.
Saltman, P., Alex, T., and McCornack, B. (1959). Arch. Biochem. Biophys. 83, 538.
Samborski, D.J. and Forsyth, F.R. (1960). Can. J. Botany 38, 467.
Sanabria, A. (1963). Brit. J. Pharmacol. 20, 352.
Sanadi, D.R., Littlefield, J.W., and Bock, R.M. (1952). J. Biol. Chem. 197, 851.
Sanborn, R.C. and Williams, CM. (1950). J. Gen. Physiol. 33, 579.
Sankar, D.V.S. (1958). Proc. Soc. Exptl. Biol. Med. 98, 198.
Sanner, T. and Pihl, A. (1962). Biochim. Biophys. Acta 62, 171.
Sanner, T. and Pihl, A. (1963). J. Biol. Chem. 238, 165.
San Pietro, A. and Lang, H.M. (1958). J. Biol. Chem. 231, 211.
Sanwal, B.D. (1961). Arch. Biochem. Biophys. 93, 377.
KEFERENCES 1051
Sanwal, B.D. and Zink, M.W. (1961). Arch. Biochem. Biophys. 94, 430.
Saroff, H.A. and Mark, H.J. (1953). J. Am. Chem. Soc. 75, 1420.
Sasaki, A. (1940). J. Biochem. (Tokyo) 32, 211.
Sasaki, S. (1962). J. Biochem. (Tokyo) 51, 335.
Sato, R., Takemori, S., and Ebata, M. (1956). J. Biochem. (Tokyo) 43, 623.
Sauer, G. and Rapoport, S. (1959). Biochem. Z. 331, 534.
Sauermann, G. (1964). Arch. Biochem. Biophys. 104, 208.
Sawai, T. (1960). J. Biochem. (Tokyo) 48, 382.
Sayre, F.W. and Roberts, E. (1958). J. Biol. Chem.. 233, 1128.
Saz, A.K., Lowery, D.L., and Jackson, L.J. (1961). J. Bacterial. 82, 298.
Saz, H.J. and Krampitz, L.O. (1955). J. Bacteriol. 69, 288.
Scala, R.A. and Lambooy, J.P. (1958). Arch. Biochem. Biophys. 78, 10.
Scarano, E., Bonaduce, L., and de Petrocellis, B. (1960). J. Biol. Chem.. 235, 3556.
Scarano, E., Bonaduce, L., and de Petrocellis, B. (1962). J. Biol. Chem. 237, 3742.
Schachter, D. and Rosen, S.M. (1959). Am. J. Physiol. 196, 357.
Schachter, D. and Taggart, J.V. (1954). J. Biol. Chem. 211, 271.
Schachter, D., Dowdle, E.B., and Schenker, H. (1960). Am. J. Physiol. 198, 275.
Schachter, H., Halliday, K.A., and Dixon, G.H. (1963). J. Biol. Chem. 238, PC3134.
Schanker, L.S. and Tocco, D.J. (1962). Biochim. Biophys. Acta 56, 469.
Schapira, G. (1946). Compt. Rend. Soc. Biol. 140, 173.
Scharff, T.G. (1961). Arch. Biochem. Biophys. 95, 329.
Schatz, V.B. (1960). In "Medicinal Chemistry" (A. Burger, ed.), 2nd ed., p. 72. Wiley
(Interscience), New York.
Schauer, R. and Hillmann, G. (1961). Z. Physiol. Chem. 325, 9.
Schayer, R.W., Smiley, R.L., and Kennedy, J. (1954). J. Biol. Chem. 206, 461.
Schellenberg, K.A. and Hellerman, L. (1958). J. Biol. Chem. 231, 547.
Schimmel, N.H., Matteucci, W.V., and Roger, W.P. (1956). Antibiot. Med. Clin.
Therapy 3, 136.
Schlegel, D.E. (1957). Virology 4, 135.
Schmidt, G. and Thannhauser, S.J. (1943). J. Biol. Chem. 149, 369.
Schneider, G. and Schmidt, S. (1959). Z. Physiol. Chem. 315, 20.
Schneiderman, H.A., Ketchel, M., and Williams, CM. (1953). Biol. Bull. 105, 188.
Scholefield, P.G. (1955). Biochem. J. 59, 177.
Scholefield, P.G. (1961). Can. J. Biochem. Physiol. 39, 1717.
Schom, R., Menke, K.-H., and Negelein, E. (1961). Z. Physiol. Chem. 323, 155.
Schonbaum, E., Birmingham, M.K., and SafFran, M. (1956). Can. J. Biochem. Physiol.
34, 527.
Schopfer, W.H. and Keller, V. (1951). Bull. Soc. Chim. Biol. 33, 1253.
Schorcher, C. and Loblich, H.J. (1960). Arch. Pathol. Anat. Physiol. 333, 587.
Schrecker, A.W. and Huennekens, F.M. (1964). Biochem. Pharmacol. 13, 731.
Schrodt, G.R., Dizon, F.Y., Peskoe, L.Y., Howell, R.S., Robie, C.H., and Stevenson,
T.D. (1960). Am. Practitioner Dig. Treat. 11, 750.
Schroeder, E.F., Woodward, G.E., and Piatt, M.E. (1933a). J. Biol. Chem. 100, 525.
Schroeder, E.F., Woodward, G.E., and Piatt, M.E. (1933 b). J. Biol. Chem. 101, 133.
Schueler, F.W. (1960). "Chemobiodynamics and Drug Design." McGraw-Hill, New
York.
Schiller, H. (1932). Biochem. Z. 255, 474.
Schuler, W. (1952). Experientia 8, 230.
1052 REFERENCES
Schultz, S.G. and Zaiusky, R. (1964). J. Gen. Physiol. 47, 1043.
Schulz, H. (1888). Arch. Ges. Physiol. 42, 517.
Schumann, E.L., Paquette, L.A., Heinzelman, R.V., Wallach, D.P., Da Vanzo, J.P.,
and Greig, M.E. (1962). J. Med. Pharm. Chem. 5, 464.
Schiitte, H.R. and Niirnberger, H. (1959). Z. Physiol. Chem. 315, 13.
Schweet, R.S. and Allen, E.H. (1958). J. Biol. Chem. 233, 1104.
Schweisfurth, R. and Schwartz, W. (1959). Acta Biol. Med. Ger. 2, 54.
Scott, R.L. and Gamble, J.L., Jr. (1961). J. Biol. Chem. 236, 570.
Scott, T.W., White, I.G., and Annison, E.F. (1962). Biochem. J. 83, 392.
Seal, U.S. and Binkley, F. (1957). J. Biol. Chem. 128, 193.
Sealock, R.R. and Goodland, R.L. (1944). J. Am. Chem. Soc. 66, 507.
Sealock, R.R. and Livermore, A.H. (1949). J. Biol. Chem. 177, 553.
Sealock, R.R. and White, H.S. (1949). J. Biol. Chem,. 181, 393.
Seaman, G.R. (1949). Biol. Bull. 96, 257.
Seaman, G.R. (1952). Arch. Biochem. Biophys. 39, 241.
Seaman, G.R. (1954). Arch. Biochem. Biophys. 48, 424.
Seaman, G.R. (1956). Exptl. Parasitol. 5, 138.
Seaman, G.R. and Houlihan, R.K. (1950). Arch. Biochem. 26, 436.
Seaman, G.R. and Naschke, M.D. (1955). J. Biol. Chem. 217, 1.
Seeley, H.W. (1955). In " Methods in Enzymology" (S.P. Colowick and N.O. Kaplan,
eds.). Vol. 1, p. 624. Academic Press, New York.
Seelich, F. and Letnansky, K. (1960). Z. Krebsforsch. 63, 236.
Seelich, F. and Letnansky, K. (1961). Naturivissenschaften 48, 408.
Seevers, M.H., Shideman, F.E., W^oods, L.A., Weeks, J.R., and Kruse, W.T. (1950).
J. Pharmacol. Exptl. Therap. 99, 69.
Segal, H.L. (1959). J. Am. Chem. Soc. 81, 4047.
Segal, H.L. and Boyer, P.D. (1953). J. Biol. Chem. 204, 265.
Segal, H.L. and Brenner, B.M. (1960). J. Biol. Chem. 235, 471.
Segal, H.L., Beattie, D.S., and Hopper, S. (1962). J. Biol. Chem. 237, 1914.
Seibert, M.A., Kreke, C.W., and Cook, E.S. (1950). Science 112, 649.
Sekuzu, I., Jurtshuk, P., Jr., and Green, D.E. (1963). J. Biol. Chem. 238, 975.
Sela, M. (1962). J. Biol. Chem.. 237, 418.
Sela, M. and Steiner, L.A. (1963). Biochemistry 2, 416.
Selim, A.S.S.M. and Greenberg, D.M. (1960). Biochim. Biophys. Acta 42, 211.
Sellinger, O.Z. and Weiler, P., Jr. (1963). Biochem. Pharmacol. 12, 989.
Sen, D.K. (1959). Nature 184, 459.
SentheShanmuganathan, S. (1960). Biochem. J. 77, 619.
Serif, G.S. and Stewart, C.J. (1958). Federation Proc. 17, 309.
Serif, G.S. and Wick, A.N. (1958). J. Biol. Chem. 233, 559.
Serif, G.S., Stewart, C.J., Nakada, H.L, and Wick, A.N. (1958). Proc. Soc. Exptl. Biol.
Med. 99, 720.
Sery, T.W. and Furgiuele, F.P. (1961). Am. J. Ophthalmol. 51, 42.
Severens, J.M. and Tanner, F.W. (1945). J. Bacteriol. 49, 383.
Sguros, P.L. and Hartsell, S.E. (1952 a). J. Bacteriol. 64, 811.
Sguros, P.L. and Hartsell, S.E. (1952 b). J. Bacteriol. 64, 821.
Shacter, B. (1953). Arch. Biochem. Biophys. 46, 324.
Shacter, B. (1957). J. Natl. Cancer Inst. 18, 455.
Shah, V.K. and Ramakrishnan, C.V. (1963). Enzymologia 26, 53.
REFERENCES 1053
Shanes, A.M. and Brown, D.E.S. (1942). J. Cellular Comp. Physiol. 19, 1.
Shannon, L.M., Young, R.H., and Dudley, C. (1959). Nature 183, 683.
Shapiro, D.M. and Gellhorn, A. (1951). Cancer Res. 11, 35.
Shapiro, D.M., Kream, J., and Dietrich, L.S. (1952). Proc. Soc. Ezptl. Biol. Med. 81, 616.
Shapiro, D.M., Shils, M.E., and Dietrich, L.S. (1953). Cancer Res. 13, 703.
Shapiro, D.M., Dietrich, L.S., and Shils, M.E. (1956). Cancer Res. 16, 575.
Shapiro, D.M., Dietrich, L.S., and Shils, M.E. (1957). Cancer Res. 17, 600.
Sharon, N. and Lipmann, F. (1957). Arch. Biochem. Biophys. 69, 219.
Shaw, E. and Woolley, D.W. (1952). J. Biol. Chem. 194, 641.
Shaw, P.D. and Hager, L.P. (1959). J. Biol. Chem. 234, 2565.
Shaw, W.H.R. (1954). Scieyice 120, 361.
Shaw, W.H.R. and Raval, D.N. (1961). J. Am. Chem. Soc. 83, 2866.
Sheets, R.F. and Hamilton, H.E. (1958). J. Lab. Clin. Med. 52, 138.
Sheets, R.F., Hamilton, H.E., and DeGowin, E.L. (1956 a). Proc. Soc. Exptl. Biol.
Med. 91, 423.
Sheets, R.F., Hamilton, H.E., DeGowin, E.L., and King, R.L. (1956 b). J. Appl.
Physiol. 9, 145.
Sheinin, R. and Crocker, B.F. (1961). Can. J. Biochem. Physiol. 39, 55.
Shemin, D. and Kumin, S. (1952). J. Biol. Chem. 198, 827.
Sherman, I.R. and Corley, R.C. (1952). Federation Proc. 11, 285.
Shibasaki, I. and Terui, G. (1953). J. Ferment. Technol. {Japan) 31, 238.
Shichi, H. and Hackett, D.P. (1962). Phytochemistry 1, 131.
Shideman, F.E. and Rene, R.M. (1951a). J. Pharmacol. Exptl. Therap. 101, 32.
Shideman, F.E. and Rene, R.M. (1951b). Am. J. Physiol. 166, 104.
Shideman, F.E., Woods, L.A., and Seevers, M.H. (1950a). J. Pharmacol. Exptl. Therap.
98, 29.
Shideman, F.E., Woods, L.A., and Seevers, M.H. (1950b). J. Pharmacol. Exptl. Therap.
99, 98.
Shifrin, S. and Kaplan, N.O. (1960). Advan. Enzymol. 22, 337.
Shigeura, H.T. and Gordon, C.N. (1962). J. Biol. Chem. 237, 1937.
Shiio, I., Otsuka, S.-I., and Takahashi, M. (1961). J. Biochem. (Tokyo) 49, 397.
Shilo, M. (1957). J. Gen. Microbiol. 16, 472.
Shimazono, H. (1955). J. Biochem. (Tokyo) 42, 321.
Shimi, I.R. and Nour El Dein, M.S. (1962). Arch. Mikrobiol. 41, 261.
Shimomura, 0., Johnson, F.H., and Saiga, Y. (1961). J. Cellular Comp. Physiol. 58,
113.
Shimomura, 0., Johnson, F.H., and Saiga, Y. (1963). J. Cellular Comp. Physiol. 61,
275.
Shinohara, K. (1935). J. Biol. Chem. Ill, 435.
Shkol'nik, M.Y. (1954). Dokl. Akad. Nauk SSSR 98, 443.
Shore, V. and Shore, B. (1960). Am. J. Physiol. 198, 187.
Shore, V. and Shore, B. (1962). Am. J. Physiol. 203, 15.
Shore, V., Shore, B., and Hsieh, K. (1959). Federation Proc. 18, 144.
Shrago, E. and Falcone, A.B. (1963 a). Biochim. Biophys. Acta 67, 147.
Shrago, E. and Falcone, A.B. (1963 b). Biochim. Biophys. Acta 73, 7.
Shug, A.L., Hamilton, P.B., and Wilson, P.W. (1956). In "Inorganic Nitrogen Meta-
bolism" (W.D. McElroy and H.B. Glass, eds.), p. 344. Johns Hopkins Press, Bal-
timore, Maryland.
1054 REFEKENCES
Shug, A.L., Fraser, D., and Mahler, H.R. (1959). Biochim. Biophys. Acta 36, 142.
Shug, A.L., Mahler, H.R., and Fraser, D. (1960). Biochim. Biophys. Acta 42, 255.
Shukuya, R. and Schwert, G.W. (1960). J. Biol. Chem. 235, 1658.
Shull, K.H. (1959). Nature 183, 259.
Shulman, A. (1956). Nature 177, 703.
Shwartzman, G. and Hift, H. (1951). J. Nutr. 44, 575.
Sibly, P.M. and Wood, J.G. (1951). Australian J. Sci. Res. Ser. B4, 500.
Sie, H.-G. and Fishman, W.H. (1954). J. Biol. Chem. 209, 73.
Siebert, G., Carsiotis, M., and Plant, G.W.E. (1957). J. Biol. Che?n. 226, 977.
Siebert, G., Schmitt, A., and Trazler, G. (1963). Z. Physiol. Chem. 332, 160.
Siegel, L. and Englard, S. (1962). Biochiyn. Biophys. Acta 64, 101.
Siegelman, H.W., Chow, C.T., and Biale, J.B. (1958). Plant Physiol. 33, 403.
Siekevitz, P., Low, H., Ernster, L., and Lindberg, O. (1958). Biochim. Biophys. Acta
29, 378.
Sih, C.J., Hamilton, P.B., and Knight, S.G. (1958). J. Bacteriol. 75, 623.
Silber, R., Huennekens, F.M., and Gabrio, B.W. (1962). Arch. Biochem. Biophys. 99,
328.
Silberman, H.R. and Wyngaarden, J.B. (1961). Biochim. Biophys. Acta 47, 178.
Sillen, L.G. (1949). Acta Chem. Scand. 3, 539.
Silverman, J.L. (1963). Biochim. Biophys. Acta 78, 94.
Simola, P.E. and Kosunen, T. (1938). Suomen Kemistilehti B11, 22.
Simon, E.W. (1957). J. Exptl. Botany 8, 20.
Simon, E.W. (1959). J. Exptl. Botany 10, 125.
Simon, L.N. and Myers, T.C. (1961). Biochim. Biophys. Acta 51, 178.
Simonds, J.P. and Hepler, O.E. (1945). Arch. Pathol. 39, 103.
Simonsen, D.H. and van Wagtendonk, W.J. (1956). J. Gen. Microbiol. 15, 39.
Simpson, F.J. (1960). Can. J. Biochem. Physiol. 38, 115.
Simpson, R.B. (1961). J. Am. Chem. Sac. 83, 4711.
Singer, B. and Fraenkel-Conrat, H. (1962). Biochemistry 1, 852.
Singer, T.P. (1948). J. Biol. Chem. 174, 11.
Singer, T.P. and Barron, E.S.G. (1944). Proc. Soc. Exptl. Biol. Med. 56, 120.
Singer, T.P. and Barron, E.S.G. (1945). J. Biol. Chem. 157, 241.
Singer, T.P. and Hofstee, B.H.J. (1948 a). Arch. Biochem. 18, 229.
Singer, T.P. and Hofstee, B.H.J. (1948 b). Arch. Biochem. 18, 245.
Singer, T.P. and Kearney, E.B. (1950). Arch. Biochem. 27, 348.
Singer, T.P. and Lara, F.J.S. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo Kyoto,
1957 p. 330. Maruzen, Tokyo.
Singer, T.P., and Lusty, C.J. (1960). Biochem. Biophys. Res. Commun. 2, 276.
Singer, T.P., Massey, V., and Kearney, E.B. (1956 a). Biochim. Biophys. Acta 19, 200.
Singer, T.P., Kearney, E.B., and Bernath, P. (1956 b). J. Biol. Chem. 223, 599.
Singer, T.P., Kearney, E.B., and Massey, V. (1957). Advan. Enzymol. 18, 65.
Sipos, J.C, Sawatzky, H., and Wright, G.F. (1955). J. Am. Chem. Soc. 77, 2759.
Siu, P.M.L. and Wood, H.G. (1962). J. Biol. Chem. 237, 3044.
Sivaramakrishnan, V.M. and Sarma, P.S. (1954). Biochim. Biophys. Acta 14, 579.
Sizer, I.W. (1942 a). J. Gen. Physiol. 25, 399.
Sizer, I.W. (1942 b). J. Biol. Chem. 145, 405.
Sizer, I.W. (1945). J. Biol. Chem. 160, 547.
Sizer, I.W. and Tytell, A. A. (1941). J. Biol. Chem. 138, 631.
REFERENCES 1055
Skarnes, R.C. and Watson, D.W. (1955). J. Bacteriol. 70, 110.
Skipper, H.E., Mitchell, J.H., Jr., and Bennett, L.L., Jr. (1950). Cancer Res. 10, 510.
Skoda, J., Kara, J., Sormova, Z., and Sorm, F. (1959). Biochim. Bilphys. Acta 33, 579.
Skou, J.C. (1963). Biochem. Biophys. Res. Commun. 10, 79.
Slater, E.C. (1949). Biochem. J. 45, 130.
Slater, E.C. and Bonner, W.D., Jr. (1952). Biochem. J. 52, 185.
Slater, E.C. and Holton, F.A. (1954). Biochem. J. 56, 28.
Slater, T.F. (1959). Nature 183, 1679.
Slechta, L. (1960). Biochem. Biophys. Res. Commun. 3, 596.
Slein, M.W., Cori, G.T., and Cori, C.F. (1950). J. Biol. Chem. 186, 763.
Slingerland, D.W. (1955). J. Clin. Endocrinol. Metab. 15, 131.
Sloane, N.H. and Boggiano, E.M. (1960). Arch. Biochem. Biophys. 87, 217.
Smalt, M.A., Kreke, C.W., and Cook, E.S. (1957). J. Biol. Chem. IIA, 999.
Smillie, R.M. (1956). Australian J. Biol. Sci. 9, 81.
Smith, C.G. and Bernstein, I. A. (1961). Biochim. Biophys. Acta 52, 184.
Smith, C.G., Lummis, W.L., and Grady, J.E. (1959). Cancer Res. 19, 847.
Smith, D.E., Czarnetzky, E.J., and Mudd, S. (1936). Am. J. Med. Sci. 192, 790.
Smith, E.L. (1958). J. Biol. Chem. 233, 1392.
Smith, E.L., Lumry, R., and Polglase, W.J. (1951). J. Phys. Colloid Chem. 55, 125.
Smith, E.L., Davis, N.C., Adams, E., and Spackman, D.H. (1954 a). In "The Mech-
anism of Enzyme Action" (W.D. McElroy and H.B. Glass, eds.), p. 291. Johns
Hopkins Press, Baltimore, Maryland.
Smith, E.L., Kimniel, J.R., and Brown, D.M. (1954b). J. Biol. Chem. 207, 533.
Smith, E.L., Kimmel, J.R., Brown, D.M., and Thompson, E.O.P. (1955). J. Biol.
Chem. 215, 67.
Smith, F. and Spriestersbach, D. (1954). Nature 174, 466.
Smith, H.M. (1958). J. Cellular Camp. Physiol. 51, 161.
Smith, J.E. (1963). Can. J. Microbiol. 9, 345.
Smith, J.T. (1963 a). Biochem. J. 87, 40p.
Smith, J.T. (1963 b). Nature 197, 900.
Smith, L. and BaltschefFsky, M. (1959). J. Biol. Chem. 234, 1575.
Smith, L.H., Jr. and Sullivan, M. (1960). Biochim. Biophys. Acta 39, 554.
Smith, M.E. and Greenberg, D.M. (1957). J. Biol. Chem. 226, 317.
Smith, P.F. (1959). J. Bacteriol. 77, 682.
Smith, P. J.C. (1960 a). Biochem. J. 76, 508.
Smith, P.J.C. (1960 b). Biochem. J. 76, 514.
Smith, S.E. (1960 a). Brit. J. Pharmacol. 15, 319.
Smith, S.E. (1960 b). Ciba Found. Symp., Adrenergic Mechanisms p. 25.
Smith, S.E. (1963). Brit. J. Pharmacol. 20, 178.
Smyth, D.H. (1940). Biochem. J. 34, 1046.
Sneed, M.C. and Brasted, R.C. (1955). "Comprehensive Inorganic Chemistry," Vol. 4,
Van Nostrand, Princeton, New Jersey.
Snodgrass, P.J. and Hoch, F.L. (1959). Federation Proc. 18, 326.
Snodgrass, P.J., Vallee, B.L., and Hoch, F.L. (1960). J. Biol. Chem. 235, 504.
Snoswell, A.M. (1959). Australian J. Exptl. Biol. Med. Sci. 37, 49.
Snoswell, A.M. (1962). Biochim. Biophys. Acta 60, 143.
Snyder, R., Vogel, W.H., and Schulman, M.P. (1963). Biochem. Biophys. Res. Com-
mun. 13, 335.
1056 REFERENCES
Snyder, S.H., Silva, O.L., and Kies, M.W. (1961). J. Biol. Chem. 236, 2996.
Sohler, M.R., Seibert, M.A., Kreke, C.W., and Cook, E.S. (1952). J. Biol. Chem. 198,
281.
Sokoloff, B., Saelhof, C, Kato, S., Chemelin, I., and Beach, J. (1956). Growth 20, 265.
Soldatenkov, S.V. and Mazurova, T.A. (1957). Biochemistry (USSR) (English Transl.)
11, 345.
Solomon, J.B. (1958). Biochem. J. 70, 529.
Sols, A. (1956). Biochim. Biophys. Acta 19, 144.
Sols, A. and Crane, R.K. (1953). Federation Proc. 12, 271.
Sols, A. and Crane, R.K. (1954). J. Biol. Chem. 206, 925.
Sols, A. and de la Fuente, G. (1957). Biochim. Biophys. Acta 24, 206.
Sols, A., Cadenas, E., and Alvarado, F. (1960 a). Science 131, 297.
Sols, A., Heredia, C.F., and Ruiz-Amil, M. (1960 b). Biochem. Biophys. Res. Commun.
2, 126.
Solvonuk, P.F. and Collier, H.B. (1955). Can. J. Biochem. Physiol. 33, 38.
Solvonuk, P.F., McRae, S.C, and Collier, H.B. (1956). Can. J. Biochem. Physiol. 34,
481.
Somers, E. (1959). Nature 184, 475.
Sondheimer, E. and Griffin, D.H. (1960). Science 131, 672.
Soodak, M. and Cerecedo, L.R. (1944). J. Am. Chem. Soc. 66, 1988.
Sophianopoulos, A.J. and Corley, R.C. (1959). Federation Proc. 18, 328.
Sorbo, B. (1963). Acta Chem. Scand. 17, S 107.
Sorbo, B.H. (1951). Acta Chem. Scand. 5, 724.
Sorsby, A., Newhouse, J.P., and Lucas, D.R. (1957). Brit. J. Ophthalmol. 41, 309.
Sourkes, T.L. (1954). Arch. Biochem. Biophys. 51, 444.
Sourkes, T.L. and Towsend, E. (1955). Can. J. Biochem. Physiol. 33, 735.
Speck, J.F., Moulder, J.W., and Evans, E.A., Jr. (1946). J. Biol. Chem. 164, 119.
Spector, A. (1963). J. Biol. Chem.. 238, 1353.
Spector, W.G. (1953). Proc. Roy. Soc. B141, 268.
Speer, H.L. and Jones, R.F. (1964). Physiol. Plantarum 17, 287.
Spencer, A.F. and Lowenstein, J.M. (1962). J. Biol. Chem. 237, 3640.
Spencer, B. and Williams, R.T. (1951). Biochem. J. 48, 538.
Spencer, D. (1959). Australian J. Biol. Sci. 12, 181.
Spencer, H.C., Rowe, V.K., and McCollister, D.D. (1950 a). J. Pharmacol Exptl.
Therap. 98, 30.
Spencer, H.C., Rowe, V.K., and McCollister, D.D. (1950 b). J. Pharmacol. Exptl.
Therap. 99, 57.
Spencer, R.P. and Brody, K.R. (1964). Am. J. Physiol. 206, 653.
Spencer, R.P. and Knox, W.E. (1962). Arch. Biochem. Biophys. 96, 115.
Spencer, R.P., Valentine, R.J., and Field, J.B. (1956). Acta Pharmacol. Toxicol. 12,
196.
Spensley, P.C. and Rogers, H.J. (1954). Nature 173, 1190.
Speyer, J.F. and Dickman, S.R. (1956). J. Biol. Chem. 220, 193.
Spiegelman, S., Kamen, M.D., and Sussman, M. (1948). Arch. Biochem. 18, 409.
Spiro, R.G. (1958). J. Biol. Chem. 233, 546.
Spizizen, J. (1943). J. Infect. Diseases 73, 222.
Spratt, N.T. (1950). Biul. Bull. 99, 120.
Srinivasan, P.R. and Sprinson, D.B. (1959). J. Biol. Chem. 234, 716.
REFERENCES 1057
Srinivasan, P.R. Brunfaut, M., and Errera, M. (1964). Exptl. Cell. Res. 34, 61.
Stadtman, E.R. (1955). In "Methods in Enzymology" (S.P. Colowick and N.O. Ka-
plan, eds.), Vol. 1, p. 596. Academic Press, New York.
Stadtman, E.R., Cohen, G.N., and LeBras, G. (1961). Ann. N. Y. Acad. Sci. 94, 952.
Staemmler, M. (1956). Arch. Pathol. Anat. Physiol. 328, 1.
Stafford, H.A. (1956). Plant Physiol. 31, 135.
Stalder, K. (1958). Z. Physiol. Chem. 311, 221.
Stalder, K. (1959). Z. Physiol. Chem. 314, 205.
Stanley, R.G. and Conn, E.E. (1957). Plant Physiol. 32, 412.
Stare, F.J. and Baumann, C.A. (1936). Pwc. Roy. Soc. B121, 338.
Stare, F.J. and Baumann, C.A. (1939). Cold Spring Harbor Symp. Quant. Biol. 7, 227.
Stare, F.J. and Baumann, C.A. (1940). J. Biol. Chem. 133, 453.
Stare, F.J., Lipton, M.A., and Goldinger, J.M. (1941). J. Biol. Chem. 141, 981.
Stark, J.B., Goodban, A.E., and Owens, H.S. (1951). Anal. Chem. 23, 413.
Starr, J.L. and Goldthwait, D.A. (1963). J. Biol. Chem. 238, 682.
Stauff, J. and Uhlein, E. (1958). Biochem. Z. 329, 549.
Stearn, A.E. (1935). J. Oen. Physiol. 18, 301.
Steberl, E.A., Wasson, G.W., and Porter, J.W. (1960). Biochem. Biophys. Res. Com-
mun. 2, 174.
Stedman, R.L. and Kravitz, E. (1955). Arch. Biochem. Biophys. 59, 260.
Stedman, R.L., Engel, S.L., and Bilse, I.M. (1954). J. Am. Pharm. Assoc, Sci. Ed.
43, 622.
Steel, K.J. (1960). J. Pharm. Pharmacol. 12, 59.
Steele, A.B., Boese, A.B., and Dull, M.F. (1949). J. Org. Chem. 14, 460.
Stein, E., Magin, J., and Kleinfeld, M. (1960). Am. J. Physiol. 199, 460.
Steinberg, D. and Fredrickson, D.S. (1955). Proc. Soc. Exptl. Biol. Med. 90, 232.
Steinberg, D. and Vaughan, M. (1956). Arch. Biochem. Biophys. 65, 93.
Steiner, D.F., Rauda, V., and Williams, R.H. (1961). J. Biol. Chem. 236, 299.
Stephenson, M.L., Thimann, K.V., and Zamecnik, P.C. (1956). Arch. Biochem. 65,
194.
Stern, J.R. (1948). Biochem. J. 43, 616.
Stern, J.R. (1956). J. Biol. Chem. ll^, 33.
Stem, J.R. and del Campillo, A. (1956). J. Biol. Chem. 218, 985.
Stern, J.R. and Friedman, D.L. (1960). Biochem. Biophys. Res. Commun. 2, 82.
Stevens, CO. and Henderson, L.M. (1959). J. Biol. Chem. 234, 1188.
Stewart, H.B. (1957). Federation Proc. 16, 256.
Steyn-Parve, E.P. (1954). Biochim. Biophys. Acta 14, 440.
Stickland, R.G. (1959 a). Biochem. J. 73, 654.
Stickland, R.G. (1959 b). Biochem. J. 73, 646.
Stjernholm, R. and Wood, H.G. (1961). Proc. Natl. Acad. Sci. U.S. 47, 303.
Stockell, A. (1959). J. Biol. Chem. 234, 1293.
Stockell, A. and Smith, E.L. (1957). J. Biol. Chem. 227, 1.
Stocken, L.A. (1947). Biochem. J. 41, 358.
Stoerk, H.C. (1947). J. Biol. Chem. 171, 437.
Stoerk, H.C. (1950). Ann. N.Y. Acad. Sci. 52, 1302.
Stone, B.A. and Hochster, R.M. (1956). Can. J. Microbiol. 2, 623.
Stone, C.A., Ross, C.A., Wenger, H.C, Ludden, C.T., Blessing, J.A., Totaro, J.A.,
and Porter, CC (1962). J. Pharmacol. Exptl. Therap. 136, 80.
1058 REFERENCES
Stone, J.E. and Potter, V.R. (1957). Cancer Res. 17, 800.
Stoneman, F., Shideman, F.E., and Rathbun, R.C. (1951). Federation Proc. 10, 337.
Stoppani, A.O.M. (1946). Anales Asoc. Quim. Arg. 34, 33.
Stoppani, A.O.M. (1948). Enzymologia 13, 165.
Stoppani, A.O.M. and Brignone, J. A. (1956). Biochem. J. 64, 196.
Stoppani, A.O.M. and Brignone, J.A. (1957). Arch. Biochem. Biophys. 68, 432.
Stoppani, A.O.M. and Milstein, C. (1957 a). Biochem. J. 67, 406.
Stoppani, A.O.M. and Milstein, C. (1957 b). Biochim. Biophys. Acta 24, 655.
Stoppani, A.O.M., Actis, A.S., Deferrari, J.O., and Gonzalez, E.L. (1953). Biochem. J.
54, 378.
Stoppani, A.O.M., Conches, L., de Favelukes, S.L.S., and Sacerdote, F.L. (1958 a).
Biochem. J. 70, 438.
Stoppani, A.O.M., de Favelukes, S.L.S., and Conches, L. (1958 b). Arch. Biochem.
Biophys. 73, 453.
Storey, I.D.E. (1964). Biochem. J. 90, 15p.
Stracher, A. and Chan, P.C. (1961). Arch. Biochem. Biophys. 95, 435.
Straessle, R. (1951). J. Am. Chem. Soc. 73, 504.
Straessle, R. (1954). J. Am. Chem. Soc. 76, 3138.
Straub, F.B. (1937). Z. Physiol. Chem. 249, 189.
Strauss, N. and Jann, G.J. (1956). J. Bacteriol. 71, 439.
Strecker, H.J. (1960). J. Biol. Chem.. 235, 3218.
Strecker, H.J. and Korkes, S. (1952). J. Biol. Chem. 196, 769.
Strength, D.R., Mondy, N.I., and Daniel, L.J. (1954). Arch. Biochem. Biophys. 48, 141.
Strickland, K.F. (1954). Can. J. Biochem. Physiol. 32, 50.
Strickler, J.C. and Kessler, R.H. (1963). Am. J. Physiol. 205, 117.
Stricks, W. and Kolthoff, I.M. (1953). J. Am. Chem. Soc. 75, 5673.
Stricks, W., Kolthoff, I.M., and Heyndrickx, A. (1954). J. Am. Chem. Soc. 76, 1515.
Strittmatter, C.F. (1959). J. Biol. Chem. 234, 2794.
Strittmatter, P. (1959). J. Biol. Chem. 234, 2661.
Strittmatter, P. (1961a). J. Biol. Chem. 236, 2329.
Strittmatter, P. (1961b). J. Biol. Chem. 236, 2336.
Strittmatter, P. and Velick, S.F. (1956). J. Biol. Chem. 221, 253.
Strohman, R.C. and Samorodin, A.J. (1962). J. Biol. Chem. 237, 363.
Strominger, J.L. and Mapson, L.W. (1957). Biochem. J. 66, 567.
Stumpf, P.K. (1948). J. Biol. Chem. 176, 233.
Stumpf, P.K. and Barber, G.A. (1956). Plant Physiol. 31, 304.
Stumpf, P.K., Zarudnaya, K., and Green, D.E. (1947). J. Biol. Chem. 167, 817.
Stumpf. P.K., Loomis, W.D., and Michelson, C. (1951). Arch. Biochem. 30, 126.
Subramanian, V., Stent, H.B., and Walker, T.K. (1929). J. Chem. Soc. p. 2485.
Sumizu, K., Yoshikawa, M., and Tanaka, S. (1961). J. Biochem. (Tokyo) 50, 538.
Summers, W.A. (1957). Exptl. Parasitol. 6, 194.
Sundaram, S. and Sarma, P.S. (1960 a). Enzymologia 21, 219.
Sundaram, S. and Sarma, P.S. (1960b). Biochem. J. 77, 465.
Sundaram, T.K., Rajagopalan, K.V., and Sarma, P.S. (1958). Biochem. J. 70, 196.
Siipfle, K. (1923). Arch. Hyg. Bakteriol. 93, 252.
Surtshin, A. (1957). Am. J. Physiol. 190, 271.
Sussman, A., Holton, R., and von Boventer-Heidenhain, B. (1958). Arch. Mikrobiol.
29, 38.
REFERENCES 1059
Sutton, C.R. and King, H.K. (1959). Biochem. J. 73, 43p.
Sutton, C.R. and King, H.K. (1962). Arch. Biochem. Biophys. 96, 360.
Sutton, W.B. (1954). J. Biol. Chem. 210, 309.
Suzuoki, T. (1912). "Morphologie der Nierensekretion unter pathologischen Bedin-
gungen." Fischer, Jena.
Suzuoki, Z. and Suzuoki, T. (1951). J. Biochem. (Tokyo) 38, 237.
Suzuoki, Z. and Suzuoki, T. (1954). Nature 173, 83.
Swan, J.M. (1959). In "Sulfur in Proteins" (R. Benesch et al., eds.), p. 3. Academic
Press, New York.
Swenson, A.D. and Boyer, P.D. (1957). J. Am. Chem. Soc. 79, 2174.
Swensson, A., Lundgren, K.-D., and Linstrom, O. (1959). A.M. A. Arch. Ind. Health
20, 432.
Swim, H.E. and Krampitz, L.O. (1954). J. Bacteriol. 67, 419.
Swingle, K.F., Axelrod, A.E., and Elvehjem, C.A. (1942). J. Biol. Chem. 145, 581.
Szabolcsi, G. and Biszku, E. (1961). Biochim. Biophys. Acta 48, 335.
Szabolcsi, G., Biszku, E., and Sajgo, M. (1960). Acta Physiol. Acad. Sci. Hung. 17, 183.
Szep, O. (1940). Biochem. Z. 307, 79.
Tabachnick, M., Srere, P. A., Cooper, J., and Racker, E. (1958). Arch. Biochem. Bio-
phys. 74, 315.
Tabor, H. and Wyngarden, L. (1959). J. Biol. Chem. 234, 1830.
Tachibana, S., Katagiri, H., and Yamada, H. (1958). Proc. Intern. Symp. Enzyme
Chem., Tokyo Kyoto 1957 p. 154. Maruzen, Tokyo.
Tager, J.M. (1963). Biochim,. Biophys. Acta 73, 341.
Takagaki, G., Hirano, S., and Tsukada, Y. (1957). Arch. Biochem. Biophys. 68, 196.
Takagaki, G., Hirano, S., and Tsukada, Y. (1958). J. Biochem. {Tokyo) 45, 41.
Takagi, Y. and Horecker, B.L. (1957). J. Biol. Chem. 225, 77.
Takahashi, H. and Nomura, M. (1952). J. Agr. Chem. Soc. Japan 26, 99.
Takahashi, N. (1960 a). J. Biochem. (Tokyo) 47, 230.
Takahashi, N. (1960 b). J. Biochem. (Tokyo) 48, 691.
Takebe, I. (1961). J. Biochem. (Tokyo) 50, 245.
Takemori, A.E. and Mannering, G.J. (1958). J. Pharmacol. Exptl. Therap. 123, 171.
Takenaka, Y. and Schwert, G.W. (1956). J. Biol. Chem. 112, 157.
Takeuchi, T. (1933). J. Biochem. (Tokyo) 17, 47.
Talalay, P. and Dobson, M.M. (1953). J. Biol. Chem. 205, 823.
Talalay, P. and Marcus, P.I. (1956). J. Biol. Chem. 218, 675.
Tanada, T. (1956). Plant Physiol. 31, 403.
Tanaka, K. (1961). J. Biochem. (Tokyo) 50, 102.
Tanaka, M., Kato, Y., and Kinoshita, S. (1961). Biochem. Biophys. Res. Commun. 4,
114.
Tanaka, S. (1938). J. Biochem. (Tokyo) 28, 119.
Tanaka, S., Shimizu, T., Ishishita, Y., Koike, M., and Hiramitsu, S. (1952). Nisshin
Igaku 39, 220.
Tanaka, Y. (1960). J. Biochem. (Tokyo) 47, 1.
Tapley, D.F. (1956). J. Biol. Chem. Ill, 325.
Tarr, H.L.A. (1959). Can. J. Biochem. Physiol. 37, 961.
Tashian, R.E. (1961). Metab., Clin. Exptl. 10, 393.
Tata, J.R. (1959). Biochim. Biophys. Acta 35, 567.
1060 REFERENCES
Tata, J.R. (1960). Biochem. J. 77, 214.
Taylor, E.L., Meriwether, B.P., and Park, J.H. (1963). J. Biol. Chem. 238, 734.
Taylor, E.S. and Gale, E.F. (1945). Biochem. J. 39, 52.
Taylor, F.J. (1960). Proc. Roy. Soc. B151, 483.
Taylor, H.D. and Austin, J.H. (1918). J. Exptl. Med. 27, 635.
Taylor, J.J. and Johnson, C.E. (1962). Enzymologia 25, 87.
Telkka, A. and Mustakallio, K.K. (1954). Experientia 10, 216.
Terner, C. (1951). Biochem. J. 50, 145.
Terner, C. (1954). Biochem. J. 56, 471.
Terner, C, Eggleston, L.V., and Krebs, H.A. (1950). Biochem. J. 47, 139.
Theis, F.V. and Freeland, M.R. (1940). Arch. Surg. 40, 190.
Theorell, H. (1956). In "Currents in Biochemical Research" (D.E. Green, ed.), p. 275.
Wiley (Interscience), New York.
Theorell, H. and Bonnichsen, R. (1951). Acta Chem. Scand. 5, 329.
Theorell, H., Yagi, K., Ludwig, G.D., and Egami, F. (1957). Nature 180, 922.
Thimann, K.V. and Bonner, W.D., Jr. (1948). Am. J. Botany 35, 271.
Thimann, K.V. and Bonner, W.D., Jr. (1949). Am. J. Botany 36, 214.
Thoelen, H. and Pletscher, A. (1953). Helv. Physiol. Pharmacol. Acta 11, 64.
Thomas, C.A. (1954). J. Am. Chem. Soc. 76, 6032.
Thomas, G.W. and Cook, E.S. (1947). J. Bacteriol. 54, 527.
Thomas, K. and Kalbe, H. (1953). Z. Physiol. Chem. 293, 239.
Thomas, K. and Stalder, K. (1958). Z. Physiol. Chem. 313, 22.
Thomas, K. and Stalder, K. (1959). Z. Physiol. Chem. 315, 256.
Thompson, E.O.P. (1954). J. Biol. Chem. 207, 563.
Thompson, R.H.S. and Whittaker, V.P. (1947). Biochem. J. 41, 342.
Thompson, R.L. (1947). J. Immunol. 55, 345.
Thompson, W. and Dawson, R.M.C. (1964). Biochem. J. 91, 237.
Thomson, J.F. (1959). Proc. Soc. Exptl. Biol. Med. 101, 1.
Thomson, J.F. and Sato, T.R. (1960). Arch. Biochem. Biophys. 89, 139.
Thorn, M.B. (1953). Biochem. J. 54, 540.
Thorn, M.B. (1959). Biochem. J. 71, 23p.
Thorne, C.J.R. and Kaplan, N.O. (1963). J. Biol. Chem. 238, 1861.
Thome, K.J.I, and Kodicek, E. (1962). Biochim. Biophys. Acta 59, 295.
Thunberg, T. (1909). Skand. Arch. Physiol. 11, 430.
Thunberg, T. (1933). Biochem. Z. 258, 48.
Thyagarajan, B.S. (1958). Chem. Rev. 58, 439.
Tietz, A. (1961). J. Lipid Res. 2, 182.
Tietze, F. and Klotz, I.M. (1952). Arch. Biochem. Biophys. 35, 355.
Tinacci, F. (1953). Boll. Soc. Ital. Biol. Sper. 29, 1457.
T'ing-seng, H. (1961a). Biochemistry (USSR) (English Transl.) 25, 870.
T'ing-seng, H. (1961b). Biochemistry {USSR) {English Transl.) 26, 493.
Tokuyama, K. (1959). J. Biochem. {Tokyo) 46, 1453.
Tokuyama, K. and Dawson, C.R. (1962). Biochim. Biophys. Acta 56, 427.
Tolba, M.K. and Eskarous, J.K. (1962). Arch. Mikrobiol. 41, 393.
Tolba, M.K. and Salama, A.M. (1962). Arch. Mikrobiol. 43, 349.
Tong, W., Taurog, A., and ChaikofF, I.L. (1957). J. Biol. Chem. Ill, 713.
Tonhazy, N.E. and Pelczar, M.J., Jr. (1953). J. Bacteriol. 65, 368.
Tonomura, Y, and Furuya, K. (1960). J. Biochem. {Tokyo) 48, 899.
REFERENCES 1061
Tonomura, Y. and Kitagawa, S. (1957). Biochim. Biophys. Acta 26, 15.
Tonomura, Y. and Kitagawa, S. (1960). Biochim. Biophys. Acta 40, 135.
Tonomura, Y. and Yoshimura, J. (1962). J. Biochem. (Tokyo) 51, 259.
Tonomura, Y., Sekiya, K., and Imamura, K. (1963 a). Biochim. Biophys. Acta 69,
296.
Tonomura, Y., Yoshimura, J., and Olmishi, T. (1963 b). Biochim. Biophys. Acta 70,
698.
Topper, Y.J. and Lipton, M.M. (1953). J. Biol. Chem. 203, 135.
Torda, C. and Wolff, H.G. (1944 a). Proc. Soc. Exptl. Biol. Med. 57, 234.
Torda, C. and Wolff, H.G. (1944 b). Proc. Soc. Exptl. Biol. Med. 57, 236.
Totter, J.R. and Best, A.N. (1955). Arch. Biochem. Biophys. 54, 318.
Tower, D.B. (1958). J. Neurochem. 3, 184.
Tower, D.B., Peters, E.L., and Curtis, W.C. (1963). J. Biol. Chem. 238, 983.
Traniello, S. and Vescia, A. (1964). Arch. Biochem. Biophys. 105, 465.
Trim, A.R. (1959). Biochem. J. 73, 298.
Troger, R. (1959). Arch. Mikrobiol. 33, 186.
Trudinger, P.A. and Cohen, G.N. (1956). Biochem. J. 62, 488.
Tsao, T.-C. and Bailey, K. (1953). Biochim. Biophys. Acta 11, 102.
Tschudy, D.P., Colhns, A., Caughey, W.S., and Kleinspehn, G.G. (1960). Science 131,
1380.
Tsen, C.C. and Collier, H.B. (1959). Nature 183, 1327.
Tsen, C.C. and Collier, H.B. (1960). Can. J. Biochem. Physiol. 38, 981.
Tsuboi, K.K. and Hudson, P.B. (1955 a). Arch. Biochem. Biophys. 55, 191.
Tsuboi, K.K. and Hudson, P.B. (1955 b). Arch. Biochem. Biophys. 55, 206.
Tsuboi, K.K. and Hudson, P.S. (1956). Arch. Biochem. Biophys. 61, 197.
Tsuboi, K.K. and Hudson, P.B. (1957). J. Biol. Chem. 224, 889.
Tsuboi, K.K., Penefsky, Z.J., and Hudson, P.B. (1957). Arch. Biochem. Biophys.
68, 54.
Tsuboi, K.K., Estrada, J., and Hudson, P.B. (1958). J. Biol. Chem, 231, 19.
Tsukada, Y. and Takagaki, G. (1955). Nature 175, 725.
Tsukada, Y., Nagata, Y., Hirano, S., and Takagaki, G. (1958). J. Biochem. (Tokyo)
45, 979.
Tsukamoto, H. (1951). J. Pharm. Soc. Japan. 71, 471.
Tsunoda, T. and Shiio, I. (1959). J. Biochem. (Tokyo) 46, 1011.
Tsurumaki, T., Ogiu, K., and Marui, E. (1928). Folia Pharmacol. Japon. 6, 329.
Tsuyuki, E., Tsuyuki, H., and Stahmann, M.A. (1956). J. Biol. Chem. Ill, 479.
Tubbs, P.K. (1962). Biochem. J. 82, 36.
Tubbs, P.K. and Greville, G.D. (1961). Biochem. J. 81, 104.
Turano, C, Giartosio, A., and Riva, F. (1963). Enzymologia 25, 196.
Turano, C, Giartosio, A., and Fasella, P. (1964). Arch. Biocheyn. Biophys. 104, 524.
Turba, F. and Kuschinsky, G. (1952). Biochim. Biophys. Acta 8, 76.
Turner, D.H. and Turner, J.F. (1961). Biochem. J. 79, 143.
Turner, J.S. and Hanly, V. (1947). Nature 160, 296.
Turner, W.A. and Hartman, A.M. (1925). J. Am. Chem. Soc. 47, 2044.
Turpaev, T.M. (1955). Biochemistry (USSE) (English Transl.) 20, 456.
Turrian, H., Grandjean, E., Blattig, K., and Turrian, V. (1956). Helv. Physiol. Phar-
macol. Acta 14, 50.
Tyler, D.B. (1942). Proc. Soc. Exptl. Biol. Med. 49, 537.
1062 REFERENCES
Udaka, S. and Moyed, H.S. (1963). J. Biol. Chem. 238, 2797.
Udenfriend, S. and Cooper, J.R. (1952). J. Biol. Chem. 194, 503.
Udenfriend, S., Creveling, C.R., Ozaki, M., Daly, J.W., and Witkop, B. (1959). Arch.
Biochem. Biophys. 84, 249.
Ukita, T., Tamura, T., Matsuda, R., and Kashiwabara, E. (1949). Japan. J. Exptl.
Med. 20, 109.
Ulbricht, T.L.V. and Gots, J.S. (1956). Nature 178, 913.
Ulrich, F. (1960). Am. J. Physiol. 198, 847.
Ulrich, J. A. and Fitzpatrick, T.B. (1951). Proc. Soc. Exptl. Biol. Med. 76, 346.
Umbreit, W.W. (1955 a). Natl. Vitamin Found. Nutr. Sym.p. Ser. 11, 21.
Umbreit, W.W. (1955 b). In "Amino Acid Metabolism" (W.D. McElroy and H.B.
Glass, eds.), p. 48. Johns Hopkins Press, Baltimore, Maryland.
Umbreit, W.W. and Waddell, J.G. (1949). Proc. Soc. Exptl. Biol. Med. 70, 293.
Umemura, Y., Nishida, J., Akazawa, T., and Uritani, I. (1961). Arch. Biochem. Bio-
phys. 92, 392.
Ungar, G. and Psychoyos, S. (1963). Biochim. Biophys. Acta 66, 118.
Unna, K. (1940). Proc. Soc. Exptl. Biol. Med. 43, 122.
Urata, G. and Granick, S. (1963). J. Biol. Chem. 238, 811.
Urivetzky, M. and Tsuboi, K.K. (1963). Arch. Biochem. Biophys. 103, 1.
Utzinger, G.E., Strait, L.A., and Tuck, L.D. (1963). Experientia 19, 324.
Vagelos, P.R. (1960). J. Biol. Chem. 235, 346.
Vagelos, P.R. and Alberts, A.W. (1960). J. Biol. Chem. 235, 2786.
Vagelos, P.R. and Earl, J.M. (1959). J. Biol. Chem. 234, 2272.
Vallee, B.L., Williams, R.J.P., and Coleman, J.E. (1961). Nature 190, 633.
VanArsdel, P.P. and Bray, R.E. (1961). J. Pharmacol. Exptl. Therap. 133, 319.
Van Baerle, R.R., Goldstein, L., and Dearborn, E.H. (1957). Enzymologia 18, 190.
Vandendriessche, L. (1956). Arch. Biochem. Biophys. 65, 347.
Vander, A.J. (1963). Am. J. Physiol. 204, 781.
Vander, A. J., Malvin, R.L., Wilde, W.S., and Sullivan, L.P. (1958). Am. J. Physiol.
195, 558.
van der Schoot, J.B., Creveling, C.R., Nagatsu, T., and Udenfriend, S. (1963). J.
Pharmacol. Exptl. Therap. 141, 74.
Van Duuren, A.J. (1953). Bee. Trav. Chim. 72, 889.
van Eys, J. (1956). Federation Proc. 15, 374.
van Eys, J. and Kaplan, N.O. (1957 a). Biochim. Biophys. Acta 23, 574.
van Eys, J. and Kaplan, N.O. (1957 b). J. Biol. Chem. 228, 305.
van Eys, J., Ciotti, M.M., and Kaplan, N.O. (1958). J. Biol. Chem. 231, 571.
van Eys, J., Nuenke, B.J., and Patterson, M.K., Jr. (1959). J. Biol. Chem. 234,
2308.
van Eys, J., Kretszchmar, R., Tseng, N.S., and Cunningham, L.W., Jr. (1962). Bio-
chem. Biophys. Res. Commun. 8, 243.
Van Grembergen, G. (1945). Enzymologia 11, 268.
Van Grembergen, G. (1949). Enzymologia 13, 241.
van Heyningen, R. and Pirie, A. (1953). Biochem. J. 53, 436.
van Heyningen, T. and Waley, S.G. (1963). Biochem. J. 86, 92.
Van Oorschot, J.L.P. and Hilton, J.L. (1963). Arch. Biochem. Biophys. 100, 289.
Vanov, S. (1962). Arch. Intern. Pharmacodyn. 138, 51.
REFERENCES 1063
Van Pilsun, J.F. (1953). J. Biol. Chem. 204, 613.
Van Thoai, N. and Olomucki, A. (1962). Biochim. Biophys. Acta 59, 533.
van Vals, G.H. and Emmelot, P. (1957). Z. Krebsforsch. 62, 63.
van Vals, G.H., Bosch, L., and Emmelot, P. (1956). Brit. J. Cancer 10, 792.
Vardanis, A. and Hochster, R.M. (1961). Can. J. Biochem. Physiol. 39, 1695.
Vargas, R. and Cafruny, E.J. (1959). Federation Proc. 18, 455.
Varner, J.E. (1960). Arch. Biochem. Biophys. 90, 7.
Varner, J.E. and Webster, G.C. (1955). Plant Physiol. 30, 393.
Vasington, F.D. (1963). J. Biol. Chem. 238, 1841.
Vasington, F.D. and Murphy, J.V. (1962). J. Biol. Chem. 237, 2670.
Vaslow, F. and Doherty, D.G. (1953). J. Am. Chem. Soc. 75, 928.
Veitch, F.P. and Brierley, G.P. (1962). Biochim. Biophys. Acta 58, 467.
Velick, S.F. (1953). J. Biol. Chem. 203, 563.
VeUck, S.F. (1958). J. Biol. Chem. 233, 1455.
Velick, S.F. and Ronzoni, E. (1948). J. Biol. Chem.. 173, 627.
Velick, S.F. and Vavra, J. (1962). J. Biol. Chem. 137, 2109.
Velluz, L. and Herbain, M. (1951). J. Biol. Chem. 190, 241.
Vennesland, B. and Evans, E.A., Jr. (1944). J. Biol. Chem. 156, 783.
Vennesland, B., Evans, E.A., Jr., and Francis, A.M. (1946). J. Biol. Chem. 163, 575.
Venturi, V.M. and Schoepke, H.G. (1960). Proc. Soc. Exptl. Biol. Med. 103, 455.
Vercauteren, R. (1957). Enzymologia 18, 97.
Vernberg, W.B. and Hunter, W.S. (1960). Exptl. Parasitol. 9, 42.
Vemon, L.P., Mahler, H.R., and Sarkar, N.K. (1952). J. Biol. Chem. 199, 599.
Vescia, A. and di Prisco, G. (1962). J. Biol. Chem. 237, 2318.
Vickery, H.B. (1959). J. Biol. Chem. 234, 1363.
Vickery, H.B. and Palmer, J.K. (1956 a). J. Biol. Chem. 218, 225.
Vickery, H.B. and Palmer, J.K. (1956b). J. Biol. Chem. 12^, 79.
Vickery, H.B. and Palmer, J.K. (1957). J. Biol. Chem. 225, 629.
Villee, C.A. and Loring, J.M. (1961). Biochem. J. 81, 488.
Villela, G.G. (1963). Enzymologia 25, 261.
Villela, G.G., Afifonso, O.R., and Mitidieri, E. (1956). Ezperientia 12, 477.
Vincent, P.O. (1958). Australian J. Exptl. Biol. Med. Sci. 36, 589.
Vincent, P.C. and Blackburn, C.R.B. (1958). Australian J. Exptl. Biol. Med. Sci. 36,
471.
Vishniac, W. (1950). Arch. Biochem. 26, 167.
Vishwakarma, P. (1957). Federation Proc. 16, 132.
Vishwakarma, P. (1962). Am. J. Physiol. 202, 572.
Vishwakarma, P. and Lotspeich, W.D. (1960). Am. J. Physiol. 198, 819.
Vitale, J.J., Jankelson, O.M., Connors, P., Hasted, D.M., and Zamcheck, N. (1956).
Am. J. Physiol. 187, 427.
Vogel, A.I. (1929). J. Chem. Soc. p. 1476.
Vogler, K.G., LePage, G.A., and Umbreit, W.W. (1942). J. Gen. Physiol. 26, 89.
Volk, W.A. (1960). J. Biol. Chem. 235, 1550.
von Bruchhausen, F. (1964). Arch. Exptl. Pathol. Pharmakol. 247, 87.
von Euler, H. (1942). Ber. 75, 1876.
von Euler, H. and Josephson, K. (1923). Z. Physiol. Chem. 127, 99.
von Euler, H. and Svanberg, O. (1920). Fermentforschung 3, 330.
Von Holt, C, Blum, K.U., and Heinrich, W.-D. (1955). Z. Physiol. Chem. 302, 68.
1064 EEFERENCES
Wachstein, M. and Meisel, E. (1954). Science 119, 100.
Wachstein, M. and Meisel, E. (1957). J. Histochem. Cytochem. 5, 204.
Wada, E. and Kobashi, Y. (1953). J. Agr. Chem. 8oc. Japan 27, 561.
Wada, H. and Snell, E.E. (1961). J. Biol. Chem. 236, 2089.
Wada, H., Yoshimatsu, H., Koizumi, T., Inoue, F., Ito, K., Morisue, T., Nasu, H.,
Ito, H., Sakamoto, Y., and Ichihara, K. (1958). Proc. Intern. Symp. Enzyme Chem.,
Tokyo Kyoto 1957 p. 148. Maruzen, Tokyo.
Wadkins, C.L. (1960). J. Biol. Chem. 235, 3300.
Wadkins, C.L. and Lehninger, A.L. (1958). J. Biol. Chem. 233, 1589.
Wadkins, C.L. and Mills, R.C. (1955). Federation Proc. 14, 299.
Wadkins, C.L. and Mills, R.C. (1956). Federation Proc. 15, 377.
Wahl, R. (1939). Compt. Rend. Soc. Biol. 131, 231.
Wakerlin, G.E. and Loevenhart, A.S. (1926). J. Pharmacol. Exptl. Therap. 27, 385.
Wakil, S.J. and Mahler, H.R. (1954). J. Biol. Chem. 207, 125.
Walaas, E. and Walaas, O. (1956). Acta Chem. Scand. 10, 122.
Wald, G. and Brown, P.K. (1951). Federation Proc. 10, 266.
Wald, G. and Brown, P.K. (1952). J. Gen. Physiol. 35, 797.
Wald, G., Brown, P.K., and Smith, P.H. (1955). J. Gen. Physiol. 38, 623.
Waley, S.G. and van Heyningen, R. (1962). Biochem. J. 83, 274.
Walker, D.G. and Rao, S. (1963). Biochim. Biophys. Acta 77, 662.
Walker, E. (1928). Biochem. J. 11, 292.
Walker, G.C. and Nicholas, D.J.D. (1961). Biochem. J. 78, lOp.
Walker, P.G., Woollen, J.W., and Heyworth, R. (1961). Biochem. J. 79, 288.
Walker, T.K., Subramaniam, V., and Challenger, F. (1927). J. Chem. Soc. p. 3044.
Wallace, A. and Mueller, R.T. (1961). Plant Physiol. 36, 118.
Wallace, R.A., Kurtz, A.N., and Niemann, C. (1963). Biochemistry 1, 824.
Wallach, D.P. (1960). Biochem. Pharmacol. 5, 166.
Wallach, D.P. (1961a). Biochem. Pharmacol. 5, 323.
Wallach, D.P. (1961b). Biochem. Pharmacol. 8, 328.
Wallenfels, K. and Miiller-Hill, B. (1964). Biochem. Z. 339, 352.
Wallenfels, K. and Sund, H. (1957 a). Biochem. Z. 329, 17.
Wallenfels, K. and Sund, H. (1957 b). Biochem. Z. 329, 48.
Wallenfels, K., Sund, H., Zarnitz, M.L., Malhotra, O.P., and Fischer, J. (1959). In
"Sulfur in Proteins" (R. Benesch et al., eds.), p. 215. Academic Press, New York.
Wallgren, H. (1960). Acta Physiol. Scand. 49, 216.
Walsh, E. O'F. and Walsh, G. (1948). Nature 161, 976.
Walter, C. (1963). Biochim. Biophys. Acta 77, 525.
Walter, P. and Kaplan, N.O. (1963). J. Biol. Chem. 238, 2823.
Waltman, J.M. and Rittenberg, S.C. (1954). J. Bacteriol. 68, 585.
Wang, T.P. and Kaplan, N.O. (1954). J. Biol. Chem. 206, 311.
Warburg, 0. and Christian, W. (1941). Naturwissenschaften 39, 589.
Warburg, O. and Christian, W. (1942). Biochem. Z. 310, 384.
Warburg, O., Gawehn, K., and Geissler, A.-W. (1957). Z. Naturforsch. 12b, 393.
Warkany, J. and Hubbard, D.M. (1948). Lancet I, 829.
Warner, C. (1951). Australian J. Sci. Res. 4, 554.
Warner, R.C. (1957). J. Biol. Chem. IV), 711.
Warnock, L.G. and van Eys, J. (1963). J. Bacteriol. 85, 1179.
Warringa, M.G.P.J. and Giuditta, A. (1958). J. Biol. Chem. 230, HI.
REFERENCES 1065
Warringa, M.G.P.J., Smith, O.H., Giuditta, A., and Singer, T.P. (1958). J . Biol. Chem.
230, 97.
Washio, S. and Mano, Y. (1960). J. Biochem. (Tokyo) 48, 874.
Watanabe, K. and Yamafuji, K. (1961). Enzymologia 23, 353.
Waterhouse, D.F. (1955). Australian J. Biol. Sci. 8, 514.
Watkins, W.M. (1959). Biochem. J. 71, 261.
Waton, N.G. (1956). Brit. J. Pharmacol. 11, 119.
Watt, D.D. and Werkman, C.H. (1954). Arch. Biochem. Biophys. 50, 64.
Weaver, M.E. (1959). Anat. Record 135, 303.
Webb, E.G. and Morrow, P.F.W. (1959). Biochem. J. 73, 7.
Webb, J.L. (1950 a). Brit. J. Pharmacol. 5, 87.
Webb, J.L. (1950 b). Brit. J. Pharmacol. 5, 335.
Webb, J.L. and Hollander, P.B. (1959). Circulation Res. 7, 131.
Webb, J.L., Saunders, P.R., and Thienes, C.H. (1949). Arch. Biochem. 22, 458.
Webb, J.L.A., Bhatia, I.S., Corwin, A.H., and Sharp, A.G. (1950). J. Am. Chem. Soc.
72, 91.
Weber, H.H. and Portzehl, H. (1954). Progr. Biophys. Biophys. Chem. 4, 60.
Weed, R.I., Eber, J., and Rothstein, A. (1962). J. Oen. Physiol. 45, 395.
Weeks, J.R. and Chenoweth, M.B. (1950). J. Pharmacol. Exptl. Therap. 98, 35.
Weeks, J.R., Chenoweth, M.B., and Shideman, F.E. (1950). J. Pharmacol. Exptl.
Therap. 98, 224.
Weill, C.E. and Caldwell, M.L. (1945 a). J. Am. Chem. Soc. 67, 212.
Weill, C.E. and Caldwell, M.L. (1945 b). J. Am. Chem. Soc. 67, 214.
Weil-Malherbe, H. (1937). Biochem. J. 31, 299.
Weil-Malherbe, H. and Bone, A.D. (1951). Biochem. J. 49, 339.
Weinbach, E.G. (1953). Arch. Biochem. Biophys. 42, 231.
Weinbach, E.G. (1961). J. Biol. Chem. 236, 1526.
Weiner, I.M. and Miiller, C.H. (1955). J. Pharmacol. Exptl. Therap. 113, 241.
Weiner, I.M., Burnett, A.E., and Rennick, B.R. (1956). J. Pharmacol. Exptl. Therap.
118, 470.
Weiner, I.M., Garlid, K., Sapir, D., and Mudge, G.H. (1959). J. Pharmacol. Exptl.
Therap. 127, 325.
Weiner, I.M., Levy, R.I., and Mudge, G.H. (1962). J. Pharmacol. Exptl. Therap. 138,
96.
Weinhouse, S., Millington, R.H., and Friedman, B. (1949). J. Biol. Chem. 181, 489.
Weinhouse, S., Millington, R.H., and Wenner, C.E. (1951). Cancer Res. 11, 845.
Weiss, B. (1951). J. Biol. Chem. 193, 509.
Weitzel, G. and Buddecke, E. (1959). Z. Physiol. Chem. 317, 150.
Weitzel, G. and Schaeg, W. (1959). Z. Physiol. Chem. 316, 250.
Welch, A.D. (1945). Physiol. Rev. 25, 687.
Welch, K. (1962). Am. J. Physiol. 202, 757.
Wells, H.G. (1916). J. Biol. Chem. 26, 319.
Wells, I.e. (1954). J. Biol. Chem. 207, 575.
Welt, L.G., Goodyer, A.V.N., Darragh, J.H., Abele, W.A., and Meroney, W.H. (1953).
J. Appl. Physiol. 6, 134.
Wenner, C.E. and Cereijo-Santalo, R. (1962). Arch. Biochem. Biophys. 98, 67.
Wenner, C.E. and Paigen, K. (1961). Arch. Biochem. Biophys. 93, 646.
Wenzel, D.G. and Siegel, LA. (1962). J. Pharm. Sci. 51, 1074.
1066 REFERENCES
Werkheiser, W.C. (1959). Proc. Am. Assoc. Cancer Res. 3, 72.
Werkheiser, W.C. (1960). Proc. Am. Assoc. Cancer Res. 3, 161.
Werkheiser, W.C. (1961). J. Biol. Chem. 236, 888.
Werkheiser, W.C. (1962). J. Pharmacol. Exptl. Therap. 137, 167.
Werle, E. and Koch, W. (1949). Biochem. Z. 319, 305.
Wessels, J.G.H. (1959). Acta Botan. Neerl. 8, 497.
Wessels, J.S.C. (1958). Biochim. Biophys. Acta 29, 113.
Wessels, J.S.C. (1959). Biochim. Biophys. Acta 35, 53.
Wessels, J.S.C. and Baltscheffsky, H. (1960). Acta Chem. Scand. 14, 233.
Wesson, L.G. (1962). J. Lab. Clin. Med. 59, 630.
Westerfeld, W.W. and Richert, D.A. (1952). J. Biol. Chem. 199, 393.
Westerfeld, W.W., Richert, D.A., and Higgins, E.S. (1959). J. Biol. Chem. 234, 1897.
Westermann, E., Balzer, H., and Knell, J. (1958). Arch. Exptl. Pathol. Pharmakol.
234, 194.
Westhead, E.W. and Boyer, P.D. (1961). Biochim. Biophys. Acta 54, 145.
Westheimer, F.H. and Shookhoff, M.W. (1939). J. Am. Chem. Soc. 61, 555.
Westlake, D.W.S., Shug, A.L., and Wilson, P.W. (1961). Can. J. Microbiol. 7, 515.
White, E.G. and Vernon, L.P. (1958). J. Biol. Chem. 233, 217.
White, H.L., Rolf, D., Bisno, A.L., Kasser, I.S., and Tosteson, D.C. (1961). Am. J.
Physiol. 200, 885.
Whitehouse, M.W., Staple, E., and Gurin, S. (1959). J. Biol. Che^n. 234, 276.
Whiteley, H.R., Osborn, M.J., and Huennekens, F.M. (1959). J. Biol. Chem. 234, 1538.
Whitmore, F.C. and Woodward, G.E. (1941). In "Organic Syntheses" (H. Gilraan
and A.H. Blatt, eds.). Collective Vol. 1, p. 159. Wiley, New York.
Whittingham, C.P. (1956). J. Exptl. Botany 7, 273.
Wick, A.N., Drury, D.R., and Morita, T.N. (1955). Proc. Soc. Exptl. Biol. Med. 89,
579.
Wick, A.N., Wolfe, J.B., and Nakada, H.I. (1956). Proc. Soc. Exptl. Biol. Med. 93,
150.
Wick, A.N., Drury, D.R., Nakada, H.I., and Wolfe, J.B. (1957). J. Biol. Chem. 224,
963.
Wick, A.N., Serif, G.S., Stewart, C.J., Nakada, H.I., Larson, E.R., and Drury, D.R.
(1959). Diabetes 8, 112.
Wickson-Ginzburg, M. and Solomon, A.K. (1963). J. Gen. Physiol. 46, 1303.
Wieland, O. and Weiss, L. (1963). Biochem. Biophys. Res. Commun. 13, 26.
Wien, R. (1939). Quart. J. Pharm. Pharmacol. 12, 212.
Wiesmeyer, H. and Colin, M. (1957). Federation Proc. 16, 270.
Wiesmeyer, H. and Cohn, M. (1960). Biochim. Biophys. Acta 39, 427.
Wiethoff, E.G., Leppla, W., and Holt, C.V. (1957). Biochem. Z. 328, 576.
Wigler, P.W. and Alberty, R.A. (1960). J. Am. Chem. Soc. 82, 5482.
Wilbrandt, W. (1941). Arch. Ges. Physiol. 244, 637.
Wilbrandt, W. and Rosenberg, T. (1961). Pharmacol. Rev. 13, 109.
Wilde, C.E. and Kekwick, R.G.O. (1964). Biochem. J. 91, 297.
Willard, H.H. and Young, P. (1930). J. Am. Chem. Soc. 52, 132.
WilHams, A.D., Slater, G.G., and Winzler, R.J. (1955). Cancer Res. 15, 532.
Williams, A.K. and Eagon, R.G. (1959). J. Bacteriol. 77, 167.
Williams, C.H., Jr. and Kamin, H. (1962). J. Biol. Chem. 237, 587.
Williams, J.N., Jr. (1952). J. Biol. Chem. 195, 629.
REFERENCES 1067
Williams, R.J.P. (1956). Chem. Rev. 56, 299.
Willing, F., Neuhoff, V., and Herken, H. (1964). Arch. Exptl. Pathol. Pharmakol. 247,
254.
Wills, E.D. (1959). Biochem. Pharmacol. 2, 276.
Wills, E.D. (1960). Biochim. Biophys. Acta 40, 481.
Wilson, A.N. and Harris, S.A. (1949). J. Am. Chem. Soc. 71, 2231.
Wilson, G.B., Morrison, J.H., and Knobloch, N. (1959). J. Biophys. Biochem. Cytol.
5, 411.
Wilson, J.E., Clark, W.C, and Hulpieu, H.R. (1962). J. Pharmacol. Exptl. Therap.
137, 179.
Wilson, P.W., Burris, R.H., and Lind, C.J. (1942). Proc. Natl. Acad. Sci. U. S. 28.
243.
Wilson, T.G.G. and Roberts, E.R. (1954). Biochim. Biophys. Acta 15, 568.
Wilson, T.H. and Landau, B.R. (1960). Am. J. Physiol. 198, 99.
Wilson, T.H., Lin, E.C.C., Landau, B.R., and Jorgenson, C.R. (1960). Federation Proc
19, 870.
Winer, A.D. (1960). Biochem. J. 76, 5p.
Winer, A.D., Schwert, G.W., and Millar, D.B.S. (1959). J. Biol. Chem. 234, 1149.
Winestock, C.H., Aogaichi, T., and Plaut, G.W.E. (1963). J. Biol. Chem. 238, 2866.
Wingo, W.J. and Awapara, J. (1950). J. Biol. Chem. 187, 267.
Winzor, D.J. and Creeth, J.M. (1962). Biochem. J. 83, 559.
Wiseman, G. (1954). J. Physiol. {London) 124, 414.
Wiseman, M.H. and Sourkes, T.L. (1961). Biochem. J. 78, 123.
Wiskich, J.T., Morton, R.K., and Robertson, R.N. (1960). Australian J. Biol. Sci.
13, 109.
Wislicenus, J. (1874). Ber. 7, 683.
Witter, A. (1960). Acta Chem. Scand. 14, 1717.
Witter, A. and Tuppy, H. (1960). Biochim. Biophys. Acta 45, 429.
Witter, R.F., Newcomb, E.H., and Stotz, E. (1950). J. Biol. Chem. 185, 537.
Wockel, W., Stegner, H.-E., and Janisch, W. (1961). Arch. Pathol. Anaf. Physiol. 334,
503.
Wohl, A. and Glimm, E. (1910). Biochem. Z. 27, 349.
Wojtczak, L., Wlodawer, P., and Zborowski, J. (1963). Biochim. Biophys. Acta 70,
290.
Wolbach, R.A. and Riggs, A. (1955). Federation Proc. 14, 306.
Wold, F. and Ballou, C.E. (1957). J. Biol. Chem. 117, 313.
Wolf, P. A. (1950). Food Technol. 4, 294.
Wolf, P.A. and Westveer, W.M. (1950). Arch. Biochem. 28, 201.
Wolfe, J.B. (1955). Arch. Biochem. Biophys. 57, 414.
Wolfe, J.B. and Nakada, H.I. (1956). Arch. Biochem. Biophys. 64, 489.
Wolfe, J.B. and Rittenberg, S.C. (1954). J. Biol. Chem. 209, 885.
Wolfe, J.B., Ivler, D., and Rittenberg, S.C. (1954 a). J. Biol. Chem. 209, 867.
Wolfe, J.B., Ivler, D., and Rittenberg, S.C. (1954 b). J. Biol. Chem. 209, 875.
Wolfe, J.B., Ivler, D., and Rittenberg, S.C. (1955). J. Bacteriol. 69, 240.
Wolfe, R.G. and Neilands, J.B. (1956). J. Biol. Chem. ll'i, 61.
Wolfe, S.J. (1957). J. Biol. Chem. 229, 801.
Wolff, J.B. and Resnik, R.A. (1963). Biochim. Biophys. Acta 73, 588.
Wood, H.G. and Stjernholm, R. (1961). Proc. Natl. Acad. Sci. U. S. 47, 289.
1068 EEFERENCES
Wood, H.G. and Werkman, C.H. (1940). Biochem. J. 34, 7.
Wood, J.D. (1959). Can. J. Biochem. Physiol. 37, 937.
Wood, R.C. and Hitchings, G.H. (1959 a). J. Biol. Chem. 234, 2377.
Wood, R.C. and Hitchings, G.H. (1959 b). J. Biol. Chem. 234, 2381.
Woodruff, L.L. and Bunzel, H.H. (1910). Am. J. Physiol. 25, 190.
Woods, D.D. (1940). Brit. J. Exptl. Pathol. 21, 74.
Woods, L.A., Shideman, F.E., Seevers, M.H., Weeks, J.R., and Kruse, W.T. (1950).
J. Pharmacol. Exptl. Therap. 99, 84.
Woodward, G.E. and Hudson, M.T. (1954). Cancer Pes. 14, 599.
Woody, B.R. and Lindstrom, E.S. (1955). J. Bacteriol. 69, 353.
Woolley, D.W. (1944 a). Proc. Soc. Exptl. Biol. Med. 55, 179.
Woolley, D.W. (1944 b). J. Biol. Chem. 154, 31.
Woolley, D.W. (1945 a). Proc. Soc. Exptl. Biol. Med. 60, 225.
Woolley, D.W. (1945 b). J. Biol. Chem. 157, 455.
Woolley, D.W. (1950 a). J. Biol. Chem. 191, 43.
Woolley, D.W. (1950 b). Ann. N. Y. Acad. Sci. 52, 1235.
Woolley, D.W. (1951). J. Biol. Chem. 191, 43.
Woolley, D.W. (1952). "A Study of Antimetabolites." Wiley, New York.
Woolley, D.W. and Merrifield, R.B. (1952). Federation Proc. 11, 458.
Woolley, D.W. and Pringle, A. (1951). Federation Proc. 10, 272.
Woolley, D.W. and White, A.G.C. (1943 a). Proc. Soc. Exptl. Biol. Med. 52, 106.
Woolley, D.W. and White, A.G.C. (1943 b). J. Biol. Chem. 149, 285.
Woolley, D.W^ and White, A.G.C. (1943 c). J. Exptl. Med. 78, 489.
Woolley, D.W., Strong, F.M., Madden, T.J., and Elvehjem, C.A. (1938). J. Biol. Chem.
124, 715.
Work, T.S. and Work, E. (1948). "The Basis of Chemotherapy." Wiley (Interscience),
New York.
Wormser, E.H. and Pardee, A.B. (1958). Arch. Biochetn. Biophys. 78, 416.
Wortman, B. (1962). Arch. Biochem. Biophys. 97, 70.
Wosilait, W.D. (1960). J. Biol. Chem. 235, 1196.
Wosilait, W.D. and Nason, A. (1954). J. Biol. Chem. 206, 255.
Wright, C.I. and Sabine, J.C. (1944). J. Biol. Chem. 155, 315.
Wright, C.I. and Sabine, J.C. (1948). J. Pharmacol. Exptl. Therap. 93, 230.
Wright, N.C. (1926). Biochem. J. 20, 524.
Wriston, J.C, Jr., Lack, L., and Shemin, D. (1955). J. Biol. Chem. 215, 603.
Wu, H.L.C. and Mason, M. (1964). J. Biol. Chem. 239, 1492.
Wu, L.-C. and Scheffer, R.P. (1960). Plant Physiol. 35, 708.
Wyatt, H.V. (1964). J. Theoret. Biol. 6, 441.
Wylie, D.W., Archer, S., and Arnold, A. (1960). J. Pharmacol. Exptl. Therap. 130,
239.
Wyngaarden, J.B. (1957). J. Biol. Chem. 224, 453.
Wyngaarden, J.B. and Ashton, D.M. (1959). Federation Proc. 18, 174.
Wyngaarden, J.B. and Greenland, R.A. (1963). J. Biol. Chem. 238, 1054.
Wynn, J. and Gibbs, R. (1963). J. Biol. Chem. 238, 3490.
Wyss, O. and Strandskov, F.B. (1945). Arch. Biochem. 6, 261.
Yagi, K. and Nagatsu, I. (1960). J. Biochem. (Tokyo) 47, 630.
Yagi, K. and Natsume, K. (1964). J. Biochem. (Tokyo) 55, 529.
REFERENCES 1069
Yagi, K. and Ozawa, T. (1959). Nature 184, 1227.
Yagi, K., Okuda, J., Ozawa, T., and Okada, K. (1957). Biochem. Z. 328, 492.
Yagi, K., Ozawa, T., and Okada, K. (1959). Biochim. Biophys. Acta 35, 102.
Yagi, K., Ozawa, T., and Harada, M. (1960). Nature 188, 745.
Yamada, E.W. and Jakoby, W.B. (1959). J. Biol. Chem. 234, 941.
Yamada, K. (1959). J. Biochem. {Tokyo) 46, 361.
Yamada, T. and Yanagita, T. (1957). J. Biochem. (Tokyo) 44, 661.
Yamamura, Y., Kusunose, M., and Kusunose, E. (1952). J. Biochem. {Tokyo) 39, 227.
Yamamura, Y., Kusunose, M., Nagai, S., and Kusunose, E. (1954). J. Biochem. {Tokyo)
41, 513.
Yamane, T. and Davidson, N. (1961). J. Am. Chem. Soc. 83, 2599.
Yamashita, J. (1960). J. Biochem. {Tokyo) 48, 525.
Yamauchi, M. (1956). Nippon Saikingaku Zasshi 11, 221.
Yanari, S. and Mitz, M.A. (1957). J. Am. Chem. Soc. 79, 1154.
Yang, W.C.T. (1957). Science 125, 1087.
Yeas, M.F. (1950). Ph.D. thesis, CaHf. Inst. Technol., Pasadena, California.
Yeas, M.F. (1954). J. Exptl. Biol. 31, 208.
Yee, R.B., Pan, S.F., and Gezon, H.M. (1958). J. Bacteriol. 75, 51.
Yonetani, T. and Theorell, H. (1962). Arch. Biochem. Biophys. 99, 433.
Yonetani, T. and Theorell, H. (1964). Arch. Biochem. Biophys. 106, 243.
Yoneya, T. and Adams, E. (1961). J. Biol. Chem. 236, 3272.
Yoshiba, A., Nakao, K., Minakami, S., and Yoneyama, Y. (1958). J. Biochem. {Tokyo)
45, 913.
Yoshida, F. and Nagasawa, M. (1958). Proc. Intern. Symp. Enzyme Chem., Tokyo
Kyoto 1957 p. 504. Maruzen, Tokyo.
Yoshida, H. and Quastel, J.H. (1962). Biochim. Biophys. Acta 57, 67.
Young, R.H. and Shannon, L.M. (1959). Plant Physiol. 34, 149.
Yushok, W.D. (1964). Cancer Res. 24, 187.
Yushok, W.D. and Batt, W.G. (1957). Ahstr. Am. Chem. Soc, 132nd Meeting, 1957
pp. 23C-24C.
Zahl, P.A. and Albaum, H.G. (1955). Proc. Soc. Exptl. Biol. Med. 88, 263.
Zakrzewski, S.F. (1963). J. Biol. Chem. 238, 1485.
Zakrzewski, S.F. and Nichol, C.A. (1958). Biochim. Biophys. Acta 27, 425.
ZambonelH, C. (1958 a). Ann. Microbiol. Enzimol. 8, 27.
ZamboneUi, C. (1958 b). Ann. Microbiol. Enzimol. 8, 98.
Zannoni, V.G. and La Du, B.N. (1959). J. Biol. Chem. 234, 2925.
Zatman, L.J., Kaplan, N.O., and Colowick, S.P. (1953). J. Biol. Chem. 200, 197.
Zatman, L.J., Kaplan, N.O., Colowick, S.P., and Ciotti, M.M. (1954 a). J. Biol. Chem.
209, 453.
Zatman, L.J., Kaplan, N.O., Colowick, S.P., and Ciotti, M.M. (1954 b). J. Biol. Chem.
209, 467.
Zelitch, I. (1955). J. Biol. Chem. 216, 553.
Zelitch, I. (1957). J. Biol. Chem. 224, 251.
Zelitch, I. (1958). Federation Proc. 17, 341,
Zelitch, I. (1959). J. Biol. Chem. 234, 3077.
Zeller, E.A. (1938). Helv. Chim. Acta 21, 1645.
Zeller, E.A. (1940). Helv. Chim. Acta 23, 1418.
1070 REFERENCES
Zeller, E.A. (1941). Helv. Chim. Acta 24, 539.
Zeller, E.A. and Maritz, A. (1944). Helv. Chim. Acta 27, 1888.
Zeller, E.A. and Maritz, A. (1945). Helv. Chim. Acta 28, 365.
Zellweger, H. and Wehrli, S. (1951). Helv. Paediat. Acta 6, 397.
Ziegler, D.M. (1961). In "Biological Structure and Function" (T.W. Goodwin and 0.
Lindberg, eds.), Vol. 2, p. 253. Academic Press, New York.
ZiflF, M. (1944). J. Biol. Chem. 153, 25.
Zimmerman, A.M., Landau, J.V., and Marsland, D. (1957). J. Cellular Comp. Physiol.
49, 395.
Zimmerman, S.B. and Kornberg, A. (1961). J. Biol. Chem. 236, 1480.
Zipkin, I., and McClure, F.J. (1957). J. Am. Dental Assoc. 55, 15.
Zipkin, I. and McClure, F.J. (1958). Proc. Soc. Exptl. Biol. Med. 97, 318.
Zittle, C.A. (1945). J. Biol. Chem. 160, 527.
ZoUner, N. and Fellig, J. (1952). Naturwissenschaften 39, 523.
Zollner, N. and Fellig, J. (1953). Am. J. Physiol. 173, 223.
ZoUner, N. and Rothemund, E. (1954). Z. Physiol. Chem. 298, 97.
Zygmunt, W.A. (1963). J. Bacteriol. 85, 1217.
I
AUTHOR INDEX
Numbers in italics refer to pages on which the complete references are listed.
Abadom, P. N., 267, 9S7
Abe, S. 982, 987
Abele, W. A., 925, 1065
Abeles, R. H., 590, 1030
Abood, L. G., 847, 873, 878, 987, 990
Abraham, E. P., 599, 987
Abraham, S., 146, 147, 273, 987, 997
Abul-Fadl, M. A. M., 440, 987
Ackermann, W. W., 30, 42, 127, 175, 193,
309, 494, 987
Acland, J. D., 910, 987
Actis, A. S., 687, 712, 716, 775, 783, 810,
853, 1058
Ada, G. L., 851, 987
Adams, E., 269, 334, 355, 549, 775, 783,
857, 987, 1013, 1055, 1069
Adams, R., 618, 1047
Adell, B., 3, 8, 987
Adinolfi, A., 615, 616, 1049
Adkins, H., 619, 987
Adler, E., 61, 668, 987, 998
Alfonso, D. R., 289, 1063
Agarwala, S. C, 692, 831, 987
Agosin, M., 18, 25, 28, 173, 203, 383, 406,
407, 411, 839, 843, 854, 882, 987, 1035
Agranoflf, B. W., 887, 987
Ahmed, K., 266, 987
Aikawa, J. K., 959, 987
Aikawa, T., 678, 727, 1022
Airan, J. W., 15, 987
Aisenberg, A. C, 126, 128, 987
Ajl, S. J., 26, 79, 177, 187, 242, 987, 1043
Akabori, S., 416, 417, 1025
Akamatsu, S., 428, 987
Akatsu, S., 983, 1042
Akawie, R. I., 310, 1016
Akazawa, T., 439, 1062
Akiba, T., 983, 984, 987
Alanis, J., 724, 987
Albaum, H. G., 169, 196, 228, 585, 987,
1069
Albers, R. W., 601, 856, 987
Albert, A., 259, 261, 972, 973, 988
Alberts, A. W., 847, 1067
Alberty, R. A., 274, 277, 278, 1066
Albrecht, A. M., 590, 988
Albrecht, M., 151, 1036
Albright, E. C., 602, 1030
Alburn, H. E., 459, 988
Aleem, M. I. H., 547, 551, 988
Alex, T., 910, 913, 1050
Alivisatos, S. G. A., 485, 487, 492, 503,
988
Allen, E. H., 307, 1052
Allen, F. W., 712, 1003
Allen, M. B., 892, 989
Allen, R. P., 698, 699, 700, 1012, 1045
AUfrey, V. G., 32, 189, 393, 622, 623, 1034
Allison, A. C., 756, 977, 978, 979, 980,
981, 988
Almkvist, J., 950, 988
Alpert, N. R., 446, 1039
AltareUi, V. R., 960, 997
Altekar, W. W., 274, 988, 1047
Altszuler, N., 401, 988
Alvarado, F., 414, 1056
Alvarez-O'Bourke, F., 505, 1047
Amaha, M., 546, 988
Ames, S. R., 593, 659, 662, 988
Anagnostopoulos, C., 63, 440, 998
Anastasi, A., 671, 1027
Anderson, B. M,, 497, 988
Anderson, D. G., 832, 886, 988, 991
Anderson, D. O., 464, 1021
Anderson, E. P., 473, 480, 988, 996
1071
1072
AUTHOR INDEX
Anderson, H. L., 217, 1006
Anderson, J. A., 38, 1003
Anderson, K. S., 625, 992
Anderson, M. V., Jr., 625, 992
Andrews, C. H., 977, 978, 979, 988
Andrews, G. S., 55, 179, 228, 988
Anfinsen, C. B., 708, 826, 840, 863, 1043
Angielski, S., 206, 988
Anichkov, S. V., 212, 988
Annau, E., 138, 179, 183, 988
Annison, E. F., 77, 1052
Anson, M. L., 661, 670, 671, 672, 680,
681, 690, 697, 754, 979, 980, 988, 1038
Anthony, D. D., 817, 820, 988
Anthony, W. L., 495, 992
Antonin, S., 657, 1003
Antopoll, W., 58, 1009
Aogaichi, T., 539, 543, 1067
Aposhian, H. V., 539, 956, 988
App, A. A., 831, 988
Appleton, J. M., 613, 1044
Applewhite, T. H., 374, 988
Appleyard, G., 836, 988
Ap Rees, T., 51, 53, 169, 172, 1016
Araki, K., 126, 988
Aravena, L., 383, 411, 843, 854, 987
Arbuthnott, J. P., 900, 901, 988
Archer, S., 611, 612, 1068
Archibald, R. M., 550, 988
Ardao, M. I., 92, 722, 882, 991
Arkin, A., 701, 727, 728, 989
Armstrong, J. McD., 787, 989
Arndt, F., 620, 989
Arnold, A., 449, 611, 612, 1008, 1068
Arnon, D. I., 892, 989
Arrigoni-MarteUi, E., 236, 1028
Asahi, T., 543, 554, 989
Asano, A., 552, 864, 989
Asano, N., 195, 1015
Ashton, D. M., 474, 480, 1068
Ashton, G. C, 422, 1019
Ashwell, G., 414, 465, 1004
Asnis, R. E., 26, 989
Astrup, T., 464, 465, 989
Atkinson, D. E., 293, 512, 851, 892, 989,
1030, 1034
Atkinson, M. R., 471, 481, 510, 989
Aubel, E., 52, 60, 74, 81, 989
Audereau, A., 62, 63, 81, 82, 1033
Auditore, G. V., 936, 989
Augustinsson, K. B., 817, 989
Austen, K. F., 374, 989
Austin, J. H., 956, 1060
Avery, G. S., Jr., 53, 60, 81, 596, 992
Avi-Dor, Y., 20, 29, 61, 80, 844, 848,
864, 872, 878, 989, 998, 1013
Avigad, G., 421, 1018
Avron, M., 74, 78, 79, 81, 82, 91, 557, 851,
891, 892, 989, 1014, 1022
Awapara, J., 328, 1067
Axelrod, A. E., 35, 36, 589, 989, 1059
Axelrod, B., 238, 411, 440, 852, 854, 887,
989, 1015, 1026, 7046
Axelrod, J., 590, 5*^/2, 597, 611, 843, 989,
996
Ayengar, P., 336, 989
Ayers, J., 639, 1027
Azarkh, R. M., 360, 995
Azoulay, E., 831, 832, 989, 1018
Azzone, G. F., 17, 18, 33, 121, 122, 865,
989, 1030
B
Bach, M. K., 169, 881, 989
Bach, S. J., 335, 989
Bach, Z. M., 611, 938, 989
Bachhawat, B. K., 816, 1044
Bachmann, H., 949, 989
Bachrach, H. L., 176, 181, 194, 401,
996, 1045
Bacila, M., 550, 1001
Bacr, J. E., 917, 993
Baer, H., 783, 854, 989
Baernstein, H. D., 696, 697, 989
Bassler, K. H., 601, 1030
Bagley, E. H., 62, 85, 87, 1013
Bagot, A. E., 660, 1014
Bahn, R. C, 926, 989
Bailey, J. H., 973, 983, 1027
Bailey, K., 684, 691, 692, 704, 705, 721,
722, 723, 865, 876, 938, 939, 989, 1061
Bain, J. A., 567, 568, 990
Baker, E. E., 196, 1040
Baker, G. D., 495, 992
Baker, N., 695, 990
Baker, R. S., 852, 989
AUTHOR INDEX
1073
Baker, Z., 500, 990
Baldwin, D. M., 976, 979, 980, 1028
Baldwin, E., 54, 174, 990
Balinsky, D., 349, 593, 604, 607, 990
Balis, M. E., 585, 990
Ball, H. A., 388, 990
Ballou, C. E., 408, 409, 1008, 1067
Balls, A. K.i 657, 667, 668, 990, 1023
Baltscheffsky, H., 445, 548, 557, 864,
990, 1066
Baltscheffsky, M., 163, 168, 557, 892.
990, 1055
Balzer, H., 314, 315, 1066
Bamberger, J. W., 198, 1038
Bandelin, F. J., 633, 990
Bandurski, R. S., 71, 172, 189, 543,
554, 852, 989, 990, 995
Banga, I., 75, 175, 181, 187, 990
Banks, J., 497, 1036
Bagtist, J. N., 437, 990
Baralt-Perez, J., 978, 980, 1044
Barany, K., 707, 939, 940, 990
Bardny, M., 663, 664, 704, 707, 723, 803,
806, 939, 940, 990
Barban, S., 32, 388, 389, 391, 400, 990
Barbato,'L. M., 847, 990
Barber, G. A., 137, 138, 389, 990, 1058
Barbour, H. G., 956, 990
Barchas, J., 310, 1033
Bargoni, N., 547, 549, 559, 990
Barker, H. A., 282, 287, 288, 995
Barker, S. B., 18, 783, 990
Barlow, A. J. E., 168, 169, 190, 999
Barman, T. E., 630, 990
Barnabas, J., 15, 987, 990
Barnard, E. A., 807, 906, 990
Barnes, H., 730, 735, 962, 990
Barnes, J. M., 225, 990
Barnes, M. N., 1046
Barnett, R. C, 117, 198, 990, 996
Barone, J. A., 531, 990
Barrett, F. R., 953, 990
Barron, E. S. G., 29, 41, 60, 74, 92, 105,
119, 121, 171, 174, 189, 227, 228, 643,
666, 667, 668, 672, 673, 705, 707, 712,
713, 714, 717, 718, 722, 781, 803, 805,
826, 831, 840, 846, 852, 855, 856, 857,
866, 870, 873, 876, 878, 882, 883, 991,
1032, 1046, 1049, 1054
Barry, R. J. C, 387, 991
Bartett, W. L., 437, 998
Bartlett, G. R., 341, 991
Bartlett, M. D., 810, 835, 870, 1001
Barton, M. N., 577, 1040
Bartley, W., 88, 991
Basciak, J., 988
Basford, R. E., 543, 553, 711, 848, 851,
1020
Bass, L., 946, 1011
Bassham, J. A., 163, 991
Bastarrachea, F., 832, 991
Batt, W. G., 385, 1069
Battaglia, F, C, 264, 911, 991
Bauchop, T., 432, 593, 600, 853, 991
Bauer, C. W., 622, 991
Bauer, S., 137, 1037
Bauer, W., 686, 1048
Bauerle, R. H., 972, 992
Baugh, C. L., 852, 991
Baum, H. M., 284, 713, 1035
Baumann, C. A., 55, 56, 79, 81, 164, 175,
176, 530, 991, 1040, 1057
Bauminger, B., 657, 658, 1032
Baur, H., 269, 353, 1005
Baxter, C. F., 64, 857, 991
Baxter, R. M., 589, 991
Bayne, S., 384, 991, 1038
Beach, J., 399, 400, 1056
Bean, R. C, 385, 991
Beaton, G. H., 566, 991
Beaton, J. R., 571, 573, 991
Beattie, D. S., 334, 810, 856, 1052
Beck, G. E., 938, 991
Beck, S. D., 22, 29, 1016
Becker, B., 209, 267, 465, 991
Becker, C. E., 383, 384, 385, 991
Becker, M., 269, 353, 1005
Becker, V., 18, 219, 220, 991
Beckmann, F., 212, 991
Beechey, R. B., 121, 991
Beers, R. F., Jr., 463, 991, 1018
Beevers, H., 61, 77, 90, 164, 171, 172, 185,
189, 191, 274, 394, 591, 848, 991, 992,
998, 1013, 1023, 1034
Behal, F. J., 195, 782, 992
1074
AUTHOR INDEX
Beher, W. T., 494, 495, 504, 992, 1010
Behne, I., 396, 1019
Beiler, J. M., 586, 587, 1036
Bein, H. J., 938, 991
Beinert, H., 138, 143, 233, 535, 668, 687,
712, 853, 992, 999, 1028
Beling, C. G., 464, 992
Belkin, M., 201, 670, 992
Bell, F. E., 831, 992
Bellin, 225, 992
Bellinger, S. B., 913, 1050
Bello, L. J., 475, 992
Benedict, F. G., 951, 998
Benesch, R. E., 638, 640, 645, 646, 746,
757, 763, 902, 903, 931, 932, 992
Ben-Gershom, R., 383, 1044
Benigno, P., 983, 984, 992
Bennett, D. R., 625, 992
Bennett, E. D., 972, 992
Bennett, H. S., 766, 767, 992
Bennett, L. L., Jr., 585, 1055
Benson, A., 888, 1032
Benson, A. A., 991
Bentley, L. E., 225, 992
Bentley, R., 226, 228, 678, 992
Ben-Zvi, E., 739, 1008
Beppu, T., 844, 992
Ber, A., 532, 992
Berends, W., 538, 1030
Beresotskaya, N. A., 395, 1006
Berg, v., 875, 1028
Bergel, F., 516, 992
Berger, A., 457, 1024
Berger, J., 53, 60, 81, 596, 992
Berger, M., 119, 992
Bergersen, F. J., 292, 294, 992
Bergquist, P. L., 881, 908, 993
Bergkvist, R., 687, 992
Bergmann, F., 281, 285, 816, 992, 993
Bergmeyer, H.-U., 841, 993
Berk, S., 228, 993
Berkowitz, J., 333, 1012
Berl, S., 574, 1046
Bertin, M., 958, 960, 993
Berliner, R. W., 917, 1043
Berman, D. A., 13, 75, 131, 191, 215, 216,
228, 941, 943, 993, 1001, 1036, 1048
Bernath, P., 18, 713, 783, 825, 856, 1054
Bernfeld, P., 454, 459, 463, 993
Bernhard, S. A., 375, 993
Bemheini, A. I., 724, 993
Bernheim, F., 225, 228, 239, 259, 272,
693, 694, 888, 993, 1003, 1040
Bernstein, D. E., 52, 74, 81, 993
Bernstein, G. S., 391, 400, 993
Bernstein, I. A., 413, 1055
Berry, L. J., 203, 221, 222, 223, 228, 993
Bersin, T., 686, 693, 993
Bertelsen, K., 817, 1017
Bertino, J. R., 581, 993
Besemer, A. R., 971, 973, 1000
Bessey, O. A., 286, 287, 288, 289, 1033
Bessman, M. J., 475, 992
Bessman, S. P., 433, 1008
Best, A. N., 585, 1061
Betzel, R., 974, 1018
Beuzeville, C., 221, 223, 228, 993
Beyer, K. H., 264, 917, 993
Beyer, R. E., 540, 544, 547, 548, 993
Bhatia, I. S., 213, 747, 1065
Bhausar, M. D., 415, 1050
Bhuvaneswaran, C, 478, 993
Bhuyan, B. K., 713, 859, 993
Biale, J. B., 74, 78, 89, 81, 82, 91, 120,
173, 182, 989, 1032, 1054
Bickers, J. N., 926, 927, 993
Bickis, I. J., 156, 1046
Bide, R. W., 553, 774, 851, 993
Bieber, R. E., 497, 1024
Bieber, S., 283, 1006
Bieleski, R. L., 910, 993
Bielinski, T. C, 587, 995
Bilinski, E., 137, 994
Bilodeau, F., 212, 994
Bilse, I. M., 632, 1057
Binkley, F., 802, 1052
Bird, H. R., 530, 1040
Birkenhager, J. C., 674, 676, 677, 994
Birmingham, M. K., 150, 1051
Birt, L. M., 74, 75, 79, 82, 171, 1002
Bishop, J. S., 401, 988
Bisno, A. L., 937, 1066
Biszku, E., 649, 788, 803, 812, 817, 827,
1059
Bivings, L., 954, 994
Bjerrum, J., 732, 739, 994
AUTHOR IXDEX
1075
Black, M. M., 202, 994
Black, R. E., 391, 400, 993
Black, S., 832, 994
Blackburn, C. R. B., 902, 908, 1063
Blattig, K., 956, 1061
Blair, D. G. R., 473, 479, 994
Blakley, E. R., 403, 404, 542, 839, 994,
999
Blakley, R. L., 137, 581, 585, 994
Blanchard, M., 60, 338, 348, 994
Blaschko. H., 308, 363, 364, 592, 694,
835, 994
Blaylock, B. A., 551, 843, 994
Blessing, J. A.. 315, 1057
Blickenstaflf. D. D., 985, 994
Bloch, D. I., 780, 798, 865, 1050
Bloch-Frankenthal, L., 383, 396, 1047
Blum, J. J., 445, 446, 816, 819, 867, 994
Blum, K. U.. 520, 1063
Blumberg, A. J., 959, 987
Blumenstein, J., 709, 994
Blumenthal, H. J., 356, 440, 443. 984,
994, 1012, 1029
Blumson, N. L., 332, 994
Bocher, C. A., 462, 1043
Bock, K. ,R., 966, 994
Bock, M., 977, 978, 1013
Bock, R. M., 549, 832, 844, 1035, 1050
Bodansky. O., 413. 994
Boeri, E., 65, 435, 437, 552, 708, 870, 994
Boese, A. B., 620, 1057
Bogdonoff, M. D., 399, 1030
Boger, W. P., 626, 1051
Boggiano, E. M., 356, 1055
Eokman, A. H., 136, 1021
Bolker, H., 202, 994
Bomford, R. R., 951, 952, 953, 954, 1021
Bonaduce, L., 469, 675, 706, 1051
Bone, A. D., 379, 1065
Bone, D. H., 878, 994
Boney, A. D., 967, 994
Bonner, B. A., 27, 168, 994
Bonner, D. M., 418, 595, 1023. 1031
Bonner, J., 53, 58, 78, 80, 93, 120, 167,
169, 170, 172, 182, 184, 189, 191, 297,
516, 911, 967, 994, 995, 996, 1000, 1023
Bonner, W. D., Jr., 32, 33, 196, 966, 968,
995, 1055, 1060
Bonnichsen, R., 784, 1060
Bonsignore, A., 412, 995
Booth, A. X., 601, 995
Borghgraef, R. R. M., 923, 928, 929, 960,
995
Borries, E., 570, 1004
Borsook, H., 887, 995
Borst, P., Ill, 152, 995
Bortz, W. M., 614, 995
Bosch, L., 130, 1063
Bose, S. M., 682, 687, 693, 1003
Boser, H., 837, 995
Bosund, I., 349, 995
Boucek, R. J., 215, 995
Bovarnik, M. R., 341, 347. 547, 549, 556,
557, 558, 560, 780, 995, 1017
Bowman, E. M.. 772, 827. 828, 838, 1011
Bowman, I. B. R., 91, 127, 560, 995
Boxer, G. E., 587, 995
Boyd, E. S., 54, 58, 166, 175, 995
Boyer, P. D., 351, 403, 404, 635, 640,
643, 645, 649, 714, 745, 760, 763, 764,
765, 766, 780, 788, 803, 806, 810, 812,
833, 853. 994, 995, 1025, 1027, 1052,
1059, 1066
Boyland, E., 138, 140, 177, 179, 201, 218,
224, 225, 428, 990, 995
Boyland, M. E., 138, 140, 177, 179, 995
Boylen, J. B., 305, 995
Bradley, L. B., 807, 816, 856, 866, 873,
1022, 1026
Bradley, R. M., 614, 1048
Bradshaw, W. H., 282, 287, 288, 995
Brady, C. J., 693, 995
Brady, R. O., 148, 613, 614, 950, 995,
1048
Brand, E., 665, 666, 667, 995
Branster, M. V., 485, 493, 995
Brasted, R. C, 736, 1055
Braun, K., 494, 995
Braune, W., 660, 995
Braunstein, A. E., 360, 561, 995
Bray, R. E., 725, 949, 995, 1062
Bregoff, H. M., 293, 996
Bremer, J.. 663, 1006
Brendel, R., 586, 587, 1036
Brenner, B. M., 472, 1052
Bresler, E. H., 926, 927, 993
1076
AUTHOR INDEX
Breslow, E., 542, 547, 550, 781, 1035
Bresnick, E., 429, 468, 469, 480, 522, 834,
996, 998, 1013
Bressler, R., 224, 234, 781, 887, 996
Breuer, H., 708, 1038
Breusch, F. L., 60, 175, 177, 996
Brewer, C. R., 78, 996
Brewer, J. H., 978, 979, 980, 1027
Brian, P. W., 618, 632, 1034
Briault, P. G., 983, 996
Bridger, W. A., 466, 481, 831, 996, 1000
Brierley, G. P., 238, 1063
Briggs, F. N., 179, 996
Brignone, J. A., 38, 41, 713, 775, 779,
783, 1058
Brindley, C; O., 538, 851, 1029, 1030
Brink, N. G., 410, 500, 996
Brixovd, E., 954, 957, 1005
Brock, N., 965, 996
Brocket!, F. P., 813, 1040
Brocklehurst, W. E., 374, 989
Brockman, R. W., 480, 996
Brode, E., 585, 1022
Brodersen, R., 617, 632, 996
Brodie, A. F., 120, 552, 846, 848, 989,
996, 1024
Brodie, B. B., 851, 1012
Brodman, K., 946, 947, 948, 1050
Brody, K. R., 267, 1056
Brody, J. M., 348, 996
Bronk, D. W., 211, 1030
Brooke, J. L., 393, 394, 999
Brooks, J., 735, 875, 1023
Brooks, J. L., 392, 403, 996
Brooks, P., 739, 740, 996
Brooks, S. A., 388, 389, 996
Broquist, H. P., 583, 996
Brosteaux,' J., 437, 1013
Brown, D. D., 843, 996
Brown, D. E. S., 211, 1053
Browm, D. H., 411, 561, 1012 1022,
Brown, D. M., 696, 770, 782, 844, 999,
1055
Brown, E. G., 162, 996
Brown, G. W., Jr., 141, 998
Brown, I. A., 954, 996
Brown, J., 401, 996
Brown, P. K., 953, 1064
Brown, R. R., 1003
Brown, W. D., 137, 996
Bruce, M., 898, 1049
Bruchmann, E. E., 692, 694, 996
Bruemmer, J. H., 17, 26, 996
Brim, C., 918, 923, 996
Brunfaut, M., 969, 1057
Brunnemann, A., 499, 996
Brunngraber, E. G., 473, 996
Bruns, F. H., 406, 855, 996, 1042
Brust, M., 198, 996
Bryce, G. F., 787, 996
Bublitz, C. 842, 996
Buch, M. L., 15, 996
Buchanan, J. M., 333, 1009, 1031
Buchman, E. R., 516, 996
Buckland, F. E., 977, 978, 979, 988
Buddecke, E., 852, 875, 881, 884, 1065
Budillon, G., 616, 1049
Budzilovich, T. V., 587, 995
Biicher, T., 61, 1027
Bueding, E., 376, 381, 382, 383, 842, 997
Bulen, W. A., 614, 708, 840, 997
Bullough, W. S., 199, 200, 225, 990, 997
Bu'Lock, J. D., 231, 997
Bunzel, H. H., 981, 1068
Burch, G., 928, 997
Burch, H. B., 540, 997
Burchenal, J. H., 582, 583, 996, 1009
Burdette, W. J., 77, 79, 82, 91, 179, 997
Burgess, E. A., 81, 176, 183, 997
Burk. D., 292, 388, 392, 400, 401, 1029,
1030, 1037
Burkard, W. P., 318, 1045
Burnett, A. E., 923, 1065
Burnett, G. H., 592, 997
Burney, T. E., 728, 979, 980, 981, 997
Burns, S., 675, 1004
Burr, M. J., 887, 997
Burris, R. H., 53, 61, 171, 291, 292, 293,
294, 997, 999, 1005, 1019, 1032, 1038,
1039, 1067
Burton, K., 268, 341, 348, 545, 547, 549,
997
Busch, H., 15, 91, 98, 100, 101, 102, 103,
104, 152, 156, 201, 217, 218, 228, 232,
437, 997, 1042, 1046
Butler, L. G., 672, 1031
AUTHOR INDEX
1077
Butt, E. M., 960, 1014
Butt, W. D., 451, 997
Buzard, J. A., 709, 718, 997
Bydgeman, S., 401, 1019
Byers, S. O., 288, 997
Byrne, W. L., 270, 390, 412, 853, 1017,
1041
Cadenas, E., 125, 414, 1039, 1056
Cady, P., 146, 997
Caffrey, R. W., 543, 553, 851, 864, 997,
1020
Cafruny, E. J., 689, 918, 920, 921, 922,
933, 997, 1063
Cailleau, R., 62, 63, 1033
Cain, R. B., 90, 997, 998
Calcutt, G., 982, 997
Caldwell, E. F., 569, 997
Caldwell, M. L., 658, 660, 662, 673, 674,
683, 684, 833, 1065
Calesnick, B., 960, 997
Caley, E. R., 948, 1016
Calkins, E., 304, 1031
Callaghan, O. H., 446, 772, 847, 997
Callahan, S., 283, 1006
Calo, N., 892, 1012
Caltrider, P. G., 133, 997
Calvin, M., 145, 163, 409, 619, 637, 646,
656, 658, 991, 997, 1002, 1036, 1043
Cama, H. R., 547, 549, 660, 685, 832,
1029, 1047
Cameron, G., 201, 998
CampbeU, A. D., 670, 690, 1006
Campbell, D. E. S., 923, 924, 998
Campbell, L. L., Jr., 269, 833, 834, 1035
Campbell, P. N., 272, 405, 998
Canellakis, Z. N., 305, 306, 998
Caniggia, A., 215, 1031
Cannata, J. J. B., 711, 852, 998
Cantero, A., 381, 1046
Cantino, E. C, 169, 1033
Capobianco, G., 616, 1049
Carafoli, E., 121, 122, 865, 989
Caravaca, J., 439, 1014
Caraway, W. T., 703, 998
Carbon, J. A., 481, 998
Carney, S. A., 391, 998
Carpenter, A. T., 600, 998
Carpenter, T. M., 951, 998
Carpenter, W. D., 61, 998
Carr, A. J., 428, 998
Carsiotis, M., 844, 852, 1054
Carson, S. F., 15, 1044
Carter, C. E., 282, 359, 1002, 1029
Carter, J. R., 650, 1047
Cartwrigt, N. J., 90, 998
Casal, A., 657, 1003
Cascarano, J., 879, 895, 998
Castelfranco, P., 136, 998
Catterson, D. A., 959, 987
Caughey, W. S., 61, 329, 591, 998, 1061
Cavallito, C. J., 621, 973, 983, 998, 1027
Cecil, R., 756, 988, 998
Cedrangolo, F., 668, 998
Cerecedo, L. R., 516, 518, 519, 522, 523,
530, 531, 533, 998, 1005, 1006, 1056
Cereijo-Santalo, R., 396, 397, 1065
Cerfontain, H., 745, 998
Cha, C.-Y., 683, 998
Chaberek, S., 13, 739, 998
Chadwick, J. B., 169, 880, 1041
Chaffee, E., 875, 1021
Chaffee, R. R., 437, 998
Chaikoff, I. L., 141, 145, 146, 147, 179,
209, 226, 234, 273, 987, 996, 997, 998,
1025, 1060
Challenger, F., 2, 225, 226, 228, 998, 1064
Chamberlain, G. T., 966, 994
Chambers, R., 201, 998
Chan, P. C, 792, 866, 867, 1058
Chance, B., 17, 786, 998
Chang, V M., 555, 1019
Chanutin, A., 68, 1039
Chaplain, R. A., 874, 1014
Chapman, D. W., 944, 957, 998
ChappeU, J. B., 86, 121, 210, 816, 998
Chari-Bitron, A., 31, 104, 239, 828, 998
Charmandairjan, M. O., 835, 999
Chattaway, F. W., 168, 169, 190, 999
Chatter jee, G. C, 529, 530, 576, 999, 1002
Chaudhuri, D. K., 835, 999
Chaure, V. J., 187, 208, 218, 883, 914,
915, 916, 1002
Cheesman, D. F., 939, 999
Cheftel, R. L., 15, 999
1078
AUTHOR INDEX
Cheldelin, V. H., 38, 52, 132, 138, 143,
705, 712, 837, 853, 999, 1002, 1023,
1026, 1033
Chemelin, I., 399, 400, 1056
Chen, K. K., 999
Chen, R. F., 509, 513, 696, 782, 844, 999
Cheniae, G., 33, 65, 850, 855, 999
Chenoweth, M. B., 212, 625, 992, 1065
Chernick, S., 179, 996
Chiba, H., 675, 685, 692, 771, 772, 835,
999
Chiba, Y., 194, 999
Chick, H., 971, 972, 999
Chiga, M., 872, 874, 999
Chin, H. P., 403, 999
Chinard, F. P., 640, 643, 657, 662, 665,
669, 697, 701, 702, 703, 727, 803, 999,
1017
Chirigos, M. A., 266, 999
Choppin, P. W., 976, 977, 979, 980, 981,
999
Chow, C. T., 173, 182, 1054
Christensen, E., 392, 393, 394, 403, 435,
996, 999
Christensen, H. N., 15, 155, 266, 575, 908,
999, 1043, 1048
Christian, W., 768, 1064
Christie, G. S., 81, 999
Christman, A. A., 236, 999
Christophers, S. R., 559, 1050
Chung, A. E., 781, 839, 999
Cianci, V., 971, 972, 999
Ciferri, O., 404, 411, 542, 839, 994, 999
CifoneUi, J. A., 385, 1004
Ciotti, M. M., 487, 496, 497, 498, 504,
508, 784, 1024, 1062, 1069
Citri, N., 249, 599, 615, 688, 999, 1011
Clagett, C. O., 61, 999
Clark, D. S., 74, 77, 190, 228, 999
Clark, I., 562, 572, 1045
Clark, W. C, 520, 1067
Clark, W. G., 310, 314, 1016, 1045
Clark, W. M., 656, 999
Clarke, D. D., 887, 1000
Clarkson, T. W., 916, 920, 1000
Claus, D., 880, 971, 972, 984, 1000
Claus, G. W., 852, 991
Clayton, R. A., 839, 885, 1000
Clayton, R. K., 53, 77, 78, 81, 146, 1000
Cleland, K. W., 22, 29, 174, 175, 817, 855,
874, 875, 882, 884, 1000
Cleland, R., 967, 1000
Cleland, W. W., 640, 1000
Clementi, A., 228, 1000
Clements, A. N., 54, 77, 1000
Cliffe, E. E., 544, 1000
Clowes, G. H. A., 198, 999, 1028
Coates, J. H., 287, 989
Cochin, J., 590, 597, 989
Cochran, D. G., 332, 446, 1038, 1050
Cochrane, V. M., 79, 97, 1000
Coe, E. L., 396, 397, 1021
Coen, L. J., 575, 1022
Coggeshall, R. E., 499, 1000
Cohen, E. M., 873, 927, 1000, 1003
Cohen, G. N., 321, 323, 356, 1057, 1061
Cohen, L. H., 466, 478, 481, 831, 996,
1000
Cohen, P. P., 116, 157, 305, 306, 336,
592, 781, 804, 840, 997, 998, 1000,
1003, 1032, 1036
Cohen, R. A., 55, 177, 1000
Cohen, S., 226, 1000
Cohen, S. S., 476, 479, 1008, 1036
Cohn, M., 400, 415, 416, 780, 1000, 1066
Coleman, G. H., 974, 1000
Coleman, J. E., 770, 818, 1000, 1062
Coleman, R., 662, 1000
Collander, R., 189, 1000
Collias, E. C, 29, 1000
Collier, H. B.. 553, 774, 825, 833, 836,
845, 851, 854, 859, 901, 906, 993, 1056,
1061
Collins, A., 591, 1061
Colowick, S. P., 273, 405, 433, 434, 435,
485, 487, 490, 493, 498, 504, 507, 508,
509, 511, 512, 783, 896, 912, 1000,
1001, 1006, 1024, 1027, 1041, 1043,
1046, 1069
Colpa-Boonstra, J., 32, 35, 1020
Colter, J. S., 791, 794, 815, 817, 820,
1000, 1006
Combs, A. M., 540, 997
Commoner, B., 18, 22, 196, 1000, 1019
Conches, L., 74, 77, 81, 92, 116, 117,
169, 1058
AUTHOR INDEX
1079
Conchie, J., 428, 429, 812, 814, 838, 1000
Conn, E. E., 27, 74, 80, 82, 119, 120, 355,
852, 1000, 1028, 1057
Connamacher, R. H., 316, 317, 318, 139,
320, 1018
Connelly, C. C, 813, 1040
Connor, R., 619, 987
Connors, P., 208, 1063
Conover, T. E., 121, 1000
Conrad, 619, 1000
Contopoulou, C. R., 136, 675, 998, 1004
Contrera, J. F., 320, 599, 1013
Converse, J. L., 971, 973, 1000
Conway, T. W., 354, 1001
Cocul, B. J., 190, 196, 1001
Cook, A. M., 971, 972, 1001
Cook, E. S., 744, 810, 835, 856, 870, 871,
972, 973, 1001, 1028, 1052, 1055, 1056,
1060
Cook, R. P., 3, 78, 228, 1001
Cook, S. F., 880, 1001
Coombs, T. L., 781, 827, 1001
Coon, M. J., 782, 834, 1029, 1048
Cooper, C, 444, 445, 705, 869, 872, 1001,
1031
Cooper, E. A., 195, 972, 1001
Cooper, J., 855, 1059
Cooper, J. A. D., 550, 1001
Cooper, J. R., 354, 1062
Cooperstein, S. J., 175, 663, 1001, 1010
Coore, H. G., 376, 1001
Copenhewer, J. H., Jr., 119, 1001
Coper, H., 499, 996
Cori, C. F., 376, 387, 388, 390, 405, 561,
648, 789, 803, 808, 811, 813, 1001,
1022, 1026, 1034, 1055
Cori, G. J., 376, 405, 1001, 1055
Cori, 0., 60, 119, 121, 707, 717, 873,
1049
Corley, R. C, 2, 219, 273, 610, 1001,
1053, 1056
Cormier, M. J., 845, 891, 1001
Corner, E. D. S., 967, 994
Corte, G., 586, 1023
Corwin, A. H., 213, 747, 1065
Corwin, L. M., 63, 878, 1001
Cotter, G. J., 696, 1007
Cottrell, T. L., 744, 1001
Cousins, F. B., 543, 1001
CoveUi, I., 546, 972, 1007
Covin, J. M., 128, 216, 1001
CowgiU, R. W., 405, 852, 853, 1001
Coxon, R. v., 76, 1001
Coyne, B. A., 575, 999
Cramer, F. B., 386, 389, 392, 1001
Cramer, H. I., 619, 987
Crandall, D. I., 541, 545, 709, 771, 772,
787, 1001, 1008
Crane, F. L., 16, 17, 26, 996, 1014
Crane, R. K., 262, 264, 379, 383, 388,
390, 782, 824, 843, 1001, 1008, 1028,
1056
Cravens, W. W., 530, 576, 1001, 1040
Crawford, E. J., 286, 287, 288, 289, 407,
474, 1024, 1033
Crawford, M. A., 109, 1001
CrawhaD, J. C, 165, 1001
Creaser, E. H., 122, 177, 200, 395, 1001
Creasey, N. H., 272, 405, 998
Creeth, J. M., 682, 1067
Cremer, J. E., 75, 76, 127, 153, 1001,
1002
Cremona, T., 65, 435, 436, 437, 510, 798,
872, 994, 1002, 1038
Creveling, C. R., 316, 320, 611, 1002,
1062
Creveling, R. K., 466, 1037
Crewther, W. G., 593, 660, 693, 708, 1002
Crocker, B. F., 418, 1053
Cronin, M. A., 358, 1002
Cross, A. C, 916, 1000
Cross, R. J., 204, 928, 1002
Crout, J. R., 592, 611, 1002
Cruickshank, C. N. D., 225, 990
Cruickshank, D. H., 64, 1002
Csonka, E., 957, 959, 961, 1038
Cumings, J. N., 952, 1002
Cummins, J. T., 154, 712, 1002, 1050
Cunningham, L. W., 681, 745, 750, 1002
Cunningham, L. W., Jr., 513, 1062
Cunningham, M., 137, 138, 141, 142,
1011
Cuong, T. C, 225, 226, 1009
Curl, A. L., 1023
Curtis, W. C, 834, 1061
Cutolo, E., 552, 708, 994
1080
AUTHOR INDEX
Czarnetzky, E. J., 956, 972, 1055
Czekalowski, J. W., 194, 1002
D
D'Abramo, F., 164, 1049
Da Costa, F. M., 315, 1012
Dahl, J. L., 478, 1002
Dahlqvist, A., 416, 1002
Dajani, R. M., 91, 1002
Dale, R. A., 920, 921, 1002
Dalgarno, L., 74, 75, 79, 82, 171, 1002
Daly, J. W., 316, 611, 1002 1062
Dalziel, K., 513, 1002
Damaschke, K., 891, 1002
Damjanovich, S., 949, 1016
Dancis, J., 585, 990
Danforth, W., 28, 51, 53, 56, 77, 228, 1002
Daniel, L. J., 530, 585, 1002, 1058
Danielski, J., 91, 1002
Danielson, L., 708, 849, 1006
Dann, O. T., 359, 1002
Darlington, W. A., 207, 1002
Darragh, J. H., 925, 1065
Das, H. K., 153, 1002
Das, N. B., 32, 38, 48, 62, 177, 178, 437,
1002
Das, S. K., 529, 1002
Das, S. N., 9, 1002
Datta, A., 548, 556, 705, 865, 869, 873,
1014, 1046
Datta, A. G., 519, 1002
Daus, L., 145, 1002
d'Aurac, J., 225, 226, 1009
Da Vanzo, J. P., 358, 1002, 1052
Davenport, H. W., 187, 208, 218, 883,
914, 915, 916, 1002
Davenport, V. D., 883, 914, 915, 1002
Davidson, E., 356, 1012
Davidson, H. M., 63, 442, 1041
Davidson, J. D., 281, 282, 466, 477, 1007
Davidson, N., 739, 740, 741, 996, 1008,
1069
Davies, D. D., 349, 593, 604, 607, 857,
990, 1006
Davies, G. E., 224, 236, 1003
Davies, J. H., 428, 995
Davies, R. E., 88, 991
Davies, R. I., 264, 911, 1031
Davis, F. F., 712, 1003
Davis, J., 348, 1024
Davis, N. C, 783, 1055
Davison, D. C., 170, 1003
Dawes, E. A., 124, 432, 593, 600, 853,
991, 1003
Dawid, I. B., 333, 1009
Dawson, C. R., 697, 1060
Dawson, R. M. C, 75, 151, 858, 1003,
1060
Day, B., 978, 979, 980, 1027
Day, H. G., 385, 520, 525, 530 991, 1010
Day, M. D., 318, 1003
Day, R. A., 333, 1009
Dearborn, E. H., 332, 550, 551, 1013, 1062
Debay, C. R., 834, 1006
de Bernardinis, C, 217, 218, 1015
De Bodo, R. C, 401, 988
De Boer, C. J., 461, 462, 1018
de Caro, L., 520, 522, 526, 527, 1003, 1048
Dechary, J. M., 593, 1028
Decker, R. H., 610, 782, 843, 1003
De Eds, F., 601, 995
de Favelukes, S. L. S., 74, 77, 81, 92,
116, 117, 169, 1058
Deferrari, J. O., 687, 712, 716, 775, 783,
810, 853, 1058
De Flora, A., 407, 1013
De Gowin, E. L., 902, 905, 1053
De Graff, A. C., 944, 1003
De Groot, C. A., 927, 1003
De Groot, N., 840, 1003
de Issaly, I. S., 74, 79, 81, 1003
Deitrich, R. A., 662, 1003
Deitz, V. R., 643, 662, 669, 803, 1017
Dejung, K., 925, 1003
de Kock, L. L., 133, 1040
de Kock, P. C, 133, 1040
de la Fuente, G., 389, 1056
Delaunay, A., 203, 1030
del Campillo, A., 708, 837, 1057
Dellert, E. E., 458, 1003
Delmonte, L., 581, 1003
De Luca, H. A., 150, 151, 1027
Demis, D. J., 876, 883, 893, 894, 895,
897, 898, 1003
De Moss, J. A., 89, 1003
AUTHOR INDEX
1081
De Moss, R. T>., 831, 1003
Dempsey, W. B., 569, 1009
Dengler, H., 314, 1003
Denison, F. W., Jr.. 15, 1003, 1044
Dennis, D., 434, 1003
Denstedt, O. F., 55, 65, 179, 436, 485,
487, 489, 492, 503, 988, 1035, 1043,
1049
de Petrocellis, B., 469, 675, 706, 1051
Derbyshire, J. E., 203, 223, 993
De Robertis, E., 203, 727, 1003
Dervartanian, D. V., 18, 38, 1003
Desnuelle, P., 649, 657, 664, 666, 668,
1003, 1028
De Stevens, G., 461, 462, 1018
Dettmer, F. H., 53, 77, 78, 81, 146, 1000
De Turk, W. E., 228, 888, 993, 1003
Devlin, J. M., 872, 1031
Dewan, J. G., 61, 594, 1014
Dewey, D. L., 593, 708, 1003
Deykin, D., 835, 1003
Dhar, S. C, 682, 687, 693, 1003
Diamant, E. J., 495, 1015
Di Carlo, F. J,, 659, 660, 662, 674, 683,
684, 792, 833, 1003
Dick, G. F., 666, 667, 991
Dickens, F., 36, 41, 503, 659, 662, 695,
711, 855, 1003
Dickerman, H. W., 490, 850, 1003
Dickman, S., 852, 991
Dickman, S. R., 273, 462, 592, 1003, 1056
Diczfaliisy, E., 461, 464, 992, 1004
Dieterle, W., 15, 1004
Dietrich, E. V., 40, 1004
Dietrich, L. S., 36, 37, 41, 240, 288, 505,
538, 569, 570, 1004, 1010, 1053
Dikstein, S., 387, 436, 437, 991, 1004
Dilley, R. A., 557, 1004
Dils, R., 147, 1004
Dilworth, M. J., 293, 1044
d'Incan, E., 236, 1049
Dintzis, H. M., 757, 1021
D'lorio, A., 612, 725, 947, 1004
Dippy, J. F. J., 299, 1004
di Frisco, G., 594, 1063
Di Sabato, G., 808, 1004
Dische, Z.. 414, 465, 1004
di Stefano, H. S., 689, 921, 922, 997
Dittrich, S., 15, 1004
Dixon, D. H., 594, 694, 1004, 1051
Dixon, K. C, 384, 1004
Dixon, M., 280, 693, 694, 993, 1004
Dizon, F. Y., 574, 577, 1051
do Amaral, D. F., 511, 710, 849, 1035
Dobson, M. M., 447, 1059
Doctor, V. M., 577, 582, 1004
Dodgson, K. S., 444, 684, 1004
Doherty, D. G., 373, 1063
Doherty, M. E., 776, 777, 788, 836, 1013
Dohi, S. R., 461, 1046
Doi, R. H.. 552, 553, 710, 848, 1004
Doi, Y., 969, 1025
Doisy, E. A., 839, 1004
Doisy, R. J., 283. 859, 1004
Dohn, M. I., 430, 547, 553, 676, 693, 847,
849, 1004
Dominguez, A. M., 1004
Domonkos, J., 179, 1004
Donovan, J. W., 638, 1004
Doran, D. J.. 173, 1004
Dorfman, A., 385, 1004
Dorman, P. J., 928, 1012
Doudoroff, M., 675, 1004
Dove, W. F., 741, 1004
Dowdle, E. B., 208, 909, 914, 1004, 1051
Downey, M., 198, 990
Downing, S. J., 264, 1049
Downs, C. E., 335, 1021
Doyle, M. L., 839, 1004
Drabikowski, W., 939, 940, 1004
Dragsdorf, R. D., 975, 1016
Dreser, H., 941, 1005
Dresse, A., 611, 989
Dreyfus, P. M., 519, 1005
Driver, G. W., 224, 236, 1003
Druckrey, H., 965, 996
Drury, D. R., 387. 390, 399, 404, 1066
Dube, S. K., 683, 783, 845, 1005
Du Bois, K. P., 22, 30, 41, 659, 676, 687,
784, 856, 1045
du Bay, H. G., 128, 177, 1005
Duchateua-Bosson, G., 297, 1008
Dudley, C., 227, 1053
Duerkson, J. D., 782, 1005
Dufait, R., 693, 1036
Duggan, D. E., 281, 282, 285, 1005
1082
AUTHOR INDEX
Dull, M. F., 620, 1057
Dulskas, A., 926, 1050
Dumont, J. E., 76, 131, 1005
Dunn, A., 401, 988
Duperon, R., 15, 1005
Durham, N. N., 267, 613, 1005, 1020
Duthie, R., 363, 994
Duysens, L. N. M.. 891, 1043
Dyer, H. M., 260, 1005
Dzurik, R., 926, 954, 957, 1005
Eadie, G. S., 459, 1025
Eagon, R. G., 389, 1066
Earl, J. M., 751, 774. 838, 1012. 1062
Eaton, M. D., 193, 1005
Ebata, M., 842, 1051
Eber, J., 902, 903, 904, 905, 906, 907, 1065
Eberson, L., 9, 1005
Ebert, H., 228. 993
Eberts, F. S., 171, 1005
Edelhoch, H., 758, 761, 1005
Edlbacher, S., 269, 335, 353, 1005
Edman, K. A. P., 938, 1005
Edsall, J. T., 638, 664. 665. 667, 758,
759, 761, 1005, 1014
Edson. N. L., 55, 56. 138, 140, 144, 177,
237, 238, 1005
Edwards, B. B., 948. 1016
Edwards, J. G., 924. 1005
Eeg-Larsen, N., 384, 1005
Egami, F., 537. 541, 544, 817, 1005,
1041, 1060
Eggerer, H., 836, 887, 987, 1005
Eggleston. L. V., 22, 27, 74, 75. 77, 94,
96, 115, 116, 124. 153, 158, 830, 1028,
1060
Ehlers, K. H., 221, 223, 993
Ehrmantraut, H., 163, 1005
Eich, S., 518, 519, 522, 523, 531, 998, 1005
Eichel, B., 60, 61, 63, 169, 196, 228, 550,
987, 1005
Eichel, H. I.. 20, 543. 547, 552, 555, 710,
848, 1005
Eisen, H. N., 263, 387, 392, 394, 1018
Eisenstadt, J. M., 269, 1006
Eisenstark, A., 975, 1016
Eldjarn, L., 639, 663, 1006
El Hawary, M. F. S., 110, 111, 1006
Elion, G. B., 283, 1006
Elkins-Kaufman, E., 365. 1006
Ellem. K. A. O., 791. 794, 815, 817, 820,
1000, 1006
Ellfolk. N.. 355, 685, 707, 1006
Ellias, L., 423, 772, 1016
Elhott. H. W., 179, 181. 1006
ElUott. K. A. C, 75, 91, 178, 185, 212,
994, 1006
Elliott, W. H., 471, 1006
Ellis, R. J., 857, 1006
Ellis, S., 217, 1006
Ellman, G. L., 640, 1006
Elodi, P.. 650, 788, 804, 809, 826, 1006,
1023
El'tsina, N. V., 395. 1006
Eltz, R. W., 886. 1006
Elvehjem, C. A., 29, 35, 36, 37, 41. 175,
240, 260, 288, 489, 503, 504, 593, 659,
662, 988, 1004, 1007, 1045, 1059, 1068
Ely, J. O., 400, 1006
Emerson, G. A., 518, 531, 538, 1006
Emerson, P. M., 436, 1006
Emery, J. F., 834, 1006
Emmelot, P., 130, 149, 150, 156, 1063
Endahl, B. R., 848, 1006
Engel, L. L.. 555, 713, 781, 1030, 1050
Engel, S. L., 632, 1057
Engelhardt, W., 865, 1006
England, S., 273, 675, 703, 706. 716, 718,
804, 833, 1006, 1054
Engle, C. G., 978, 1015
Ennor, A. H., 467, 685, 704, 707, 710,
833, 836, 845, 1006, 1010, 1014, 1039
Eny, D. M., 228, 1006
Eppley, R. W., 881, 908, 909, 912, 1006
Erbland, J., 151, 1036
Erdos, F. G., 834, 1006
Erf. L. A., 217, 218, 1015
Erlenmeyer, H., 518, 530, 1006
Ernster, L., 17, 18, 33, 444, 445, 708,
849. 865, 872, 989, 1006. 1030. 1033,
1054
Errera, M., 969. 1057
Eskarous, J. K., 973, 978, 1060
Estabrook, R. W., 395, 1035
AUTHOR INDEX
1083
Estes, E. H., Jr., 399, 1030
Estler, C. J., 750, 881, 884, 885, 1000
Estrada, J., 711, 839, 1061
Eubank, L. L., 670, 690, 1006
Eusebi, A. J., 518, 530, 533, 998, 1000
Evang, A., 662, 663. 1046
Evans, E. A., Jr., 53, 63, 73, 74, 78. 91,
93, 124, 226, 1007, 1056. 1063
Evans, H. J.. 33, 65, 850, 855, 999
Evans, J. B.. 78, 83, 1044
Evans, J. I., 11, 13, 1007
Evert, H. E., 773, 1007
Ewers, A., 865. 912. 1030
Eyer, H., 694. 1009
Eyring, H., 5. 1007
Eysenbach, H., 49, 1008
Ezaki, S., 271, 417, 1007
Fahey, J. L.. 538. 1030
Fahrlander, H., 157, 1007
Fain, J. N., 391, 1007
Fairclough, R. A., 3, 1007
Fairhurst, A. S., 1035
Falcone, A. B., 846, 851, 1053
Falcone, G., 546, 972, 1007
Falk, J. E., 972. 973, 988
Fanestil, D. D., 869, 1007
Fanshier, D. W., 334, 1029
'Fantes, K. H., 422, 1019
Farah, A., 204, 205. 214. 626, 689, 918.
921, 922, 943. 944, 945, 957, 997, 1007,
1011, 1032
Farah, A. E., 626, 917, 934, 935, 1007,
1038
Farkas. G. L.. 169. 170, 195, 1007
Farkas, W., 451, 1034
Farnham, A. E., 435, 1043
Farrar, W. V., 576, 1015
Fasella, P., 788, 803, 827, 1007, 1061
Fasold, H., 641, 1007
Fastier. F. X.. 363, 364. 365, 994, 1007
Favarger, P., 157, 1007
Favelukes, G., 708, 717. 778, 781, 838,
1007
Fawaz, E. X., 41, 55, 56, 64, 95, 112, 128,
129, 221, 236, 925, 927, 1007
Fawaz, G., 41, 55, 56, 64, 95, 112, 128,
129, 221, 236. 925, 927, 1007
Fazekas. J. F., 500, 990
Featherstone, R. M., 40, 596, 1004, 1028
Feeney, R. E., 745, 754, 760, 1034
Feher, 0.. 949, 1016
Feigelson, P., 281, 282, 466, 477, 503,
603, 1007, 1014
Feigl, F.. 15, 1007
Feinberg, R. H.. 660, 693, 713, 1007
Feinstein, R. X., 696, 1007
Feist, E., 146, 224, 234, 1007
Feist, F., 617, 618, 1007
Fekete, J., 460, 1015
Feldberg, W., 949, 1007
Feldman, W.. 848, 855, 1007
Felenbok, B.. 842. 1007
Feller, D. D., 146, 224, 234, 1007
Fellig, J., 461, 462, 712, 1008, 1070
Fellman. J. H.. 314. 1008
Felsher. R. Z., 687. 1013
Felton, S. P., 539, 543, 544, 1020
Felts, J. M., 613, 1008
Feng, J. Y., 853, 1016
Fenn, P., 965, 1048
Ferguson, J. H., 456, 1012
Fenley, H. X.. 772, 791, 793, 795, 796,
797, 1008
Ferno, 0., 461, 464, 1004, 1008
Ferrari, G., 520, 526, 578, 1003, 1048
Ferrari. R. A., 379, 390, 449, 1008
Ferreira, R., 739, 1008
Festenstein, G. X., 417, 429. 1008
Fewson, C. A., 547, 554, 1008
Fewster, J. A., 842, 1008
Fex, H., 461, 464, 1004, 1008
Field, J. B., 268, 1056
Fildes, P.. 971, 975, 1008
Filho, J. B. M., 510, 552, 850, 1047
Finamore, F. J., 887, 997
Findlay, J., 429, 1008
Fine, A.. 19, 29, 1044
Finean. J. B., 950, 1038
Fink. J., 210, 1021
Finkelman, F., 704, 723, 939, 990
Finkle, B. J., 804, 1008
Fischer, A.. 465, 1008
Fischer, E. H., 453, 887, 995, 1028
1084
AUTHOR INDEX
Fischer, F. G., 49, 1008
Fischer, J., 810, 1064
Fischer, P., 697, 698, 699, 700, 1012
Fishbein, W. N., 433, 1008
Fisher, A. L., 40, 1004
Fisher, F. M., Jr., 839, 1008
Fisher, H. F., 293, 1008
Fishgold, J. T., 30, 50, 200, 1008
Fishman, R. A.. 401, 1008
Fishman, W. H., 63, 428, 442, 1008,
1041, 1054
Fiskin, R. D., 913, 1050
Fitzgerald, R. J., 632. 1008
Fitzpatrick, J. B., 304, 529, 1031, 1062
Fiume, L., 238, 1008
Flaks, J. G., 476, 1008
Flamm, W. G., 771, 772, 1008
Flatow, L., 670, 1008
Flavin, M., 64, 145, 224, 226, 234, 235,
357, 1008
Fleischer, S., 16, 1014
Flesch, P., 767, 1008, 1037
Flickinger, R. A., Jr., 203, 1008
Flood. A. E., 195, 1026
Florkin, M., 297, 1008
Fluharty, A. L., 408, 1008
Fluri, R., 529, 1008
Foa, P. P., 394, 1038
Fodor, P. J., 595, 1008
Foldeak, S., 503, 1037
Folk, J. E., 367, 1008
Folkers, K., 538, 1019
Foltz, V. D., 974, 1033
Fonnesu, A., 210, 1050
Fonnum, F., 857, 1008
Ford, L., 873, 883, 1021
Forney, R. B., 960, 1008
Forsen, S., 619, 1009
Forssman, S., 18, 48. 98, 104, 105, 213,
215, 217, 218, 959, 1009
Forster, R. P., 205, 921, 1009
Forsyth, F. R., 196, 529, 1050
Forti, G., 623, 1009, 1036
Foss, O., 639, 1009
Foster, C., 530, 1023
Foster, R. J., 349, 372, 373, 374, 1009
Foulerton, A. G. R., 954, 1009
Foulkes, E. C, 912, 1009
Fountain, J. R., 582. 1009
Fourneau, E., 952, 1009
Fournier, P., 225, 226, 1009
Fonts, J. R., 429, 1013
Fraenkel-Conrat, H., 681, 741, 980, 1009,
1054
Francis, A. M., 226, 1063
Francis, M. J. O., 845, 1009
Frank, E., 461, 1015
Frank, S., 695, 1020
Franke, W., 18, 32, 37, 40, 44, 664, 665,
1009, 1028
Frankel, S., 569, 1048
Franshier, D. W., 596, 1028
Eraser, D., 877, 981, 1054
Eraser, D. M., 551, 1009
Frear, D. S., 597, 846, 1023
Fredrickson, D. S., 614, 1057
Freeland, M. R., 696, 1060
Freedland, R. A., 325, 1009
Freedlander, B. L., 202, 218, 1009
Freeman, M., 581, 1043
French, D., 421, 1038
French, F. A., 202, 218, 1009
French, R. C, 591, 991
French, T. V., 333, 1009
Freudenberg, K., 694, 1009
Friberg, L., 959, 960, 1009
Fridhandler, L., 179, 183, 391, 393, 574,
1009
Fridovich, I., 288, 451, 549, 555, 803,
807, 1009, 1034, 1046
Fried, G. H., 58, 1009
Friedemann, T. E., 349, 1009
Frieden, C., 508, 514, 1009, 1010
Frieden, E., 325, 744, 778, 1010
Friedenwald, J. S., 444, 465, 991, 1034
Friedland, I. M., 505. 1004, 1010
Friedman, B., 137, 141, 1065
Friedman, D. L., 228, 233, 235, 1037,
1057
Friedmann, E., 34, 35, 1039
Friedmann, H. C, 470, 1010
Frimmer, M., 548, 552, 553, 676, 793, 850,
1010
Frisell, W. R., 60, 341, 346, 601, 704,
706, 717, 772, 780, 1010
Fritz, C. T., 175, 1010
AUTHOR IXDEX
1085
Frohman. C. E., 89, 520, 525, 530. 1010
Fromageot, C, 60, 357. 1010
Fromm. H. J.. 357. 376, 1010
Frommel, E.. 835, 1010
Fruton, J. S.. 375, 1010
Fu, T.-H., 792, 813, 1043
Fuentes, V., 877, 1010
Fujie. Y., 865, 1010
Fujimoto, D., 554. 684, 1010, 1022
Fukui, T., 887, 1016
Fukunaga, K., 843, 1029
Fuld, M., 71, 140, 995, 1010
Fulton, J. D., 168, 179, 559, 693, 882, 1010
Furgiuele, F. P., 978, 981, 1052
Furst, A., 202, 218. 1009
Furuya, K., 820, 867. WHO
Futterman, S., 581, 582, 1010
Gabrio, B. W.. 543, 553. 581. 582. 711,
848. 851, 864. 9V3, 997, 1020. 1054
Gaebler, O. H., 494, 504, 992, 1010
Gaffney, P. E., 978, 1015
Gaffney, T. J., 710, 845, 1010
Gage, J. C, 961, 1010
Gajdos, A., 62, 1010
Gale. E. F., 660, 845. 1060
Gale, G. R., 430, 1010
Galoyan, A. A., 951, 956, 985, 1010
Galoyan, S. A., 951, 958, 959, 1010
Galston, A. W., 750, 1010
Gamble. J. L., Jr., 383, 722, 872, 909, 914,
1010, 1031, 1052
Gamble, W., 91, 1002
Gamborg, O. L., 599, 1011
Gammon, G. D., 573, 1011
Gane, R., 5., 1011
Ganguly, J., 547, 554, 1035
Garber, N., 249, 599, 615, 688, 999, 1011
Garcia -Hernandez, M., 355. 1011
Gardier, R. W., 921, 1011
Gardner, E. A., 214, 1011
Garen, A., 439, 1011
Garfinkel. D., 849, 1011
Garlid, K., 924, 1065
Gause, G. F., 981, 1011
Gawehn, K., 695, 1064
Gayer. J., 922. 1011
Gehrig, R. F., 353. 1042
Geiger, P. J.. 789, 819, 1048
Geissler, A.-W., 695, 1064
Gelfant, S., 200, 968, 1011
Gellert, M.. 803, 807, 814, 1012
Gelles, E.. 4, 1011
Gellhorn, A., 546, 576, 577, 1011.
1053
Gemma, F. E., 594, 1034
Gemmill, C. L., 772, 827, 828, 838,
1011
Geppert, J., 971, 1011
Gerard, R. W., 55, 177, 212, 1000,
Gerarde, H. W., 156. 1011
Gerardin, C, 983. 984, 1011
Gergely. J., 475, 866. 868, 939.
1004, 1011
Gerhardt, P., 768, 1011
Gerhart, J. C, 468, 480, 481, 816,
Gerlach, D., 802, 1044
Gershanovich. V. N., 387, 1011
Gershenfeld, L., 690, 1011
Gassier, U., 945, 946, 1011
Gest, A., 49, 294, 1011, 1044
Geuther, 617, 619, 1011
Gey, K. F., 318, 1045
Gey, M, K., 878, 1018
Geyer, R. P., 114, 137, 138, 141,
148, 150, 595, 1011, 1033, 1049
Gezon, H. M., 90, 1069
Ghiretti, F., 54, 92, 105, 114, 174,
228, 336, 709, 718, 840, 991, 1011,
Ghiretti-Magaldi, A., 54, 114, 174,
Ghosh, A. K., 660, 662, 675, 692,
773, 833, 1011
Ghosh, S., 356, 1012
Giartosio, A., 808, 827, 1061
Gibbins, L. N., 833, 1012
Gibbs, M., 838, 892, 1012
Gibbs, R., 602, 1068
Gibson, F.. 321, 1012
Gibson, K. D., 751, 888, 1012
Gibson, Q. H., 757, 1012
Gibson, S., 960, 993
Giebel, O., 187, 188, 1012
Giebisch, G., 923, 928, 936. 1012
Gilbert, D. A., 797, 803, 1012
1047,
876,
1032
940.
1011
142,
227,
1048
1011
772,
1086
AUTHOR INDEX
Gilfillan, R. F., 224, 1012
Gillespie, J. M., 712, 1012
Gillespie, L., 315, 1042
Gillespie, R. E., 710, 1030
Gillette, J. R., 851, 1012
Gilman, A., 698, 699, 700, 1012, 1045
Gilmour, D., 803, 807, 814, 816, 819,
820, 821, 866, 869, 1012
Gil y Gil, C, 985, 1012
Ginoza, Y. W., 737, 973, 1050
Ginsberg, T., 458, 459, lOlH
Ginsburg, S., 532, 1041
Giovanelli, J., 145, 228, 231, 1012
Girerd, R. J., 333, 1012
Giri, K. V., 660, 839, 854, 1012, 1047
Gitlin, J., 548, 551, 1024
Giuditta, A., 33, 38, 49, 773, 850, 855,
1012, 1064, 1065
Gladner, J. A., 367, 369, 370, lOOS, 1041
Glahn, P. E., 268, 340, 104S
Glaid, A. J., 432, 433, 435, 436, 1015, 1042
Glaser, L., 411, 1012
Glasziou, K. T., 834, 1012
Glazko, A. J., 456, 1012
Glenn, J. L., 17, 26, 996
Glick, M. C, 26, 989
Glimm, E., 259, 421, 1067
Glock, G. E., 503, 708, 711, 838, 839,
1003, 1012
Glynn, I. M., 705, 864, 1012
Gnuchev, N. V., 359, 1026
Goddard, A. E., 195, 1001
Godeaux, J., 938, 1012
Godzeski, C, 26, 36, 1012
Goebel, W. F., 657, 1012
Goedde, H. W., 432, 1020
Goedkoop, J. A., 4, 1012
Gonnert, R., 977, 978, 1013
Goerz, R. D., 292, 1012
Goffart, M., 697, 698, 699, 700, 1012
Gold, A. H., 641, 1012
Goldberg, G., 206, 1012
Goldberg, L., 63, 1012
Goldberg, L. I., 315, 1012
Goldemberg, S. H., 476, 1031
Goldin, A., 400, 401, 496, 1024, 1030
Goldinger, J. M., 75, 79, 115, 174, 991,
1057
Goldman, D. S., 60, 268, 596, 831, 832,
843, 854, 991, 1010, 1013
Goldschmidt, E. P., 77, 237, 992, 1013
Goldstein, A., 776, 777, 788, 836, 1013
Goldstein, L., 205, 332, 550, 551, 1009,
1013, 1062
Goldstein, M., 320, 599, 1013
Golstein, M. H., 920, 921, 1013
Goldstone, A., 334, 355, 857, 987, 1013
Goldthwait, D. A., 817, 820, 988, 1057
Gollub, E. G., 480, 1013
Golub. 0. J., 728, 979, 980, 981, 997
Gonda, O., 20, 29, 61, 80, 844, 864, 872,
989, 1013
Gonnard, P., 308, 1013, 1045
Gonzales, E. L., 687, 716, 775, 783, 810,
853, 1058
Gonzalez-Monteagudo, O., 574, 1046
Goodban, A. E., 15, 1043, 1057
Gooder, H., 268, 324, 1013
Goodland, R. L., 524, 1052
Goodman, D. S., 835, 886, 887. 1003,
1013
Goodman, I., 429, 696, 948, 1013
Goodwin, M. E., 571, 573, 991
Goodyer, A. V. N., 925, 1065
Gopinathan, K. P., 490, 847, 1013
Gordon, C. N., 467, 1053
Gordon, H., 407, 684, 692, 833, 1018
Gordon, J. J., 699, 865, 1013
Gore, I. Y., 886, 1045
Gore, M. B. R., 445, 1013
Gorin, G., 671, 672, 673, 1025, 1031
Gosselin, L., 611, 886, 989, 1045
Goth, A., 401, 1013
Gots, J. S., 480, 530, 1013, 1062
Gotto, A. M., 602, 1013
Gottlieb, D., 125, 133, 997, 1013, 1047
Gould, R. G., 886, 1045
Gourevitch, A., 623, 1013
Gouvea, M. A., 868, 939. 940, 1011, 1036
Govorov, N. P., 948, 1013
Grady, J. E., 201, 577, 968, 1055
Graff, M., 587, 1036
Grafflin A. L., 62, 85, 87, 1013
Grana, E., 520, 522, 526, 1003
Grand, R., 60, 357, 1010
Grandjean, E., 956, 1061
AUTHOR INDEX
1087
Granick, S., 161, 162, 674, 859, 888,
1013, 1037, 1062
Grant, B. R., 394, 1013
Grant, P. T., 560, 693, 842, 995, 1013
Grant, W. M., 687, 1013
Grassetti, D. R., 334. 596. 102S, 1029
Grassmann, W., 368, 1013
Graves, D. J., 453, 709, 1028, 1030
Gray, C. T., 52, 187, 228, 229, 1013
Gray, N. M., 62, 85, 87, 1013
Gray, S. J., 687, 1013
Grazi, E., 407, 412, 995. 1013
Green, A. A., 845, 1013
Green, D. E., 16, 60, 61, 62, 64, 146,
237, 259, 338, 348, 407, 437, 509, 549,
555, 599, 684, 692. 832, 833, 843, 994,
1013. 1014. 1018, 1034, 1035, 1052,
1058
Green, D. M., 333, 1012
Green, D. W., 755, 1014
Green, I., 445, 1014
Green, J. W., 908, 1014
Green, L. S., 15, 1049
Green, N. M., 735, 797, 798, 1014
Green, S., 428, 1008
Greenawalt, J. W., 548, 864, 1014
Greenbaum, L. M., 367, 1014
Greenberg, D. M.. 155. 156, 200, 336,
351, 357, 507, 582, 660, 684, 685, 687,
693, 706, 709, 712, 713, 717, 718, 787,
841, 843, 847, 856, 1007, 1014, 1010,
1024, 1026, 1037, 1041, 1042, 1044,
1046, 1052, 1055
Greengard, O., 603, 1014
Greengard, P., 211, 266, 949, 999, 1014
Greenland, R. A., 467, 480, 1068
Greenstein, J. P., 664, 665, 666, 667, 1014
Gregg, D. C, 297, 1014
Gregg, J. R., 199, 882, 964, 965, 1014,
1043
Gregolin, C, 693, 845, 1014
Gregory, M. E., 475, 1033
Greif, R. L., 873, 923, 927, 429, 1014
Greig, M. E., 75, 91, 178, 185, 358, 1002,
1006
Grein, L., 810, 857, 1014
Gremels, H., 879, 927, 1014
Greville, G. D., 128, 175, 177, 180, 183,
187, 210, 552, 789, 793, 816, 819,
866, 869, 998, 1014, 1038, 1061
Grey, E. C, 2, 228, 1014
Griffin, D. H., 595, 1056
Griffith, G. C, 960, 1014
Griffith, W. H., 348, 1014
Griffiths, D. E., 467, 707, 833, 848, 874,
1014, 1017, 1039
Griffiths, M., 816, 820, 821, 866, 869, 1012
Grillo, M. A., 712, 1031
Grimm, M. R., 304, 1028
Grisolia, S., 413, 439, 1014, 1023
Groschel-Stewart, U., 641, 1007
Gromet-Elhanen, Z., 557, 1014
Gros, F., 479, 1014
Gross, R. E., 259, 335, 1014
Grossman, L., 269. 487, 1006, 1014, 1015
Grossowicz, X., 269, 428, 504, 1015
Grouge, V., 978. 1015
Grove,W. E., 701, 702, 723, 724, 725, 1032
Gruber, C. M., 217, 218, 1015
Gruber, W., 122, 814, 861, 1015
Grunberg-Manago, M., 705, 830, 1049
Grunert, R. R., 751, 956, 1015
Guarino, A. J., 413, 1015
Gubler, C. J., 520, 521, 532, 533, 534, 1015
Giinther, G., 61, 987
Guerra, F., 210, 1021
Guest, J. R., 590, 1015
Guggenheim, K., 495, 1015
Guha. S. R., 550, 841, 1015
Gulick, Z. R., 458, 459. 461, 462, 1018
GuUand, J. M., 576, 1015
Gumbmann, M., 75, 80, 91, 121, 1015
Gumnit, R., 573, 1011
Gunja, Z. H., 429, 843, 1015
Gunsalus, I. C, 430, 1004
Gurd, F. R. N., 649, 739, 759, 789, 853,
862, 1015, 1034
Gurd, R. S., 924, 1026
Gurin, S., 22, 27, 74, 77, 124. 148, 150,
995, 1028, 1066
Gurley, H., 926, 1018
Gurtner, H. P., 526, 1015
Gutfreund. H.. 85, 105, 152, 375, 453,
993, 1023
Guthrie, R., 531, 990
Gwin, B. A., 15, 1044
1088
AUTHOR INDEX
H
Haan, J., 1020
Haarniann, W., 737, 753, 1015
Haas, E., 547, 549, '^550, 1015
Haas, H. T. A., 965, 1015
Haas, W. J., 657, 1015
Haavaldsen, R., 857, 1008
Haavik, A. G., 848, 849, 1017
Habeeb, A. F. S. A., 762, 1015
Hachisuka, Y., 195, 1015
Hackett, D. P., 120, 553, 848, 873, 878,
881, 1015, 1041, 1053
Hackney, F. M. V., 296, 1015
Haddow, A., 225, 990
Hafez, E.S.E., 179, 183, 1009
Haft, D. E., 883, 884, 1015
Hagen, U., 956, 957, 1015
Hager, L. P., 453, 1053
Hagihira, H., 265, 394, 1015
Hagstrom, B. E., 726, 1015
Hahn, H., 950, 1015
Hahn, L., 460, 461, 1015
Hahn, M., 974, 1015
Haining, J. L., 887, 1015
Hakala, M. T., 432, 1015
Halasz, P., 949, 1016
Haider, D. K., 530, 576, 999
Halenz, D. R., 853, 1016
Haley, T. J., 942. 948, 1016
Hall, A. N., 321, 1016
Hall, G. A., Jr., 3, 1016
Halliday, K. A., 694, 1051
Halliday, S. L., 504, 505, 1016
Hallman, N., 105, 108, 110, 1016
Halpern, C, 195, 1038
Halpern, Y. S., 269, 498, 504, 1015
Halvorson, H., 326, 351, 423, 552, 553,
710, 772, 782, 848, 1004, 1005, 1016
Halvorson, H. 0., 236, 237, 270, 354,
674, 705, 831, 1040, 1042
Hamilton, H. E., 902, 905, 1053
Hamilton, P. B., 293, 294, 708, 837, 870,
1053, 1054
Hamilton, R. D., 782, 992
Hamilton-Miller, J. M. T., 598, 615, 1016
Hammes, G. G., 788, 803, 1007
Hamolsky, M., 497, 1024
Hampton, A., 466, 467, 481, 1016
Hampton, M. M., 696, 1007
Hand, W. C., 948, 1016
Handler, P., 138, 144, 217, 218, 219,
332, 451, 485, 488, 512, 549, 555, 772,
783, 803, 807, 1009, 1016, 1023, 1027
1034, 1046
Handley, C. A., 856, 925, 933, 1016, 1039
Handschumacher, R. E., 472, 478, 480,
1016, 1044, 1049
Hanly, V. F., 22. 97, 171, 181, 182, 185,
189, 190, 1016, 1061
Hannoun, C, 193, 1016
Hanshoff, G., 707, 779, 780, 1047
Hanson, A., 329, 1016
Hanzlik, P. J., 952, 1016
Happold, F. C, 268, 324, 858, 1013, 1016
Harada, M., 347. 1069
Harary, I., 705, 1016
Hard, J. A., 400, 1006
Hardy, W. G., 201, 670, 992
Harger, R. N., 960, 1008
Hargreaves, A. B., 675, 835, 1016
Harigaya, S., 428, 987
Harkness, D. R., 817, 1018
Harlan, W. R., 399, 1030
Harley, J. L., 51, 53, 169, 172, 1016
Harman, J. W., 119, 992
Harper, E., 553, 1016
Harpur, R. P., 381, 593, 1016
Harris, J., 668, 669, 713, 718, 1016
Harris, J. E., 911, 1016
Harris, J. O., 975, 1016
Harris, S. A., 518, 1067
Harrison, K., 384, 1004
Harting, J., 721, 876, 1016
Hartman, A. M., 2, 225, 1061
Hartman, S. C, 333, 853, 1016
Hartman, W. J., 310, 1016
Hartmann, K.-U., 476, 1016
Hartree, E. F., 259, 283, 696, 698, 1025
Hartsell, S. E., 168, 225, 1052
Hartshorne, D., 713, 856, 1016
Harvey, G. T., 22, 29, 1016
Harvey, S. C., 173, 1017
Hasenfuss, M., 125, 1029
Hashimoto, T., 817, 1017
Hashizume, T., 969, 1025
Haskell, T. H., 621, 998
AUTHOR INDEX
1089
Haskins, F. A., 53, 1017
Haslam, R. J., 153, 1017
Hass, L. F., 390, 412, 853, 1017, 1025
Hassall, K. A., 414, 1017
Hasse, K., 485, 487, 490, 1017
Hasselbach, W., 938, 1017
Hasselbring, H., 2, 225, 1041
Hassid, W. Z., 385. 991
Hastings, A. B., 869, 1007
Hatch, M. D., 53, 77, 79, 80, 82, 173,
830, 853. 1017
Hatefi, Y., 848, 849, 1017
Hatt, D. L., 980, 1047
Haugaard, N., 178, 185, 659, 1017
Hauge, J. G., 591, 839, 1017
Haughton, B. G., 352, 578, 1017
Haurowitz, F., 645, 1017
Hauser, A. D., 920, 921, 1013
Hawtrey, A. O., 79, 1017
Hay, A. J., 427, 429, 812, 814, 838, 1000,
1032
Hayaishi. O.. 226, 228, 230, 324, 853,
1017
Hayano, M., 116, 157, 1000
Hayano, S.,401,m5(?
Hayashi, T., 802, 844, 1019
Hayes, A. D., 958, 959, 960, 1049
Heald, P. J., 134, 176, 184, 1017
Hearon, M., 92, 991
Hecht, L., 798, 865, 1042
Heegaard, E., 516, 996
Hegre, C. S., 853, 1016
Heidelberger, C, 476, 1016
Heilbrunn, L. V., 963, 965, 1017
Heim, F., 750, 881, 884, 885, 927, 1006,
1031
Heimann-Hollaender, E., 340, 1017
Heimbiirger, G., 817, 989
Heiney. R. E., 639, 1027
Heinrich, W.-D., 520, 1063
Heinz, F., 817, 1017
Heinz. R., 701, 724, 1017
Heinzelman, R. V., 358, 1052
Hellerman, L., 38, 40, 42, 60, 61, 76,
240, 329, 340, 341, 346, 347, 547, 549,
556, 557, 558, 560, 640, 643, 657, 658,
662, 664, 665, 666, 668, 669, 673, 676,
683, 686, 697, 701, 702, 703, 704, 706,
713, 717, 718, 742, 772, 780, 789, 803,
816, 819, 876, 878, 950, 995, 998,
999, 1003, 1010, 1011, 1016, 1017,
1018, 1041, 1045, 1047, 1048, 1051
Hellig, H.,711,852, iW<5
Helmert, E., 685, 1036
Helmreich, E., 263, 387, 392, 394, 1018
Hemker, H. C, 547, 549, 550, 556, 558,
1018
Hemming, H. G., 618, 632, 1034
Henderson, J. F., 481, 1018
Henderson, J. H. M., 171, 1018
Henderson, L. M., 610, 709, 772, 782,
843, 1003, 1038, 1057
Henderson, M. J.. 125, 1039
Hendley. D. D., 463, 1018
Hendlin, D., 582, 589, 1018
Hendricks, S. B., 122, 1022
Henle, W., 530, 1023
Henning, U., 887, 987
Hepler, O. E., 924, 926, 1018, 1054
Hepp, P., 931, 941, 1018
Heppel, L. A., 471, 473, 817, 988, 1018,
1021
Herbain, M., 518, 1063
Herbert, D., 407, 684, 692, 833, 1018
Herbert, E., 864, 1018
Herbert, M., 77, 78, 1019
Heredia, C. F., 387, 388, 400, 547, 554,
1018, 1056
Herken, H.. 499, 965, 996, 1018, 1067
Herman, E., 927, 947, 1043
Herner, B., 348, 1018
Herring, P. J., 384, 1018
Herriott, R. M., 683, 688, 1018
Herrmann, H., 465, i005
Herschberg, A. D., 835, 1010
Herz, R., Jr., 208, 911, 913, 1018
Herzog, R. O., 974, 1018
Herzog, U., 750, 881, 884, 885, 1006
Hess, R., 850, 1018
Hess, S. M., 316, 317, 318, 319, 320, 1018
Hesselbach, M. L., 128, 177, 1005
Hested, D. M., 208, 1063
Hestrin, S., 421, 1018
Heydeman, M. T., 831, 832, 989, 1018
Heymann, H., 297, 458, 459, 461, 462,
1018
1090
AUTHOR INDEX
Heymans, J. F., 1, 187, 218, 1018
Heyndrickx, A., 747, 748, 1058
Heyworth, R., 419, 420, 1064
Hiai, S., 292, 293, 1018
Hiatt, A. J., 33, 166, 855, 1018
Hiatt, R. B., 696, 948, 1013
Hicks, S. P., 499, 504, 670, 724, 965,
1018
Hierholzer, K., 924, 1026
Hietanen, S., 736, 739, 1018
Hift, H., 569, 1054
Higgins, E. S., 554, 614, 814, 1018, 1066
Hildebrandt, A. C, 197, 1019
Hilden. T., 918, 923, 996
Hill, B. R., 772, 774, 845, 1019
Hill, R. L., 1019
Hill, R. M., 41, 177, 1020
Hillman, R. S. L., 574, 575, 1022
Hillmann, G., 910, 1051
Hilmoe, R. J., 817, 1018
Hilton, J. L., 588, 597, 1019, 1062
Hilz, H., 693, 818, 1019
Himms, J. M., 363, 994
Himwich, H. E., 500, 990
Hinman, R. L., 326, 1016
Hino, S., 292, 293, 1018
Hirade, J., 802, 844, 873, 1019
Hiramitsu, S., 179, 1059
Hirano, S., 126, 127, 135, 153. 176, 1059,
1061
Hirashima, K., 968, 969, 1019
Hird, F. J. R., 657, 1019
Hirohata, R., 792, 813, 1043
Hirschhorn, L., 724, 993
Hitchings, G. H., 283, 429, 584, 1006,
1013, 1019, 1068
Hitchcock, P., 212, 1019
Hjort, A. M., 956, 990
Ho, J. Y. C, 639, 1027
Hoadley, L., 963, 964, 1019
Hoare, D. S., 593, 708, 1003
Hoare, J. L., 762, 1019
Hoberman, H. D., 61, 1019
Hoch, F. L., 743, 746, 780, 785, 789,
806, 825, 831, 1019, 1055
Hoch, G. E., 293, 1019
Hochachka, P. W., 169, 1019
Hochster, R. M., 18, 30, 31, 132, 261,
474, 511, 553, 555, 847, 878, 994,
1019, 1057, 1063
Hockenhull, D. J. D., 77, 78, 422, 1019
Hofling, E., 4, 1019
Hogberg, B., 461, 464, 1004, 1008
Hohl, R., 122, 1015
Hokfelt, B., 401, 1019
Hofmann, E., 396, 1019
Hofmann, E. C. G., 487, 489, 493, 1019
Hofmann, K., 589, 989
Hofstee, B. H. J., 286, 288, 289, 457,
676, 718, 721, 1019, 1054
Hogg, J. F., 263, 389, 390, 391, 1041
Hoggarth, E., 224, 236, 1003
Hohnholz-Merz, E., 802, 1044
Holden, M., 693, 712, 1019
Holland, J., 71, 74, 79, 80, 81. 82, 173,
203, 273, 274, 1019
Holland, J. F., 531, 581, 990, 1019
Holland, W. C, 936, 989
Hollander, P. B., 214, 625, 896, 943,
944, 945, 1065
Holldorf, A., 430, 841, 1020
Holliday. W. M., 494, 992
HoUis, V. W., Jr., 449, 1034
HoUocher, I. C, Jr., 18, 22, 1000, 1019
Hollunger, G., 17, 998
Holly, F. VV., 538, 1019
Holmberg, C. G., 691, 693, 1019
Holmes, W. L., 473, 1020
Holms, W. H., 124, 1003
Holt, A., 803, 1020
Holf, C. v., 518, 1066
Holtkamp, D. E., 41, 177, 1020
Holtman, D. F., 224, 1012
Holton, F. A., 32, 35, 61, 80, 83, 121, 122,
1020, 1055
Holton, R., 195, 1058
Holtz, P., 359, 577, 1020
Holz, G., 841, 993
Holzer, E., 1020
Holzer, H., 430, 432, 695, 817, 841, 852,
1020
Hommes, F. A., 18, 1020
Honda, S. I., 23, 27, 33, 46, 47, 51, 170,
1020
Hopkins, F. G., 18, 25, 34, 35, 41, 661,
662, 664, 1020
AUTHOR INDEX
1091
Hopper, S., 334, 358, 810, 856, 1020,
1052
Horecker, B. L., 412, 413, 471, 855,
995, 1021, 1059
Hori, K., 549, 554, 555, 837, 1020
Horibata, K., 400, 1000
Horio, T., 228, 229, 557, 880, 1020
Horning, M. G., 234, 1020
Horowitz, M. G., 639, 760, 767, 1020,
1027
Horwitt, M. K., 456, 1020
Horwitz, L., 881, 892, 1020
Hoshino, M., 543, 1020
Hoskins, D. D., 783, 847, 1020
Hosoya, N., 55, 78, 147, 149, 179, 228,
233, 1020
Hosoya, T., 686, 1020
Hospelhorn, V. D., 681, 703, 762. 1023
Hostynova, D., 954, 957, 1005
Hotchkiss, R. D., 910, 1020
Houck, C. R., 888, 889, 890, 897, 1020
Houck, J. C, 460, 461, 462, 1020
Houlihan, R. K., 37, 241, 1052
Houwing, C, 980, 1024
Howard, R. L., 849, 1026
Howe, W. B., 359, 1020
Howell, R. S., 574, 577, 1051
Howes, W. v., 61, 1034
Hsieh, K., 927, 1053
Huang, H. T., 271, 1020
Hubbard, D. M., 953, 1064
Hubbard, J. S., 267, 613, 1005, 1020
Hubbard, N., 888, 1029
Hubbard, R. W., 459, 993
Hudson, M.T., 385, 391, 392, 1020, 1068
Hudson, P. B., 440, 441, 442, 711, 713,
788, 839, 842, 1032, 1061
Hudson, P. S., 452, 1061
Hiibscher, G., 284, 662, 713, 1000, 1035
Hulsmann, W. C, 86, 547, 549, 550, 556,
558, 1018, 1021
Huennekens, F. M., 539, 543, 544, 553,
581, 582, 585, 711, 848, 851, 864, 993,
997, 1020, 1043, 1051, 1054, 1066
Huf, E. G., 670, 690, 1006
Huffaker, R. C, 225, 226, 1021
Huggins, C, 681, 703, 762, 1023
Hughes, A. F. W., 199, 742, 1021
Hughes, C, 852, 1021
Hughes, D. E., 550, 845, 1009, 1021
Hughes, W. J., Jr., 757, 1021
Hughes, W. L., Jr., 681, 743, 744, 748,
750, 753, 754, 755, 757, 758, 759, 761,
862, 930, 1005, 1021
Huisman, I. H. J., 755, 760, 1021
Hulme, A. C, 120, 1023
Hulpieu, H. R., 520, 1067
Hultquist, G., 401, 1019
Hummel, J. P., 464, 475, 1021, 1041
Humphrey, B. A., 28, 1021
Humphrey, G. F., 22, 28, 71, 74, 79, 80,
81, 82, 173, 174, 203, 273, 274, 713,
882, 884, 1019, 1021
Humphreys, S. R., 400, 401, 496, 1024,
1030
Humphreys, T. E., 136, 1024
Hundley, J. M., 495, 1033
Hunter, A., 335, 1021
Hunter, D., 951, 952, 953, 954, 1021
Hunter, F. E., Jr., 210, 873, 883, 1021
Hunter, F. R., 22, 23, 54, 58, 1021
Hunter, G. J. E., 164, 238, 1005, 1021
Hunter, N. W., 27, 28, 1021
Hunter, R., 27, 1021
Hunter, W. C, 985, 1021
Hunter, W. R., 882, 961, 1021
Hunter, W. S., 173, 183, 1063
Hurd, C. D., 619. 997
Hurlbert, R. B., 15, 28, 997
Hurwitz, A., 210, 1021
Hurwitz, J., 413, 471, 475, 477, 564,
832, 855, 1021
Hurwitz, L., 875, 1021
Husa, W. J., 196, 1033
Huszak, S., 124, 138, 140, 177, 1021
Hutchinson, D. J., 582, 583, 996, 1009
Huxley, J. S., 964, 1021
Hylin, J. W., 33, 244, 1021
Hynd, A., 384, 1018, 1021
lacono, J., 574, 1040
Ibsen, K. H., 396, 397, 1021
Ichihara, A., 857, 1021
1092
AUTHOR INDEX
Ichihara, E. A., 857, 1021
Ichihara, K., 64, 564, 578, 676, 858,
1022, 1039, 1043, 1064
Igarasi, H., 225, 1022
Iliffe, J., 149, 1022
Illingsworth, B., 561, 1022
Imaizumi, R., 611, 612, 1042
Imamoto, F., 27, 1022
Imamura, K., 868, 1061
Imshenetsky, A. A., 983, 984, 985, 1022
Inada, A., 194, 999
Inagaki, T., 741, 1022
Ingold, C. K., 5, 1011
Ingraham, R. C, 910, 1022
Ingram, V. M., 649, 755, 1014, 1022
Inkson, R. H. E., 133, 1040
Inoue. F., 578, 676, 858, 1064
Irvin, E. M., 556, 1022
Irvin, J. L., 556, 1022
Isaka, S., 678, 727, 1022
Ishervvood, F. A., 64, 773, 774, 1002,
1035
Tshii, K., 983, 984, 987
Ishikawa, S., 873, 1022
Ishimoto, M., 551, 554, 555, 684, 1010.
1022
Ishishita, Y., 179, 1059
Isles, T. E., 663, 1022
Itatani, M. K., 175, 194, 1039
Ito, E., 439, 1022
Ito, H., 578, 676, 858, 1064
Ito, K., 578, 676, 858, 1064
Ives, D. H., 481, 1022
Ives, D. J., G., 9, 1002
Ivler, D., 228, 230, 231, 1067
Iwainsky, H., 168, 228, 237, 504, 1040
Iwasa, K., 27, 1022
Iwatsubo, M., 547, 559, 1022
Iwatsuka, H., 892, 1022
Izaki, K., 349, 1038
Izar, G., 920, 1022
J
Jackson, D. E., 918, 944, 946, 947, 952,
1022
Jackson, J. F., 510, 989
Jackson, L. J., 599, 1051
Jackson, P. C., 122, 1022
Jackson, W. T., 709, 1030
Jacob, H. S., 877, 904, 906, 908, 912,
1022
Jacob, M., 856, 873, 1022
Jacobs, E. E., 856, 873, 1022
Jacobs, F. A., 574, 575, 1022
Jacobs, G. S., 873, 927, 929, 1014
Jacobs, H. I., 212, 1022
Jacobsen, C. F., 646, 1032
Jacobsohn, K., 279, 1022
Jacobson, K. B., 154, 503, 511, 586,
1022
Jacobson, L., 170, 209, 273, 274, 1043
Jacoby, G. A., 305, 1022
Jacoby, M., 794, 1022
Jacquez, J. A., 266, 1022
Jaattela, A. J., Q\\, 1022
Jaenicke, L., 585, 1022
Janisch, W., 923, 1067
Jarnefett, J., 548. 1023
Jagendorf, A. T., 851, 874, 891, 892,
989, 1022, 1024
Jahn, F., 701, 723, 724, 725, 727, 1022
Jakoby, W. B., 60, 547, 550, 595, 1022,
1023, 1041, 1069
James, W. O., 171, 1023
Janacek. K., 912, 950, 1023, 1032
Jandl, J. H., 877, 904, 900, 908, 912,
1022
Jang, R., 411, 657, 989, 1023
Jankelson, O. M., 208, 1063
Jann, G. J., 33, 38, 1058
Jansen, B. C. P., 519, 774, 775, 854,
1043
Jansen, E. F., 657, 1023
Jansz, H. S., 561, 1022
Jasmin, R., 586, 1023
Jecsai, G., 804,-809, 1023
Jedeikin, L. A., 149, 151, 1023
Jeffree, G. M., 443, 464, 1023
Jencks, W. P., 887, 1023
Jenerick, H. P., 211, 1023
Jenkins, W. T., 64, 334, 1023
Jenrette, W. V., 664, 667, 1014
Jensen, E. V., 681, 703, 762, 1023
Jensen, K. B., 15, 225, 1023
Jermstad, A., 15, 225, 1023
Jerstad, A. C, 961, 1038
AUTHOR INDEX
1093
Jervis, E. L., 265, 1023
Jiracek, V., 389, 1027
Joachimoglu, G., 875, 1023
Jocelyn, P. C, 663, 1022
Jodrey, L. H., 54, 174, 1023
Johnson, C. E., 859, 1060
Johnson, F. H., 676, 1053
Johnson, G. T., 28, 1023
Johnson, J. E., 941, 943, 1049
Johnson, M., 199. 997
Johnson, M. A., 597, 846, 1023
Johnson, M. P., 911, 1023
Johnson, O. H., 538, 1006
Johnson, R., 944, 945, 1007
Johnson, W., 586, 1023
Johnson, W. A., 55, 58, 98, 99, 104, 110,
1028
Johnson, W. J., 504, 505, 601, 1023
Johnston. R. L., 942. 1023
Johnstone. J. H., 384. 1023
Johnstone, R. M., 338, 1023
Jokhk, W. K., 685, 709, 711, 1023
Jolley, R. L., 132, 1023
Jonas, R. E. E., 137, 994
Jones, E. A., 85, 105, 152, 453, 1023
Jones, J. D., 120, 1023
Jones, J. H., 530. 1023
Jones, J. R., 882, 963, 1023
Jones, L. O., 576, 1011
Jones, M., 156, 1011
Jones, O. T. G., 267, 547, 553. 1023, 1041
Jones, R. F., 169, 1056
Jordan, H. V., 632, 1008
Jorgenson, C. R., 263, 403, 1067
Josephson, K., 685, 1063
Joshi, J. G., 512, 783. 1023
Joshi, J. v., 15, 987
Josten, J. J., 547, 553, 847, 1047
Jowett, M., 78, 87, 138, 144, 176, 177,
238, 349, 613, 735, 875, 1023
Joyce, B. K., 413, 439, 1014, 1023
Joyce, C. R. B., 901, 905, 1024
Judah, J. D., 81, 999
Judis, J., 294, 1011
Jukes, T. H., 581, 1003
Jung, F., 900, 901, 902, 906, 1024
Junowicz-Kocholaty, R., 349, 1039
Jurtshuk, P., Jr., 843, 849, 1017, 1052
K
Kappner, W., 982, 1024
Kagawa, Y., 163, 888, 1024
Kahana, S. E., 407, 474, 1024
Kahn, J. S., 874, 1024
Kaisch, K., 674, 816, 1028
Kaiser, C, 261, 1024
Kaiser, E., 459, 1024, 1043
Kalbe. H.. 225. 1060
Kalckar, H. M., 285, 286, 287, 288, 859,
1024, 1037
Kaldor, G., 548, 551, 556, 1024
Kallen, R. G., 147, 1024
Kalmus. A., 285, 992
Kalner, H. S.. 383, 396, 1047
Kalnitsky, G., 18, 26, 90, 650, 826, 855,
991, 1024, 1047
Kaltenbach, J. P., 18, 26, 1024
Kalyankar, G. D., 15, 1024
Kamen, M. D., 293, 557, 875, 880, 996,
1020, 1056
Kameyama, T., 551, 555, 1022
Kamin, H., 341, 817, 850, 1027, 1066
Kamrin, A. A., 573, 1011, 1024
Kamrin, R. P., 573, 1011, 1024
Kanamori, M., 708, 1024
Kandler, O., 163, 1024
Kang, H. H., 610, 772, 782, 843, 1003,
1038
Kann. E. E., 189, 1024
Kantarjian, A. D., 952, 954, 1024
Kaper, J. M., 980, 1024
Koplan, C., 979, 1024
Kaplan, E. H., 348, 1024
Kaplan, L. A., 505, 1004, 1010
Kaplan, N. O., 211, 434, 485, 487, 490,
493, 496, 497, 498, 500, 503, 504,
507, 508, 509, 511, 512, 784, 786, 807,
808, 846, 849, 850, 851, 980, 981,
988, 1003, 1004, 1014, 1015, 1022,
1024, 1025, 1041, 1053, 1060, 1062,
1064, 1069
Kara, J., 474, 1055
Karasek, M. A., 706, 713, 718, 1024
Karlsson, J. L., 227, 228, 1024
Karmen, A., 234. 1020
Karnofsky, D. A., 576, 1024
1094
AUTHOR INDEX
Karnovsky, M. L., 435, 104S
Karpeiskii, M. Y., 359, 102G
Karunaivatnam, M. C, 61, 424, 1024
Kasamaki, A., 78, 79, 86, 349, 1024
Kashiwabara, E., 618, 632, 1062
Kashket, E. R., 120, 1024
Kashket, S., 487, 492, 988
Kassanis, B., 976, 979, 1024
Kassel, B., 665, 666, 667, 995
Kasser, I. S., 937, 1066
Kassowitz, H., 952, 1044
Katagiri, H., 543, 554, 1059
Katagiri, M., 853, 1017
Katchalski, E., 457, 758, 761, 1005, 1024
Kato, H., 195, 1015
Kato, I., 416, 417, 1025
Kato, S., 399, 400, 1056
Kato, Y., 542, 1059
Katoh, S., 787, 1025
Katoh, T., 554, 1025
Katsuya, H., 969, 1025
Katyal, J. M., 671, 672, 673, 1025
Katz, A. M., 939, 940, 1025
Katz, H., 664, 1028
Katz, J., 145, 226, 234, 1025
Katz, R. L., 532, 1041
Katz, S., 741, 980, 1025
Katzman, P. A., 839, 1004
Kauder, E. M., 841, 993
Kaufman, B., 849, 851, 1025
Kaufman, B. T., 211, 816, 1025
Kaufman, S., 369, 1025
Kavanau, J. L., 937, 1025
Kawabata, S., 224, 1025
Kawada, N., 55, 78, 147, 149, 179, 228,
233, 1020
Kawai, F., 675, 685, 692, 771, 772, 835,
999
Kawano, Y., 224, 1025
Kawasaki, E. H., 52, 1026
Kay, C. M., 759, 1025
Kayser, F., 983, 984, 1011
Kaziro, Y., 475, 853, 1025
Kearney, E. B., 16, 17, 18, 32, 33, 38,
42, 45, 46, 48, 539, 541, 542, 545,
547, 549, 713, 783, 825 856, 1025,
1054
Kearns, C. W., 675, 1032
Keay, L., 551, 709, 843, 1048
Keech, D. B., 676, 853, 1025
Keelee, M. M., 939, 999
Keighley, G., 887, 995
Keil, J. G., 226, 228, 992
Keilin, D., 22, 23, 259, 283, 696, 698,
1025
Keister, D. L., 510, 891, 1025
Kekwick, R. G. O., 836, 1066
Keleti, T., 409, 1025
Kellaway, C. H., 949, 1007
Keller, D. M., 81, 210, 1025
Keller, V., 529, 1051
Kellerman, G. M., 466, 1025
Kelley, J. F., 463, 993
Kelly, F. J., 928, 997
Kelly, S., 169, 209, 1025
Kelner, A., 349, 1039
Kench, J. E., 920, 1000
Kennedy, E. P., 137, 1031
Kennedy, J., 348, 363, 1024, 1051
Kenney, F. T., 713, 857, 1025
Kenten, R. H., 524, 1025
Keodara, J. C, 1025
Kerby, G. P., 459, 1025
Kerly, M., 81, 176, 183, 997
Kermack, W. 0., 551, 560, 593, 995,
1009, 1025
Kernan, R. P., 212, 1026
Kerr, D. S., 817, 820, 988
Keskin, H., 60, 996
Kessler, D., 980, 1026
Kessler, R. H., 206, 918, 923, 924, 928,
929, 930, 933, 935, 995, 1026, 1058
Keston. A. S., 413, 414, 1026
Ketchel, M., 199, 1051
Kharasch, N., 637, 639, 1026, 1043
Khomutov, R. M., 359, 1026
Khorana, H. G., 473, 1047
Kielley, R. K., 121, 445, 446, 1026
Kielley, W. W., 121, 445, 446, 807,816,
866, 1026
Kies, M. W., 693, 1056
Kiesow, L., 387, 395, 678, 1026
Kiessling, K. H., 515, 518, 1026
Kihara, H., 833, 1026
Kilbourne, E. D., 400, 1026
Kilsheimer, G. S., 238, 440, 1026
AUTHOR INDEX
1095
Kimmel, J. R., 375, 769, 770, 804, 1026,
1055
Kimura, H., 137, 1026
Kimura, T., 845, 1026
Kimura, Y., 124, 135, 176, 184, 1026
King, C. G., 259, 1040
King, E. E., 943, 944, 956, 957, 10:il
King, E. J., 440, 987
King, H. K., 352, 578, 780, 783, 810, 814,
1017, 1059
King, R. L., 1053
King, T. E., 22, 33, 38, 552, 705, 712,
832, 849, 853, 1002, 1025, 1026, 1033,
1050
Kini, M. M., 76, 135, 153, 1026
Kinoshita, S., 542, 1059
Kinoshita, T., 969, 1025
Kjaer, A., 617, 632, 996
Kjeldgaard, N. D., 285, 286, 287, 288,
1024
Kinsey, V. E., 687, 1013
Kinsky, S. C, 850, 1026
Kipnis, D. M., 387, 388, 390, 1026
Kiraly, Z., 170, 1007
Kirkham, D. S., 195, 1026
Kirkman, H. N., 839, 1026
Kirkpatrick, H. C, 979, 1032
Kirshner, N., 399, 1030
Kishimoto, U., 866, 1026
Kistiakowsky, G. B., 64. 603, 610,
1026
Kistner, S., 547, 1026
Kit, S., 155, 200, 1026
Kitagawa, S., 816, 866, 867, 868, 1061
Kiyomoto, A., 428, 987
Klavano, P. A., 957, 959, 961, 1038
Kleczkowski, A., 976, 979, 1024
Klein, A. O., 675, 1026
Klein, H. P., 269, 1006
Klein, J. R., 152, 341, 342, 347, 350,
485, 488, 1016, 1027
Klein, M., 978, 979, 980, 1027, 1044
Klein R. F., 399, 1030
Klein, R. M., 74, 171, 189, 595, 1032,
1046
Klein, W., 472, 1027
Kleiner, I. S., 202, 994
Kleinfeld, M., 942, 945, 946, 1027, 1057
Kleinspehn, G. G., 591, 1061
Kleinzeller, A., 267, 908, 1027
Kleitman, N., 942, 944, 946, 1050
Klemperer, H. G., 688, 881, 910, 912,
1027
Klempien, E. J., 693, 818, 1019
Klenk, L., 368, 1013
Klenow, H., 285, 286, 287, 288, 1024
Kleppe, K., 389, 1044
Klimek, J. W., 973, 983, 1027
Kline, D. L., 150, 151, 1027
Klingenberg, M., Gl., 178, 1027
Klingman, J. D., 332, 1027
Klotz, I. M., 29, 32, 35, 37, 242, 243,
639, 757, 760, 767, 1020, 1027, 1049,
1060
Klotz, T. A., 757, 1027
Klybas, V., 408, 409, 1046
Knell, J., 314, 315, 1066
Knight, S. G., 708, 837, 870, 1054
Knobloch, N., 197, 1067
Knopfmacher, H. P., 683, 685, 691, 692,
1027
Knox, R., 615, 697, 699, 1016, 1045
Knox, W. E., 686, 1056
Kobashi, Y., 225, 362, 1027, 1064
Koch, R., 970, 1027
Koch, W., 352, 1066
Kochakian, C. D., 848, 1006
Kocholaty, W., 349, 687, 1027, 1039
Kocouvek, J., 389, 1027
Kodicek, E., 886, 1060
Koditschek, L. K., 582, 1018
Koedam, J. C, 521, 522, 525, 1027
Koelle, E. S., 698, 699, 700, 1012, 1045
Konig, S., 396, 1019
Koeppe, O. J., 812, 1027
Koffler, H., 77, 237, 992, 1013
Kohler, A., 291, 1041
Kohler, A. R., 583, 996
Kohn, P., 376, 388, 1030
Koike, K., 817, 1027
Koike, M., 179, 578, 1035, 1059
Koishi, T., 204, 883, 921, 1027
Koisumi, T., 578, 1064
Koivusalo, M., 349, 1027
Koizumi, T., 676, 858, 1064
Kolesar, P., 954, 957, 1005
1096
AUTHOR INDEX
Kolthoff. I. M., 671. 747, 748, 763, 1027,
1058
Kominz, D. R., 789, 866, 868, 939,
1027
Kondo, K., 675, 685, 692, 771, 772, 835,
999
Kondo, S., 439, 1022
Kondo, Y., 551, 555, 1022
Kono, M., 382, 1027
Kono, T., 896, 912, 1027
Konrad, fi., 170, 1007
Konsanszky, A., 503, 1037
Kopac, M. J., 201, 998
Korey, S., 696, 723, 938, 1027
Korey, S. R., 705, 830, 1049
Korkes, S., 410, 1058
Korn, E. D., 463, 1027
Kornberg, A., 226, 228, 446, 470, 1017,
1042, 1070
Kornberg, H. L., 71, 594, 602, 845. 1004,
1009, 1013, 1027
Kornguth, S. E., 456, 1027
Koshland, D. E., Jr., 868, 1032
Kostif, J., 389, 1027
Kostytschew, S., 875, 1028
Kosunen, T., 109, 1054
Kotaka, S., 541, 1028
Kotyk, A., 267, 1027
Koukol, J., 355, 852, 1028
Kovac, L., 49, 267, 1027, 1028
Kovachevich, R., 885, 1028
Koval, G. J., 601, 856, 987
Kowa, Y., 148, 228, 1029
Krahe, E., 974, 1028
Krahl, M. E., 198, 1028
Krajci-Lazary, B., 926, 954, 957, 1005
Kramer, M., 236, 1028
Kramer, P. J., 209, 1028
Kramer, S., 29, 1034
Krampitz, L. O., 26, 89, 168, 1051, 1059
Krane, S. M., 262, 1028
Krasna, A. I., 279, 293, 294, 676, 1008,
1028
Krause, R. M., 980, 1026
Kravitz, E., 74, 77, 1057
Kream, J., 288, 1053
Krebs, E. G., 289, 453, 1028
Krebs, H. A., 22, 27, 55, 58, 70, 71, 72,
74, 75, 77, 89, 94, 96, 98, 99, 104,
110, 115, 116, 124, 153, 158, 332, 333,
769, 830, 1017, 1027, 1028, 1060
Krejci, L. E., 687, 1027
Kreke, C. W., 744, 810, 835, 856, 870,
871, 880, 1001, 1028, 1052, 1055, 1056
Kretszchmar, R., 513, 1062
Krishna Murti, C. R., 692, 831, 987
Krishnaswamy, P. R., 15, 660, 854,
1012, 1024
Kriszat, G., 726, 964, 1028, 1049
Kroger, M. H., 856, 870, 871, 1028
Kronenberg, G. H. M., 891, 1043
Krop, S., 956, 957, 1038
Krueger, A. P., 976, 979, 980, 1028
Krueger, R., 876, 938, 1028
Krueger, R. C, 300, 1028
Kruse, R., 922, 1007
Kruse, W. T., 620, 622, 623, 624, 625,
627, 628, 630, 631, 1052, 1068
Krusius. F. E., 109, 110, 1028
Kubowitz, F., 768, 1028
Kuby, S. A., 60, 836, 1028
Kuczynski, M., 878, 989
Kuhn, J., 791, 794, 817, 820, 1000
Kuhn, R., 259, 421, 664, 665, 666, 668,
687, 712, 853, 1028
Kulka, R. G., 444, 445, 1001
Kull, F. C, 304, 1028
Kumar, S. A., 660, 685, 1047
Kumin, S., 160, 1053
Kummerovv, F. A., 137, 1026
Kun, E., 334, 355, 593, 596, 767, 1008,
1011, 1028, 1029
Kuner, E., 945, 946, 1011
Kuno, M., 892, 1022
Kuno, T., 195, 1015
Kunz, H. A., 519, 531, 1029
Kunz, W., 40, 1029
Kuo, M. H., 440, 443, 1029
Kupiecki, F. P., 706, 834, 1029
Kuratomi, K., 519, 676, 774, 783, 843,
854, 1029
Kurtz, A. N., 373, 1029, 1064
Kurup, C. K. R., 615, 1029
Kuschinsky, G., 937, 938, 939, 1029,
1061
AUTHOR INDEX
1097
Kusunose, E., 26, 36, 62, 63, 148, 228,
855, 1029, 1069
Kusunose, M., 26, 36, 62, 63, 148, 228,
855, 1029. 1069
Kutscha, W., 938, 1029
Kutscher, W., 125, 1029
Kuttner, E., 298, 299, 301, 1029
Kvam, D. C, 478, 1029
Kvamme, E., 104, 111, 126, 1029
Kwientny, H., 281, 992
Kwienthy-Govrin, H.. 285, 992, 993
Labbe, R. F., 888, 1029
Labeyrie, F., 435, 437, 547, 559, 1022,
1029
Lachnit, V., 1045
Lack, L., 160, 1068
Ladd, J. N., 15, 52, 1029
La Du, B. N., 272, 305, 306, 595, 851,
1012, 1022, 1029, 1069
Ladygina, M. E., 170, 1049
Lagnado, J. R., 816, 1029
Laki, K., 63, 1029
Lakshmanan, M. R., 547, 549, 837,
1029
Laland, S. G., 384, 1005
Lamberg, S. L., 540, 544, 993
Lambie, A. T., 921, 1029
Lambooy, J. P., 537, 538, 539, 988,
1029, 1051
Lamprecht, W., 817, 1017
Landau, B., 392, 400, 401, 1030
Landau, B. R., 263, 387, 388, 401, 403,
404, 1029, 1067
Landau, J. V., 964, 965, 1029, 1070
Lands, W. E. M., 372, 1029
Landon, E. J., 282, 864, 1029
Lane, M., 538, 1029, 1030
Lane, M. D., 853, 1016
Lang, C. A., 851, 1030
Lang, H. M., 891, 1050
Lang, K., 601, 1030
Langdon, R. G., 412, 781, 815, 817,
839, 841, 999, 1030, 1038, 1045
Lange, C. F., Jr., 376, 388, 1030
Lange, R., 662, 663, 697, 699, 1045
Langer, L. J., 781, 1030
Lansford, E. M., Jr., 354, 1001
Lara, F. J. S., 16, 49, 225, 226, 435, 547,
551, 557, 558, 710, 717, 773, 778, 781,
844, 1030, 1037, 1038, 1054
Lardy, H. A., 60, 63, 77, 87, 119, 124,
128, 176, 203, 377, 381, 382, 383, 475,
710, 779, 783, 798, 836, 846, 869,
1001, 1028, 1030, 1035, 1042, 1044,
1045, 1050
Laris, P. C., 865, 912, 1030
Larner, J., 391, 706, 709, 710, 718, 1030,
1049
Laroche, M. J., 592, 611, 989
Larrabee,M.G., 211, 1050
Larson, E. R., 404, 1066
Larson, F. C., 602, 1030
La Sala, E. F., 622, 991
Lascelles, J., 163, 888, 1030
Lassen, U. V., 267, 1030
Lasser, N. L., 38, 40, 42, 240, 1018
Laszlo, J., 388, 392, 399, 400, 401,
1029, 1030
Lathe, G. H., 428, 1030
Laties, G. G., 78, 79, 87, 91, 94, 132,
170, 171, 173, 181, 182, 183, 185, 189,
616, 1030, 1044
Latuasan, H. E., 538, 1030
Latzkovits, L., 179, 1004
Launoy, L., 984, 985, 1030
Laurence, E. B., 200, 997
Laver, W. G., 751, 888, 1012
Lavik, P. S., 856, 925, 1016
Lawrence, J. C, 388, 389, 391, 996, 998
Lazarow, A., 175, 1000, 1010
Lazarus, A. S., 968, 972, 1050
Lazdunski, M., 711, 772, 773, 1030
Lazzarini, R. A., 512, 553, 851, 1030
Lea, D. J., 321, 1016
Leaback, D. H., 419, 1046
Leach, F. R., 610, 782, 843, 1003
Leach, S. J., 763, 1030
Le Bras, G., 356, 1057
Lebrun, J., 203, 1030
Leder, I. G., 781, 839, 1030
Lederer, E., 683, 1040
Ledingham, G. A., 169, 195, 1007
Ledoux, L., 712, 715, 716, 744, 815, 1030
1098
AUTHOR INDEX
Lee, C. P., 18, 33, 1030
Lee, H. A., Jr., 590, 1030
Lee, J. S., 228, 1031
Lee, Y.-P., 674, 697, 774, 804, 852, 1031
Lees, H., 450 451, 547, 551, 988, 997,
1031
Le Fevre, C. G., 618, 1031
Le Fevre, P. G., 264, 690, 905, 906, 911,
912, 1031
Le Fevre, R. J. W., 618, 1031
Legge, J. W., 707, 1031
Lehman, I. R., 462, 849, 1031
Lehman, R. A., 943, 944, 956, 957,
1003, 1031
Lehnann, J., 1031
Lehmnger, A. L., 91, 119, 137, 138, 178,
705, 872, 873, 874, 1001, 1022, 1031,
1064
Leiby, C. M., 316, 317, 318, 1045
Leigh, J., 63, 1012
Lein, J., 623, 1013
Leloir, L. F., 61, 144, 476, 594, 1005,
1014, 1031
Lembeck, F., 611, 1031
Lenney, J. F., 791, 793, 1031
Lenti, C, 712, 1031
Lentz, C. P., 677, 1042
Lenz, G. R., 739, 1031
Lenzi, F., 215, 1031
Leopold, A. C, 881, 967, 1041
Le Page, G. A., 168, 1063
Leppla, W., 518, 1066
Lerner, A. B., 304, 1031
Leslie, J., 672, 1031
Lester, G., 418, 1031
Letnansky, K., 125, 399, 1031. 1052
Leuschner, F., 750, 927, 1031
Leuthardt, F., 157, 158, 1007, 1040
Levaditi, C, 984, 985, 1030
Levenberg, B., 333, 1031
Levey, H. A., 31, 50, 176, 1031
Levey, S., 458, 1031, 1036
Levi, A. A., 224, 995
Levin, G., 281,992
Levine, H. B., 168, 1011
Levine, M., 189, 1044
Levine, S., 705, 714, 717, 718, 803, 805,
831, 991
Levinthal,C., 439, 1011
Levitt, M. F., 920, 921, 1013
Levvy, G. A., 61, 272, 424, 426, 427,
428, 429, 1000, 1008, 1024, 1031,
1032, 1036
Levy, H. M., 868, 1032
Levy, H. R., 449, 712, 713, 1032
Levy, J. F., 210, 1021
Levy, R. I., 932, 933, 934, 935, 1032,
1065
Lewin, J., 758, 1032
Lewis, G., 954, 994
Lewis, H. B., 236, 999
Lewis, S. E., 80, 85, 121, 1032
Li, C. H., 681, 1032
Li, S. C, 838, 1032
Li, T.-K.. 785, 1032
Li, Y., 838, 1032
Lichtenstein, N., 336, 340, 1017, 1032
Lieb, H., 4, 1019
Liebecq, C, 63, 1032
Lieben, F., 657, 658, 1032
Lieber, E., 362, 1032
Lieberman, I., 467, 1032
Lieberman, M., 120, 1032
Liener, I. E., 770, 804, 1032
Lifson, N., 228, 232, 1031, 1032
Lilly, J. H., 29, 1000
Lin, E. C. C, 263, 265, 394, 403, 1015,
1067
Lind, C. J., 292, 1067
Lindberg, O., 444, 445, 865, 872, 1033,
1054
Lindell, S. E., 363, 1032
Linderholm, H., 882, 912, 1032
Linderdt, T., 461, 464, 1004, 1008
Linderstrom-Lang, K., 646, 1032
Lindner, R. C, 979, 1032
Lindsay, A., 341, 347, 547, 549, 556,
557, 558, 560, 780, 995, 1017
Lindsten, T., 105, 213, 215, 217, 218,
1009
Lindstrom, E. S., 26, 293, 1032, 1068
Lineweaver, H., 227, 228, 667, 668,
990, 1032
Ling, G., 212, 1032
Link, G. K. K., 74, 171, 189, 1032
Linskens, H. F., 967, 1044
AUTHOR INDEX
1099
Linstrom, 0., 959, 960. 1059
Lipke, H., 675, 70-32
Lipmann, F., 326, 556, 887, 1023, 1032,
1053
Lipsett, M. N., 741, 1032
Lipten, M. A., 75, 79, 115, 1057
Lipton, M. M., 385. 1061
Lister, A. J., 603, 610, 1032
Litchfield, J. H., 53, 74, 77, 79, 1032
Lito, E., 975, 1033
Littlefield, J. W., 844, 1050
Liverman, J. L., 451, 1046
Livemiore, A. H., 518, 524, 1032, 1052
Ljunggren, M., 708, 849, 1006
Llinas, J. M., 708, 837, 1045
Lloyd, D. R., 9, 1032
Lockwoad, W. H., 888, 1032
Lodin, Z., 950, 1032
Loevenhart, A. S., 956, 1064
Loblich, H. J., 924, 1051
Loefer, J. B., 576, 1032
Loevenhart, A. S., 701, 707, 723, 724,
725, 1032
Levtrup, S., 76, 81, 152, 1033
Low, H., 444, 445, 547, 548, 549, 551,
555, 556, 864, 865, 872, 1014, 1033, 1054
Logemann, W., 686, 693, 993
London, M., 440, 441, 442, 1032
Long, W. K., 957, 1032
Longley, J. B., 926, 989
Loomis, W. D., 61, 1058
Loomis, W. F., 556, 1032
Long, J. P., 40, 1004
Long, M. v., 15, 1044
Long, W. K., 943, 1032
Loofbourovv, J. R., 657, 1015
Lorand, L., 375, 1033
Lord, C. F., 196, 1033
Lorenz, B., 461, 1033
Lorenz, R., 461, 1033
Loring, J. M., 393, 399, 1063
Lotspeich, W. D., 81, 207, 210, 709, 844,
1025, 1033, 1063
Loureiro, J. A., de, 975, 1033
Louwrier, K. P., 409, 1043
Loveless, L. R., 195, 727, 972, 973, 1033
Lovenberg, W., 310, 1033
Low, B. W., 757, 758, 1033
Lowe, H. J., 60, 341, 346, 1010
Lowenstein, J. M., 70, 72, 89, 147,
1024, 1028, 1056
Lowery, D. L., 599, 1051
LowTy, O. H., 89, 286, 287, 288, 289,
385, 407. 474. 540, 997, 1024, 1033,
1044
Lozano, R., 918, 923, 924, 929, 930, 935,
1026
Lucas, D. R., 953, 1056
Ludden, C. T., 315, 1057
Ludowieg, J., 385, 1004
Luduena, F. P.. 901. 1038
Ludwig, B. J., 297. 1033
Ludwig. G. D.. 544. 1060
Luebering, J., 817, 818, 820, 1047
Lubke, M., 891, 1002
Liick, H., 623, 1033
Liillmann, H., 937, 938. 1029
Luh. B. S., 421, 1033
Lukomskaia, I. S., 417, 1033
Lumnis. W. L.. 201, 577, 968, 1055
Lumry, R., 60, 365, 1055
Lund, P., 594, 1004
Lundbom, S., 293, 1039
Lundgren, K.-D., 959, 960, 1059
Lusty, C. J., 46, 1054
Luteraan, P. J., 195, 1033
Lutwak-Mann, C, 18, 25, 34. 35, 41, 179,
661, 662, 664, 1020, 1033
Luukkainen, T.. 349, 1027
Luzzati, M., 552, 708, 994
Luzzato, R., 726, 1033
Lwoff, A., 62, 63, 81, 82, 1033
Lyman, C. M., 666, 667, 991
Lynch, J. L., 359, 360, 1041
Lynen, F., 27, 77, 92, 187, 614, 751, 887,
987, 995, 1033
Lyon, I., 148, 150, 1033
M
Mc AUan, A., 429, 1032
Mc Calla, T. M., 974, 1033
Mc Clure, F. J., 630, 631, 1070
Mc Coll, J. D., 504, 505, 1023
Mc CoUister, D. D., 627, 1056
Mc Comb, R. B., 381, 382, 388, 395, 1033
1100
AUTHOR INDEX
Mc Cormack, B. R. S., 175, 184, 1039
Mc Cormick, D. B., 358, 475, 539, 542,
564, 565, 578, 1033
Mc Cormick, N. G., 675, 1033
Mc Cornack, B., 910, 913, 1050
Mc Crea, F. D., 944, 945, 1033
Mc Curdy, M. D., Jr., 169, 1033
Mc Daniel, E. G., 495, 1033
Mc Devitt, M., 810, 835, 870, 1001
Macdonald, K., 63, 439, 711, 778, 1034
Mc Donald, J. K., 38, 1033
Mac Donnell, L. R., 745, 754, 760, 1034
McDougall, B. M., 581, 994
Macdowall, F. D. H., 891, 1034
Mc Elroy, W. D., 845, 850, 1013, 1026
Mc Ewen, B. S., 32, 189, 393, 622, 624,
1034
Mc Fadden. B. A., 61, 892, 1034
Macfarlane, E. W. E., 964, 966, 1034
Macfarlane, W. V., 938, 1034
Mc Garrahan, J. F., 382, 1034
Mac Gillavry, C. H., 4, 1012
Mc Gilvery, R. W., 1038
Mc Gowan, J. C, 618, 632, 1034
Mc Grath, H., 351, 1046
Mc Guire, J. S., Jr., 449, 1034
Machado, A. L., 685, 1040
Macheboeuf, M., 15, 939, 999
Mc Henry, E. W., 556, 569, 573, 991, 997
Machlis, L., 27, 116, 168, 170, 185,
994, 1034
Macht, D. I., 944, 956, 957, 966, 1034
Mc Hugh, R., 440, 441, 442, 1032
Mcllwain, H., 350, 485, 491. 497, 504,
587, 637, 1034
Mackay, D., 310, 1034
Mc Kee, R. W., 396, 397, 1021
Mackenzie, C. G., 601, 1010
Mc Kinney, G. R., 434, 594, 1034
Mac Kinnon, J. A., 376, 381, 382, 383,
842, 997
Mackler, B., 549, 552, 553, 555, 783,
832, 847, 848, 849, 1020, 1034, 1035,
1044
Mc Lean, P., 708, 711, 838, 839, 1012
Mac Lean, P. D., 499, 1000
Mac Lennan, D. H., 274, 1034
Mac Leod, J., 699, 1034
MacLeod, R. M., 451, 452, 1034
Mc Mahon, P., 173, 882, 987
Mc Mahon, R. E., 592, 1034
Mc Manus, T. T., 149, 232, 887, 1039
Mac Nider, W. de B., 954, 956, 985, 1034
Mc Rae, S. C, 836, 1056
Mc Shan, W. H., 29, 30, 31, 1000, 1034
Madden, T. J., 260, 489, 504, 1068
Madinaveitia, J., 538, 546, 1034
Madoff, M., 495, 992
Madonska, L., 206, 988
Madsen, N. B., 19, 26, 547, 555, 648,
649, 789, 803, 808, 811, 813, 853, 862,
1034
Maengwyn-Davies, G. D., 444, 1034
Magee, P. N., 225, 990
Magel, T. T., 619, 997
Mager, J. ,848, 878,989
Maggiolo, I. W., 778, 1010
Magin, J., 942, 945, 1057
Mahadevan, S., 547, 554, 1035
Mahler, H. R., 16, 284, 511, 549, 555
707, 710, 713, 781, 832, 849, 977, 981
1034, 1035, 1054, 1063, 1064
Mahowald, T. A., 869, 1007
Maio, J. J., 394, 1035
Maitre, L., 318, 1035, 1040
Maitra, P. K., 53, 228, 395, 838, 839,
1035
Maizels, M., 177, 209, 928, 1035
Makino, K., 578, 1035
Malachowski, R., 619, 1035
Maley, F., 381, 382, 469, 474, 781, 837,
1034, 1035
Maley, G. F., 469, 781, 837, 1035
Malhotra, 0. P., 810, 1064
Malkin, A., 485, 489, 492, 1035
Mallus, E., 968, 1035
Malmstrom, B. G., 789, 810, 837, 1035
Malvin, R. L., 205, 924, 1035, 1062
Manaker, R. A., 978, 1015
Manchol, P., 268, 340, 1048
Mancilla, R., 406, 407, 839, 1035
Mandel, H. G., 478, 1016
Mandelstam, J., 155, 156, 1035
Mandelstam, P., 379, 380, 1008
Manery, J. F., 121, 1036
Mangelsdorf, P. C, 64, 1026
AUTHOR INDEX
1101
Mann, F. C, 914, 1035
Mann, F. D., 914, 1035
Mann, K. M., 377, 1030
Mann, P. J. G., 485, 1035
Mannering, G. I., 597, 1059
Manners, D. J., 429, 843, 1015
Manning, D. T., 271, 1035
Manning, G. B., 269, 833, 834, 1035
Mano, Y., 518, 522, 523, 854, 1035, 1065
Manoukian, E., 129, 1007
Mansour, J. E., 474, 1035
Mapson, L. W., 542, 547, 550, 662, 713.
773, 774, 778, 781, 858, 859, 1035,
1058
Marchenko, N. K., 194, 1050
Marcus, G. J., 121, 1036
Marcus, P. I., 449, 1036, 1059
Mardashev, S. R., 307, 1036
Margulies, S. I., 856, 1040
Marinetti, G. V., 151, 1036
Marini, M., 458, 1036
Maritz, A., 338, 348, 1070
Mark, H. J., 763, 1051
Marks, P. A., 166, 497, 1018, 1036
Markwardt, F., 675, 707, 836, 1036
Marre, E., 623, 850, 1036
Marsh, B. B., 704, 705, 721, 722, 865,
876, 989
Marsh, C. A., 272, 426, 427, 428, 429,
1008, 1031, 1032, 1036
Marsh, J. B., 178, 185, 1017
Marsh, M. M., 759, 1025
Marshall, J. K., 264, 1031
Marshall, M., 781, 804, 1036
Marshall, P. B., 560, 1036
Marsland, D., 964, 965, 1029, 1070
Martell, A. E., 13, 658, 739, 998, 1031,
1036
Martin, A. R., 224, 236, 1003
Martin, D. B., 234, 1020
Martin, G. J., 259, 261, 586. 587, 1036
Martin, L. E., 63, 1012
Martin, R. B., 374, 988
Martin, R. G., 351, 861, 1036
Martin, S. M., 27, 677, 1036
Martin, S. P., 434, 1031
Martonosi, A., 868, 939, 940, 1011, 1036
Marui, E., 985, 1061
Maruyama, K., 866, 939, 1011, 1042
Maruyama, M., 892, 1022
Maschmann, E., 685, 793, 1036
Mason, H. L., 670, 1036
Mason, J., 972, 1001
Mason, M., 64, 595, 600, 607, 608, 609,
1036, 1068
Mason, ,S. F., 280, 1036
Masoro, E. J., 613, 1008
Massart, L., 693. 1036
Massen, V.. 16, 18, 48, 54, 60, 65, 79,
80, 94. 174, 183, 243, 273, 275, 276,
277, 774, 787, 788, 805, 856, 1036,
1043, 1054
Massini, P., 892, 1036
Master, R.W. P., 15,987
Masuda, T., 535, 1036
Masuoka, D. T., 191, 215, 1036
Matheson, N. A., 593, 1025
Mathews, A. P., 963, 1036
Mathews, C. K., 479, 1036
Matkovics, B., 503, 1037
Matsuda, R., 618, 632, 1062
Matsumoto, H., 33, 244, 1021
Matsumura, Y., 78, 149, 179, 228, 1070
Matsuo, Y., 709, 843, 1037
Matsuoka, M., 611, 612, 1042
Matsushima, T., 416, 417, 1025
Matteucci, W. V., 626, 1051
Matthes, K. J., 147, 273, 987
Matthews, J., 387, 991
Matthew, M., 238, 1037
Matthews, L. W., 114, 141, 1011
Matthews, R. E. F., 194, 261, 1037
Matthews, R. J., 358, 1002
Matthies, H., 946, 1045
Maung, K., 429, 843, 1015
Mauzerall, D., 859, 888, 1037
Maver, M. E., 688, 707, 1037
Mavrides, C, 612, 1004
Maw, G. A., 356, 1037
Maxwell, E. S., 858, 859, 1037
May, C. E., 383, 991
Maybury, R. H., 758, 759, 761, 1005
Mayeda, M., 385, 1004
Mayer, A. M., 847, 1037
Mayer, R. L., 297, 304, 458, 459, 461, 462,
1018, 1028
1102
AUTHOR INDEX
Mazelis, M., 151, 466, 1037
Mazia, D., 969, 1037
Mazur, A., 711, 1037
Mazurova, T. A., 224, 225, 1056
Meares, J. D., 938, 1034
Mechler, F., 949, 1016
Medina, A., 547, 553, 554, 782, 842,
1018, 1037, 1041
Medina, H., 550, 1001
Meek, W. J., 944, 945, 1033
Mehl, J. W., 691, 692, 1037
Meinke, M., 145, 1002
Meier, R., 875, 880, 883, 884, 968, 1037
Meisel, E., 926, 927, 1064
Meiss, A. N., 831, 988
Meister, A., A., 437, 1037
Mello Ayres, G. C, 778, 781, 1037
Melnick, I., 333, 1031
Melson, G. L., 359, 1020
Melville, K. I., 952, 1009
Mendel, B., 673, 677, 1037
Mendelsohn, M. L., 928, 1037
Mendez, R., 669, 723, 724, 942, 944,
1037
Mendicino, J., 470, 1037
Menegas, R., 429, 1013
Meneghetti, E., 902, 1037
Menke, K. H., 875, 1051
Menon, G. K. K., 228, 233, 234, 1037
Menon, V. K. N., 64, 1040
Mercker, H., 937, 1029
Meridith, C. H., 359, 1020
Meriwether, B. P., 409, 1060
Meroney, W. H., 925, 1065
Merrett, M. J., 237, 1037
Merrifield, R. B., 533, 1068
Merritt, P., 221, 222, 993
Mescon, H., 767, 1037
Messina, R. A., 770, 1037
Metzenberg, R. L., 781, 804, 838, 1036,
1037
Metzger, R. P., 410, 839, 1037
Meyer, D. K., 909, 1037
Meyer, H., 137, 868, 969, 1036, 1037
Meyer, J. R., 966, 1037
Meyer, R. K., 30, 31, 1034
Meyer, V., 701, 1037
Meyerhof, O., 292, 1037
Michaelis, L., 259, 271, 416, 421, 1037
Michel, O., 678, 689, 1047
Michel, R., 678, 689, 1047
Michelson, C, 61, 1058
Mickelson, M. N., 77, 87, 104, 168, 1038
Middlebrook, M., 689, 1038
Mil, S., 463, 1038
Mikulski, P., 988
Mildvan, A. S,, 789, 793, 816, 819,
1014, 1038
Milholland, R. J., 571, 574, 1049
Millar, D. B. S., 787, 802, 1038, 1067
Millbank, J. W., 51, 1038
Miller, A., 783, 1038
Miller, J. J., 195, 1038
Miller, D. N., 75, 1038
Miller, T. B., 917, 934, 935, 1007, 1038
MiUer, V. L., 957, 959, 961, 1038
Millerd, A., 18, 19, 27, 53, 77, 78, 79,
80, 82, 120, 172, 173, 189, 855, 994,
995, 1017, 1038
Millington, P. F., 950, 1038
Millington, R. H., 137, 141, 273, 274,
1065
Mills, G. T., 61, 427, 1038
Mills, J., 592, 1034
Mills, J. M., \\,2Z6,1044
Mills, R. C, 18, 187, 543, 555, 840, 1024
1047, 1064
Mills, R. R., 332, 1038
Milofsky, E., 435, 1043
Milstein, C, 662, 705, 706, 711, 716, 717,
780, 804, 810, 832, 1038, 1058
Minakami, S., 163, 510, 798, 872, 888,
1024, 1038, 1069
Minatoya, H., 901, 1038
Mirsky, A. E., 32, 189, 393, 622, 624,
661, 670, 671, 672, 690, 988, 1034,
1038
Mirsky, I. A,, 883, 884, 1015
Misaka, E., 817, 1038
Misra, U. K., 421, 1038
Mitchell, H. K., 53, 1017
Mitchell, I. L. S., 384, 1023, 1038
Mitchell, J. H., Jr., 585, 1055
Mitchell, M. B., 53, 1017
MitcheU, P., 267, 690, 910, 912, 1038
MitcheU, R. A., 772, 782, 1038
AUTHOR INDEX
1103
Mitchell, R. B., 221, 222, 223, 993
Mitidieri, E., 289, 1063
Mittermayer, C, 708, 1038
Mitz, M. A., 367, 1069
Miura, N., 985, 1038
Miyachi, S., 892, 1038
Mize, C. E., 841, 1038
Mizushima, S., 349, 1038
Mochizuki, Y., 156, 1042
Modell, W., 956, 957, 1038
Modignani, R. L., 394, 1038
Mollering, H., 841, 993
Moetsch, J. C, 957, 1046
Mokrasch, L. C, 1038
Molinari, R., 435, 511, 547, 551, 557, 558,
709, 710, 717, 773, 843, 844, 849,
1035, 1038, 1048
Molnar, D. M., 291, 292, 294, 1038
Mommaerts, W. F. H. M., 445, 939, 940,
1014, 1025
Momose, G., 2, 138, 1038
Mondy, N. I., 585, 1058
Moniz, R., 519, 1005
Monk, C. B., 11, 13, 1007
Monroy, A., 726, 1038
Monson, W. J., 36, 37, 41, 240, 288, 1004
Montgomery, C. M., 31, 35, 36, 46, 47.
48, 51, 69, 70, 71, 75, 76, 81, 82, 83,
88, 198, 240, 241, 1039, 1038
Montgomery, R., 15, 996
Mook, W., 943, 944, 945, 1007
Mookerjea, S., 138, 144, 149, 218, 1039
Moore, B. W., 901, 902, 905, 1039
Moore, H., 901, 905, 1024
Moos, C, 28, 446, 1023, 1039
Moosburger, A., 750, 881, 884, 885, 1006
Mora, P. T., 459, 462, 1039
Morales, M. F., 446, 447, 1041, 1042
Morawetz. H., 457, 1039
Morgan, E. J., 18, 25, 34, 35, 41, 661,
662, 664, 1020, 1039
Morgan, H. E., 125, 263, 1039, 1045
Morgan, H. R., 194, 1039
Morhara, K., 660, 1039
Mori, T., 292, 293, 1018
Morino, Y., 564, 1039
Morisue, T., 564, 578, 676, 858, 1039,
1064
Morita, T. N., 387, 1066
Moriyama, H., 977, 980, 1039
Morren, L., 747, 763, 1027
Morris, M. P., 244, 1039
Morrison, J. F., 707, 833, 1039
Morrison, J. H., 197, 1067
Morrison, J. F,, 171, 185, 189, 467, 1014,
1039
Morrison, J. R., 359, 1020
Morrow, P. F., W., 444, 1065
Morse, P. A., Jr., 481, 1022
Morton, H. E., 349, 1039
Morton, R. K., 20, 27, 63, 471, 481,
485, 493, 510, 686, 787, 989, 995,
1039, 1067
Mosbach, E. H., 15, 1044
Moser, J. C, 780, 798, 865, 1050
Moses, F. E., 385, 1004
Moses, v., 51, 53, 187, 236, 1039
Motoyama, H., 969, 1025
Mott, J. C, 962, 1046
Moudgal, N. R., 681, 1046
Moudler, J. W., 53, 73, 74, 78, 91, 93,
124, 125, 194, 360, 1039, 1056
Mounter, L. A., 68, 662, 675, 684, 685,
836, 1039
Moussatche, H., 176, 212, 213, 1039
Moyed, H. S., 321, 481, 854, 1039, 1062
Moyer, I. H., 933, 1039
Mozen, M. M., 293, 1039
Mozingo, R., 538, 1019
Mudd, J. B., 148, 149, 226, 232, 887,
1039
Mudd, S., 956, 972, 1055
Mudge, G. H., 78, 137, 179, 205, 623,
626, 883, 917, 924, 928, 931, 932, 933,
934, 935, 936, 1032, 1039, 1040, 1065
Miiller, A. F., 158, 1040
Miiller, E., 664, 1040
Miiller, F., 941, 951, 956, 957, 1040
Miiller, G., 168, 228, 237, 504, 1040
Miiller, J., 950, 1032
Mueller, J. F., 574, 577, 1040
Miiller, O. H., 929, 1065
Mueller, R. T., 852, 1064
Muller-Rodloff, I., 664, 1040
Muenst«r, A. M., 23, 33, 46, 47, 1020
Muir, C, 133, 1040
1104
AUTHOR INDEX
Mullnos, M. G., 724, 993
Mullins, L. J., 905, 1040
Munier, R., 15, 999
Murachi, J., 340, 810, 1040
Murayama, M., 755, 1040
Murphy, B., 946, 1027
Murphy, J. V., 209, 909, 1063
Murray, A. W., 471, 481, 989
Murray, D. R. P., 259, 1040
Murthy, S. K., 547, 554, 1035
Muscatello, U., 865, 989
MuschoU, E., 318, 938, 1040
Mushett, C. W., 577, 1040
Mustakallio, K. K., 925, 1040, 1060
Muus, J., 813, 1040
Myant, N. N., 149, 1022
Mycek. M. J., 375, 887, 1000, 1010
Myerhof, O., 815, 866, 1045
Myers, C. D., 974, 1000
Myers, D. K., 60, 126, 816, 820, 867, 869,
988, 1040
Myers, T. C, 446, 474, 1039. 1054
Myrback, K., 464, 660, 675, 683, 685,
688, 691, 781, 791, 792, 838, 1040
N
Naber, E. C, 530, 1040
Nachlas, M. M., 856, 1040
Nachmansohn, D., 683, 685, 1040
Nadai, Y., 338, 1040
Nadeau, L. V., 964, 966, 1034
Nadkarni, S. R., 833, 1040
Nadler, J. E., 944, 1050
Nagai, S., 26, 36, 62, 63, 855, 1029, 1069
Naganna, B., 64, 1040
Nagasawa, M., 662, 687, 1069
Nagata, Y., 153, 1061
Nagatsu, I., 544, 1068
Nagatsu, T., 320, 1062
Nagler, H., 947, 1050
Naik, K. G., 240, 1040
Naite, B., 744, 1010
Nair, P. M., 547, 549, 1040
Nair, P. V., 437, 997
Najjar, V. A., 383, 1010
Nakada, H. I., 78, 152, 178, 187, 218,
219, 228, 232, 233, 234, 387, 390, 392,
398, 399, 404, 407, 857, 1040, 1052,
1066, 1067
Nakahara, T., 546, 988
Nakamura, H., 847, 1040
Nakamura, K., 225, 856, 1040
Nakamura, M., 61, 196, 1040
Nakamura, M., 61, 196, 1040
Nakanishi, K., 817, 1038
Nakao, K., 163, 1069
Nakao, M., 817, 1017
Nakata, H. M., 236, 237, 1040
Nakayama, T., 832, 898, 1041
Nanninga, L. B., 445, 1041
Naoi, M., 537, 1005
Naoi-Tada, M., 817, 1041
Naono, S., 479, 1014
Narrod, S. A., 547, 550, 1041
Narurkav, M. V., 415, 1050
Naschke, M. D., 71, 1052
Nason, A., 487, 490, 493, 551, 552, 553,
843, 849, 851, 994, 1024, 1031, 1049,
1068
Nasu, H., 578, 676, 858, 1064
Nath, R., 582, 1041
Nathan, H., 283, 1006
Nathans, D., 207, 265, 1041
Natsume, K., 340, 1068
Naurath, H., 365, 1006
Navazio, F., 518, 1041
Neame, K. D., 266, 1041
Needham, D. M., 866, 869, 1014
Negelein, E., 875, 1051
Neidle, A., 887, 1000
Neilands, J. B., 437, 686, 710, 802, 825,
845, 1041, 1067
Neims, A. H., 340, 1041
Neish, A. C, 599, 1011
Nelson, C. A., 475, 1041
Nelson, E. K., 2, 225, 1041
Nelson, J. M., 297, 1014, 1033
Nelson, L., 722, 864, 882, 991
Nelson, M., 859, 1041
Nelson, M. M., 576, 1041
Nelson, W. L., 837, 1049
Nerurkar, M. K., 415, 1050
Neuberger, A., 238, 751, 888, 1012, 1037
Neufeld, E. F., 507, 508, 409, 511, 512,
842, 1007, 1041
AUTHOR INDEX
1105
Neuhaus, F. C, 270, 359, 360, 853, 1041
Neuhaus, H., 981, 1041
NeuhofF, v., 499, 1018, 1067
Neuman, W. F., 54, 58, 166, 175, 995
Neumann, H., 457, 1024
Neurath, H., 369, 370, 373, 735, 797,
798, 810, 1014, 1025, 1040, 1041
Newburgh, R. W., 132, 1023
Newcomb, E. H., 136, 143, 1021, 1067
Newey, H., 265, 1041
Newhouse, J. P., 953, 1056
Newman, M. D., 950, 1017
Newmark, M. Z., 773, 817, 1041
Newton, G. G. F., 599, 987
Neyman, M. A., 540, 544, 993
Ngai, S. H., 532, 1041
Nichol, C. A., 571, 574, 582, 583, 1041,
1049, 1069
Nicholas, D. J. D., 547, 553, 554, 782,
787, 842, 1008, 1037, 1041, 1064
Nickerson, M., 217, 1041
Nickerson, W. J., 169, 880, 1041
Niderland, J. R., 954, 957, 1005
Niedergang-Kamien, E., 881, 967, 1041
Niederpruem, D. J., 881, 1041
Nielsen, H., 157, 1007
Niemann, C, 271, 349, 372, 373, 374,
988, 1009, 1020, 1029, 1035, 1064
Niemer, H., 291, 1041
Nigam, V. N., 63, 442, 1041
Nihei, J., 446. 447, 1041, 1042
Xilsson, K., 363, 1032
Nilsson, M., 619, 1009
Nimmo-Smith, R. H., 772, 1041
Ninomiya, H., 882, 1041
Nirenberg, M. W., 263, 389, 390, 391,
1041
Nishi, A., 368, 842, 1041, 1042
Xishida, J., 439, 1062
Nishimura, J. S., 357, 787, 1042
Nishimura, S., 463, 1042
Nishiyama, T., 156, 1042
Nistratova, S. N., 947, 1042
Niwa, T., 124, 135, 176, 184, 1026
Nocito, v., 60, 338, 348, 994
Noda, H., 939, 1042
Noda, L., 60, 446, 447, 836, 847, 1028,
1041, 1042
Noe, F. E., 952, 954, 1042
Noguchi, H., 983, 1042
Nogueira, O. C, 510, 552, 850, 1047
Nohara, H., 816, 1042
Noltmann, E., 406, 855, 996, 1042
Nomaguchi, G. M., 217, 1041
Nomura, M., 51, 74, 77, 1059
Nordlie, R. C, 357, 475, 1010, 1042
Norris, E. R., 289, 1028
Norris, J. L., 864, 1029
Norris, L. C, 530, 1002
Norton, G., 91, 105, 106, 111, 132, 149,
154, 172, 182, 1048
Nosoh, Y., 768, 1042
Nossal, P. M., 15, 62, 1029, 1042
Nour El Dein, M. S., 91, 678, 1053
Novikoff, A. B., 798, 865, 1042
Novinger, G., 865, 912, 1030
Novoa, W. B., 433, 435, 436, 1042
Novosel, D. L., 360, 1039
Nozzolillo, C. G., 511, 553, 847, 1019
Nuenke, B. J., 681, 745, 750, 804, 812,
1002, 1062
Xiirnberger, H., 803, 1052
Nukada, T., 611, 612, 1042
Nutting, M.-D. F., 657, 1023
Nygaard, A. P., 62, 435, 437, 683, 1042
Nyhan, W. L., 152, 156, 1042
Nyteh, P. D., 709, 718, 997
Gates, J. A., 315, 1042
Ochoa, S., 62, 63, 64, 75, 83, 145, 175,
181, 187, 224, 226, 234, 235, 463, 469,
474, 597, 705, 830, 853, 990, 1008,
1025, 1035, 1038, 1042, 1049, 1050
O'Connor, R. J., 270, 354, 674, 705, 831,
1042
Odeblad, E., 959, 1009
Oesper, P., 711, 1047
Officer, J. E., 360, 1039
Ogata, K., 156, 816, 1042
Ogata, M., 156, 1042
Ogilvie, R. F., 954, 1042
Oginsky, E. L., 353, 1042
Ogiu, K., 985, 1061
Ogston, D., 560, 995
1106
AUTHOR INDEX
Ohashi, S., 977, 980, 1039
Ohnishi, T., 868, 1061
Ohno, M., 328, 1042
Ohr, E. A., 907, 1042
Ohshima, S., 428, 987
Ohta, J., 173, 1042
Okada, K., 347, 1069
O'Kane, D. J., 848, 854, 855, 1007, 1039
Okazaki, R., 470, 1042
Okuda, J., 347, 1069
Okui, S., 817, 1027
Okunuki, K., 26, 27, 228, 229. 328,
1020, 1022, 1042
Olavarria, J. M., 476, 1031
Oleson, J. J., 504, 505, 1016
Oliphant, J. F., 982, 1042
Olivard, J., 575, 1042
Oliver, I. T., 603, 1042
Olemucki, A., 779, 780, 1063
Olsen, N. S., 350, 1027
Olson, E. J., 714, 1042
Olson, J. A., 708, 709, 826, 840, 863,
1042, 1043
Olson, J. M., 891, 1043
Olson, M. E., 336, 351, 1046
Olson, R. E., 75, 103H
Omote, Y., 781, 827, 1001
Onrust, H., 519, 774, 775, 854, 1043
Ordal, E. J., 675, 1033
Ordal, Z. J., 53, 74, 77, 79, 1032
Ordin, L., 170, 209, 273, 274, 1043
Oren, R., 435, 1043
OrlofF, J., 917, 1043
Ornstein, N., 199, 887, 964, 965, 1014,
1043
Orten, J. M., 89, 91, 104, 109, 1002,
1010, 1043
Ortiz, P. J., 64, 145, 224, 226, 234, 235,
1008
Osborn, M. J., 581, 585, 1043, 1066
Ota, S., 64, 792, 813, 1043
Otsuka, S., 554, 1043
Otsuka, S.-I., 880, 1053
Ott, J. L., 692, 1043
Ott, P., 768, 1028
Ott, W. H., 562, 1043
Ottolenghi, P., 55, 65, 179, 436, 485,
1043, 1049
Ouellet, L., 711, 772, 773, 1030
Overall, B. T., 15, 1043
Overgaard-Hansen, K., 267, 1030
Owens, H. S., 15, 1043, 1057
Owens, R. G., 797, 1043
Oxender, D. L., 266, 1043
Ozaki, K., 706, 709, 841, 1043
Ozaki, M., 315, 316, 317, 318, 319, 320,
611, 1012, 1018, 1062
Ozawa, G., 573, 991
Ozawa, T., 347, 772, 1069
Paasonen, M. K., 611, 7022
Packer, L., 31, 154, 177, 1043, 1050
Padilla, A. M., 540, 997
Padykula, M. A., 927, 947, 1073
PafF, G. H., 215, 462, 995, 1043
Pagan, C., 244, 1039
Pahl, J. I., 228, 232, 1045
Paigen, K., 125, 1065
Paige, M. F. C, 224, 236, 1003
Paine, C. M., 912, 1009
Palm, D., 359, 577, 1020
Palmer, G., 772, 774, 805, 1043
Palmer, J. F., 918, 920, 997
Palmer, J. K., 91, 105, 106, 172, 190, 225,
228, 1063
Pan, S. F., 90, 1069
Pan, Y. L., 984, 994
Panagos, S. S., 613, 1008
Pantlitschko, M., 459, 695, 1043
Papa, M. J., 177, 1043
Papacenstantinou, J., 433, 434, 435, 1043
Pappas, A., 158, 1047
Paquette, L. A., 358, 1052
Pardee, A. B., 36, 38, 63, 67, 68, 73, 75,
76, 82, 88, 105, 117, 177, 178, 326, 357,
468, 480, 481, 816, 1011, 1043, 1068
Park, C. R., 125, 263, 1039, 1045
Park, J. H., 409, 714, 1042, 1060
Park, R. B., 409, 1043
Parke, D. V., 630, 990
Parker, A. J., 639, 1043
Parker, C. A., 292, 293, 294, 1043, 1044
Parks, R. E., Jr., 383, 478, 1000, 1002,
1029, 1044
AUTHOR INDEX
1107
Parmar, S. S., 38, 40, 42, 240, 1018
Parpart, A. K., 908, 1014
Parr, C. W.. 272, 405, 406, 998, 1044
Partowi, R., 922, 1011
Partridge, A. D., 973, 983, 985, 1044
Passey, R. D., 225, 990
Passonneau, J. V., 385. 407, 474, 1024,
1044
Passow, H., 187, 188, 898, 899, 908.
1012, 1044
Pasternak, C. A. 472, 478, 1016, 1044
Patel, C, 464, 1021
Patel, R. P., 240, 1040
Pattabiraman, T. N., 816, 1044
Patterson, M. K., Jr., 804, 842, 1062
Patterson, P. A., 576, 1024
Patterson, W. B., 208, 1044
Paul, J., 61, 428, 1038
Paul, M. H., 71, 140, 995. 1010
Pauling, L., 43, 1044
Payes, B., 616, 1044
Pazur, J. H., 389, 1044
Pearse, A. G. E., 850, 1018
Pearson, A. M., 603, 1046
Pearson, D. E., 189, 1044
Pearson, J. A., 53, 77, 79, 80, 82, 173, 1017
Pearson, R. G., 11, 236, 1044
Pease, D. C, 198, 678, 1044
Pece, G., 623, 1036
Pechstein, H., 259, 271, 421, 1037
Peck, H. D., Jr., 49, 79, 1000, 1011, 1044
Pedersen, T. A., 231, 1044
Peel, E. W., 538, 1019
Pelczar, M. J., Jr., 79, 1060
Pelizza, G., 615, 1049
Peluffo, C. A., 203, 727, 1003
Pendergast, J., 143, 1011
Penefsky, H. S., 548, 556, 705, 865, 869,
873, 1044, 1046
Penefsky, Z. J., 713, 1061
Pengra, R. M., 292, 1012
Penn, N., 552, 849, 1044
Pennington, R. J., 548, 556, 613, 1044
Pentschew, A., 952, 1044
Peralta, B., 669, 723, 942, 1037
Pereira, A. S. R., 967, 1044
Perez, J. E., 978, 979, 980, 1027, 1044
Perisutti, G., 870, 1001
Perkins, D. J., 737, 739, 748, 753. 1044
Perkins, M. E., 658, 676, 683, 686, 1017
Perlmann, G. E., 657, 1012
Perova, K. Z., 983, 984, 985, 1022
Perri, V., 520, 526, 527, 528, 532, 578,
1003, 1048
Perry, J. J., 78, 83, 1044
Perry, S. V., 684, 691, 692, 723. 816.
938, 939, 989, 998
Pershin, G. N., 774, 976, 1044
Person, P., 19, 29, 1044
Persson, B., 464, 1040
Perutz, M. F., 755, 1014
Peskoe, L. Y., 574, 577, 1051
Petering, H. G., 287, 289, 538, 851, 1030,
1044
Peters, E. L.. 834, 1061
VeteTS,J.M., 582, 1044
Peters, R. A., 36, 63, 75, 125, 181, 187,
709, 844, 980, 1032, 1033, 1044
Peters-Mayr, T., 368, 1013
Peterson, E. A., 156, 1044
Petrucci, D., 174, 1044
Pette, D., 1020
Pfleiderer, G., 802, 810, 814, 857, 861,
1014, 1015, 1044
Phaff, H. J., 421, 1033
Phares, E. F., 15, 1003, 1044
Philips, F. S., 698, 699, 700, 1012, 1045
Philipsen, L., 976, 977, 979, 980, 981, 999
Phillips, A. H., 815, 817, 1045
Phillips, P. H., 77, 87, 124, 128, 176, 203,
1030
Philpot, J. St. L., 658, 1045
Phizackerley, P. J. R., 845, 1009
Pierpont, W. S., 28, 79, 80, 82, 87, 1045
Pietschmann, H., 1045
Pihl, A., 639, 662, 663, 691, 697, 699,
750, 751, 804, 1006, 1045, 1050
Piloty, O., 664, 1045
Pincus, G., 179, 183, 1009
Pine, M. J., 911, 1045
Piquet, J., 835, 1010
Fine, A., 596, 846, 1062
Pirie, N. W., 693, 712, 1019
Pisanty, J., 723, 724, 1037, 1045
Pitot, H. C, 593, 1028
Pitts, R. F., 917, 918, 919, 923, 924,
1108
AUTHOR INDEX
928, 929, 930, 934, 935, 960, 995,
1014, 1026, 1045
Pizer, L. I. 852, 1001
Plant, G. W. E., 63, 509, 513, 539, 543,
696, 705, 782, 844, 852, 872, 874,
999, 1045, 1054, 1067
Plass, M., 61, 987
Piatt, M. E., 689, 1051
Piatt, M. H., 359, 1020
Pletscher, A., 318, 954, 1045, 1060
Plummer, D. T., 433, 1045
Podber, E., 798, 865, 1042
Poertzel, H., 750, 927, 1031
Pogell, B. M., 564, 1045
Pogrund, R. S., 314, 1045
Pohle, W., 946, 1045
Pointer, N. S., 956, 988
Polatnick, J., 176, 181, 194, 1045
Polglase, W. J., W, 365, 1055
Polimeros, D., 920, 921, 1013
Polis, B. D., 815, 866, 1045
Pollock, M. R., 697, 699, 1045
Polonovski, M., 308, 1045
Polynovskii, O. L., 675, 857, 1045
Pon, N. G., 409, 1043
Pontremoli, S., 407, 412, 995, 1013
Ponz, F., 708, 837, 1045
Pope, H., 228, 993
Popinigis, J., 988
Popjak, G., 146, 147, 711, 852, 886, 887,
1004, 1013, 1018, 1045
Porter, C. C, 315, 316, 317, 318, 562,
572, 664, 1045, 1057
Porter, J. W., 234, 886, 988, 1057
Porter, W. L., 15, 996
Portzehl, H., 938, 1045, 1065
Post, R. L., 1045
Postgate, J. R., 699, 1045
Potter, V. R., 3, 15, 22, 29, 30, 31, 35,
36, 38, 41, 42, 63, 67, 68, 73, 75, 76,
82, 88, 91, 98, 100, 101, 102, 103, 104,
105, 112, 117, 126, 128, 138, 139, 175,
177, 178, 201, 217, 218, 228, 232,
309, 473, 478, 479, 481, 659, 676, 687,
784, 856, 987, 994, 997, 1022, 1043,
1045, 1046, 1047, 1058
PoweU, G. W., 444, 1004
Prado, J. L., 272, 273, 274, 1050
Prairie, R. L., 121, 1000
Pratt, E. A., 462, 1031
Preiss, B., 137, 1037
Pressman, D., 683, 783, 845, 1005
Prestidge, L. S., 326, 1043
Price, C. A., 61, 84, 117, 1046
Price, J. M., 1003
Prince, R. H., 9, 1032
Pringle, A., 589, 1068
Proctor, C. H., 194, 1037
Prouvost-Danon, A., 176, 212, 213, 1039
Psychoyos, S., 387, 1062
Pubols, M. H., 854, 1046
Putter, J., 695, 1046
Pugh, D., 419, 1046
Pujarniwcle, S., 225, 226, 1009
Pullman, M. E., 548, 556, 705, 865, 869,
873, 1044, 1046
Pupilli, G., 228, 1046
Purpura, D. P., 574, 1046
Pursiano, T. A., 623, 1013
Purvis, S. E., 589, 989
Putnam, F. W., 635, 1046
Pyefinch, K. A., 962, 1046
Quaestel, J. H., 2, 18, 20, 21, 26, 30, 31,
32, 34, 35, 36, 40, 52, 55, 62, 76, 78,
87, 113, 115, 135, 138, 144, 151, 152,
153, 156, 176, 177, 184, 207, 237, 238,
261, 305, 349, 381, 382, 432, 485,
574, 589, 593, 613, 699, 991, 995,
1002, 1009, 1013, 1016, 1019, 1023,
1026, 1027, 1035, 1046, 1048, 1069
Quesnel, V. C. J., 529, 1046
Quinn, J. R., 603, 1046
Raaflaub, J., 210, 1046
Raaschou, F., 918, 923, 996
Rabin, B. R., 787, 996
Rabin, R. S., 595, 1046
Rabinovitch, M., 461, 712, 1046
Rabinovitz, M., 336, 351, 1046
Rabinowitch, E., 163, 1005
AUTHOR INDEX
1109
Rabinowitz, J. C, 575, 1046
Racker, E., 121, 408, 409, 519; 548,
556, 705, 837, 855, 865, 869, 873,
1000, 1002, 1044, 1046, 104S, 1059
Rafter, G. W., 439, 714, 783, 1046
Raghupathy, E., 681, 1046
Ragland, J. B., 451, 1046
Rahatekar, H. I., 673, 674, 830, 1046
Rahman, M. A., 81, 176, 183, 997
Rahn, O., 1046
Raiziss, G. W., 957, 1046
Rajagopalan, K. V., 549, 711, 772, 803,
1046, 1058
Rail, J. E., 678, 689, 1047
Ram, D., 383, 396, 1047
Ramachandran, S., 125, 1047
Ramakrishnan, C. V., 677, 830, 1047,
1052
Ramasarma, T., 839, 1047
Ramel, A., 807, 990
Ramsdell, P. A., 657, 701, 1017
Rand, M. J., 318, 1003
Randa, V., 391, 1057
Randle, P. J., 264, 376, 911, 991, 1001
Ranzi, S., 726, 1047
Rao, D. R., 711, 1047
Rao, M. R. R., 274, 673, 674, 830, 988,
1046, 1047
Rao, N. A., 660, 685, 854, 1012, 1047
Rao, S., 389, 1064
Rapkine, L., 661, 662, 683, 687, 1047
Rapoport, S., 489, 493, 817, 818, 820,
1019, 1047, 1051
Rapp, G. W., 406, 1047
Rapport, D., 613, 1008
Raska, S. B., 513, 1047
Rassaert, C. L., 333, 1012
Rassweiler, C. F., 618, 1047
Rathbun, R. C, 626, 1047
Ratner, S., 60, 116, 158, 338, 348, 994,
1047
Rauschke, J., 18, 991
Raval, D. N., 603, 610, 1053
Raw, I., 510, 511, 552, 710, 849, 850,
1035, 1047
Ray, T., 928, 997
Razzel, W. E., 473, 1047
Read, C. P., 28, 173, 228, 1047
Recknagel, R. 0., 138, 139, 1047
Redetzki, H. M., 505, 1047
Redfearn, E. R., 16, 1047
Redfern, S., 659, 660, 662, 674, 683,
684, 792, 833, 1003
Rees, K. R., 80, 81, 111, 114,174, 175,
999, 1047
Rege, D. V., 478, 993
Regen, D. M., 125, 1039
Reichard, P., 707, 711, 779, 780, 783,
851, 1047
Reichel, G., 314, 1003
Reichert, E., 751, 1033
Reichmann, M. E., 980, 1047
Reif, A. E., 100, 1046
Reinafarje, B., 126, 128, 987
Reinbothe, I. H., 15, 1047
Reiner, L., 546, 1047
Reinhardt, F., 1045
Reisberg, R. B., 707, 813, 835, 1047
Reiss, 0. K., 38, 40, 42, 76, 240, 702, 816,
878, 1017, 1018, 1047
Rem, L. T., 552, 710, 1005
Remberger, U., 836, 1005
Remington, M., 177, 209, 928, 1035
Remy, C. N., 966, 471, 1047
Remy, E., 974, 1015
Rendina, G., 840, 1047
Rene, R. M., 32, 204, 205, 626, 1053
Rennels, E. G., 925, 1047
Rennick, B. R., 204, 205, 626, 921, 923,
1007, 1065
Renson, J., 611, 989
Repaske, R., 18, 26, 292, 293, 547, 553,
847, 1047
Repasky, W., 574, 1040
Resch, H., 611, 1031
Resnick, H., 650, 1047
Resnik, R. A., 833, 1067
Revel, H. R., 865, 1048
Reynard, A., 480, 1049
Reznikoff, P., 982, 1048
Rhoads, C. P. 261, 1048
Rhoads, W. A., 225, 1048
Rice, B., 120, 873, 878, 1015
Rice, L. I., 13, 75, 131, 216, 228, 1048
Rice, M. S., 886, 988
Rich, A. E., 973, 983, 985, 1044
1110
AUTHOR INDEX
Richards, 0. C, 407, 1048
Richert, D. A., 287, 614, 783, 814, 859,
1004, 1018, 1066
Richter, D., 259, 296, 1048
Rickenberg, H. V., 394, 1035
Ricketts, C. R., 388, 389, 391, 996, 998
Ridgway, L. P., 576, 1024
Rieken, E., 219, 220, 991
Rieser, P., 267, 1048
Rigbi, M., 457, 1048
Riggs, A., 756, 757, 1048, 1067
Riggs, A. F., 756, 1048
Riggs, T. R., 155, 399, 575, 908, 999,
1048
Riker, A. J., 171, 197, 1005, 1019
Riklis, E., 207, 676, 1028, 1048
Rimington, C, 678, 1048
Rimon, S., 816, 992
Rindi, G., 520, 522, 526, 527, 528, 532,
578, 1003, 1048
Ringler, R. L., 500, 798, 872, 1038
Ritchie, J. L., 838, 1012
Rittenberg, D., 61, 293, 294, 676, 1008
1019, 1028
Rittenberg, S. C, 137, 228, 230, 231,
1064, 1067
Riva, F., 808, 1061
Rivera, G. F., 173, 882, 987
Robbins, W. J., 516, 528, 1048
Robert, B., 772, 773, 836, 1048
Robert, L., 772, 773, 836, 1048
Robert, M., 772, 773, 836, 1048
Roberts, E., 64, 269, 327, 332, 336, 569,
772, 782, 857, 989, 991, 1048, 1051
Roberts, E. R., 292, 294, 1067
Robertson, A. E., Jr., 53, 77, 78, 81, 146,
1000
Robertson, D. H., 625, 992
Robertson, M. E., 971, 1048
Robertson, R. N., 20, 27, 53, 63, 77, 79,
80, 82, 173, 1017, 1067
Robertson, W. van B., 686, 1048
Robie, C. H., 574, 577, 1051
Robins, R. K., 281, 282, 1007
Robinson, B., 353, 1048
Robinson, J. B. D., 966, 994
Robinson, J. C., 551, 709, 843, 1048
Robinson, J. D., 614, 1048
Robinson, J. R., 883, 928, 1048
Robinson, R. J., 76, 1001
Robinson, W. G., 782, 1048
Robson, J., 965, 1048
Robson, J..S., 921, 1029
Rocca, E., 336, 709, 718, 840, 1048
Roche, J., 268, 340, 678, 689, 1047,
1048
Rodnight, R., 485, 1034
Rogach, Z., 297, 1018
Rogers, E. F., 530, 1048
Rogers, H. J., 453, 459, 1048, 1056
Rogers, K. S., 789, 816, 819, 1048
Rogers, L. L.. 588, 1048
Rogers, W. I., 640, 645, 992
Rogers, W. P., 54, 79, 80, 94, 174, 183,
1036
Rogulski, J., 206, 988
Rohdenburg, E. L., 751, 1015
Roholt, D. A., Jr., 660, 1014
Rohutt, O., 683, 783, 845, 1005
Rolf, D., 937, 1066
RoHnson, G. N., 169, 1048
Romanchek, L., 873, 878, 987
Romberger, J. A., 91, 105, 106, 111, 132,
149, 154, 172, 182, 1048
Rona, P., 259, 271, 416, 1037
Ronzoni, E., 766, 1063
Roos, B.-E., 363, 1032
Ropes, M. W., 686, 1048
Rosa, N., 122, 1049
Roscoe, H. E., 837, 1049
Rose, C. L., 999
Rose, F. L., 224, 236, 1003
Rose, I. A., 705, 830, 1049
Rose, W. C., 2, 219, 1001, 1049
Rosell-Perez, M., 391, 1049
Roseman, S., 356, 385, 1004, 1012
Rosen, F., 571, 574, 1049
Rosen, S. M., 208, 909, 914, 1051
Rosen, S. W., 32, 35, 243, 1049
Rosenberg, A. J., 52, 64, 74, 81, 195, 989,
1026, 1049
Rosenberg, H., 685, 704, 707, 710, 836,
845, 1006, 1010
Rosenberg, L. E., 264, 1049
Rosenberg, T., 262, 461, 464, 1004, 1008,
1049, 1066
AUTHOR INDEX
nil
Rosenthal, A., 595, 1049
Ross, C. A., 315, 1057
Ross, H. E., 336, 1032
Ross, J. E., 207, 208, 265, 911, 913,
1018, 1041
Ross, R. T., 224, 1012
Ross, W. F., 15, 999, 1049
Rossi-Fanelli, A., 518, 1041
Rostorfer, H. H., 845, 1001
Roth, J. S., 461, 462, 815, 1043, 1049
Rothberg, S., 853, 1017
Rothemund, E., 18, 30, 1070
Rothschild, A. M., 353, 1049
Rothschild, H. A., 60, 119, 121, 707, 717,
781, 873, 1049
Rothschild, Lord, 175, 875, 882, 1000
Rothstein, A., 876, 883, 893, 894, 895,
897, 898, 899, 900, 902, 903, 904, 905,
906, 907, 908, 958, 959, 960, 1003, 1044,
1049, 1065
Roughton, F. J. W., 757, 1012
Roussos, G. G., 462, 552, 553, 1031,
1049
Rovery, M., 649, 1003
Rowan, K. S., 22, 97, 171, 181, 182, 185,
189, 190, 1016
Rowe, A. W., 803, 1049
Rowe, V. K., 627, 1056
Rowland, R. L., 744, 1049
Roy, A., 177, 178, 1002
Roy, A. B., 443, 444, 1049
Roy, S. C, 53, 153, 228, 838, 839, 1002,
1035
Rozenfel'd, E. L., 417, 1033
Rubbo, S. D., 972, 973, 98S
Rubin, B. A., 170, 1049
Rubin, R. J., 480, 1049
Rubino, R., 877, 1010, 1049
Rubinstein, D., 55, 179, 485, 1049
RuefF, L., 751, 1033
RuflFo, A., 164, 615, 616, 1049
Ruiz-Amil, M., 387, 388, 400, 1056
Rule, N. G., 375, 1033
Rulon, O., 198, 1049
Rumpf, P., 236, 1049
Rundles, R. W., 283, 1006
Runnstrom, J., 726, 964, 1028, 1038,
1049
Ruska, H., 975, 1049
Ruskin, A., 856, 883. 925, 941, 943, 948,
1047, 1049, 1050
Ruskin, B., 856, 883, 948, 1050
Russell, D. S., 951, 952, 953, 954, 1021
Russell, P., 983, 1050
Russo, H. F., 264, 993
Rust, J. H., Jr., 177, 1043
Rutter, W. J., 407, 710, 779, 783, 846,
1048, 1050
Ryan, C. A., 33, 38, 552, 1050
Ryan, E. M., 868, 1032
Ryan, J., 798, 865, 1042
Ryan, K.-J., 555, 713, 1050
Rydon, H. N., 321, 1016
Ryley, J. F., 173, 1050
Ryzhkov, V. L., 194, 1050
Sabato, G. di, 210, 1050
Sabine, J. C, 545, 549, 559, 1068
Sable, H. Z., 413, 1015
Sacerdote, F. L., 1058
Sachs, G., 751, 1050
Sachs, H. W., 926, 1050
Sacktor, B., 154, 446, 780, 798, 865, 1050,
1054
Sadhu, D. P., 138, 144, 149, 218, 1039
Saelhof, C, 399, 400, 1056
SafFran, M., 150, 272, 273, 274, 1050,
1051
Sage, H., 457, 1039
Sahasrabudhe, M. B., 415. 1050
Saiga, Y., 676, 1053
St. John, E., 699, 700, 1012
Saito, K., 435, 1043
Sajgo, M., 817, 827, 1059
Sakai, H., 727, 965, 1050
Sakami, W., 570, 1050
Sakamoto, Y., 64, 564, 578, 676, 858,
1022, 1039, 1043, 1064
Sakata, K., 401, 1050
Salama, A. M., 911, 1060
Salaman, M. H., 225, 990
Salant, W., 942, 944, 946, 947, 948, 1050
Salle, A. J., 683, 685, 691, 692, 737, 968,
972, 973, 1027, 1050
1112
AUTHOR INDEX
Salles, J. B. V., 63, 1050
Saltman, P., 383, 842, 852, 910, 913,
989, 1050
Salvatore, G., 546, 972, 1007
Salvin, E., 98, 99, 104, 110, 1028
Samborski, D. J., 196, 529, 1050
Samiy, A. H., 265, 1015
Samorodin, A. J., 939, 1058
Sanabria, A., 924, 1050
Sanadi, D. R., 844, 856, 873, 1022, 1050
Sanborn, R. C, 21, 29, 1050
Sanders, C. R., 388, 990
Sanderson, P. H., 920, 921, 1002
Sankar, D. V. S., 520, 529, 1050
Sanner, T., 691, 750, 751, 804, 1050
San Pietro, A., 490, 510, 553, 850, 851,
891, 1003, 1025, 1030, 1050
Santi, R., 983, 984, 992
Santilli, V., 741, 980, 1025
Sanwal, B. D., 332, 840, 845, 1050
Sagir, D., 924, 1065
Sargent, J. R., 693, 842, 1013
Sarkar, N. K., 710, 849, 1063
Sarma, P. S., 522, 676, 681, 711, 712,
1046, 1054, 1058
Saroff, H. A., 763, 1051
Sarreither, W., 125, 1029
Sasaki, A., 794, 1051
Sasaki, S., 78, 79, 86, 349, 844, 1024, 1051
Sato, R., 842, 1051
Sato, T. R., 447, 1060
Sato-Asano, K., 817, 1041
Satta, G., 726, 1033
Sauer, G., 818, 1051
Sauermann, G., 397, 1051
Saunders, P. R., 55, 56, 76, 81, 178, 181,
183, 191, 214, 215, 993, 1036, 1065
Sawai, T., 420, 1051
Sawatzky, H., 744, 1054
Sayre, F. W., 332, 1051
Saz, A. K., 599, 1051
Saz, H. J., 26, 168, 1051
Scala, A. R., 193, 1005
Scala, R. A., 539, 1051
Scarano, E., 469, 675, 706, 1051
Schaal, R., 236, 1049
Schachter, D., 208, 355, 909, 914, 1004,
1051
Schacter, H., 694, 1051
Schaefer, M. A., 870, 1028
Schaeg, W., 776, 1065
Schanker, L. S., 911, 1051
Schapira, G., 308, 1045, 1051
Scharff, T. G., 263, 1051
Schatz, V. B., 257, 261, 1051
Schatzberg, G., 878, 989
Schauer, R., 910, 1051
Schayer, R. W., 353, 363, 1049, 1051
Scheffer, R. P., 27, 1068
Schellenberg, K. A., 673, 702, 703, 816,
1017, 1051
Schenker, H., 208, 909, 914, 1004, 1051
Scheraga, H. A., 683, 998
Schimmel, N. H., 626, 1051
Schlegel, D. E., 194, 1051
Schlegel, V., 29, 1034
Schleyer, H., 485, 487, 490, 1017
Schlieselfeld, L. H., 706, 718, 1030
Schmid, C, 120, 873, 878, 1015
Schmidt, G., 439, 1051
Schmidt, S., 297, 1051
Schmitt, A., 707, 1054
Schmitt, J. A., 287, 289, 1044
Schneider, G., 297, 1051
Schneider, K. C, 293, 1019
Schneider, S., 432, 817, 1020
Schneider, W. C., 3, 1045
Schneiderman, H. A., 199, 1051
Schoeller, W., 941, 951, 956, 957, 1040
Schom, R., 875, 1051
Schonbaum, E., 150, 1051
Schoniger, W., 4, 1019
Schoepke, H. G., 214, 1063
Schorcher, C, 924, 1051
Scholefield, P. G., 62, 122, 177, 200. 238,
265, 266, 267, 395, 432, 596, 987,
1001, 1046, 1051
Schollmeyer, P., 178, 1027
Schopfer, W. H., 529, 1051
Schor, J. M., 325, 1010
Schramm, M., 408, 409, 1046
Schrauth, W., 941, 951, 956, 957, 1040
Schrecker, A. W., 582, 1051
Schrodt, G. R., 574, 577, 1051
Schroeder, E. A. R., 837, 1046
Schroeder, E. F., 689, 1051
AUTHOR INDEX
1113
Schueler, F. W., 44, 261, 1051
Schiller, H., 670, 1051
Schiitte, H. R., 803, 1052
Schuler, M. N., 77, 87, 104, 168, 1038
Schuler, W., 362, 1051
Schulman, M. P., 831, 1055
Schultz, G., 1020
Schultz, S. G., 387, 1052
Schulz, D. W., 407, 474, 1024
Schulz, H., 875, 1052
Schuize, H. O., 32, 388, 389, 391, 400,
990
Schumann, E. L., 358, 1052
Schutz, B., 210, 1021
Schwarz, K., 878, 1001
Schwartz, W., 195, 1052
Schwarz, D. R., 1000
Schwarzenbach, G., 732, 739, 994
Schweet, R. S., 307, 1052
Schweisfurth, R., 195, 1052
Schwerin, B. G., 664, 1045
Schwert, G. W., 373, 432, 433, 435, 436,
782, 787, 802, 811, 1015, 1038, 1041,
1042, 1054, 1059, 1067
Scott, J. J., 600, 998
Scott, R. L., 722, 909, 914, 1052
Scott, J. W., 77, 1052
Scutt, P. B., 292, 294, 1044
Seal, U. S., 802, 1052
Sealock, R. R., 518, 524, 1032, 1052
Seaman, G. R., 18, 28, 37, 71, 74, 91,
173, 241, 242, 243, 1052
Sebrell, W. H., 495, 1033
Seeley, H. W., 591, 830, 1052
Seehch, F., 125, 399, 695, 1031, 1043,
1052
Seevers, M. H., 620, 622, 623, 624, 625,
627, 628, 629, 630, 631, 1052, 1053,
1068
Segal, H. L., 334, 358, 412, 472, 641,
714, 760, 766, 806, 810, 856, 995,
1012, 1020, 1052
Segal, R., 816, 992
Segal, S., 264, 1049
Seibert, M. A., 835, 870, 871, 1028, 1052,
1056
Seibert, R. A., 933, 1016, 1039
Sekiya, K., 868, 1061
Sekuzu, I., 26, 843, 1042, 1052
Sela, M., 457, 459, 462, 1048, 1052
Seligman, A. M., 856, 1040
Selim, A. SS. M., 357, 712, 1052
Sellinger, 0. Z., 335, 1052
Semina, L. A., 307, 1036
Sen, D. K., 338, 1052
Senthe Shanmuganathan, S., 783, 857,
1052
Serif, G. S., 392, 393, 404, 1052, 1066
Servettaz, O., 850, 1036
Sery, T. W., 978, 981, 1052
Severac, M., 957, 1046
Severens, J. M., 972. 983, 984, 985, 1052
Severin, E. S., 359, 1026
Sgaros, P. L., 168, 225, 1052
Shacter, B., 722, 879, 880, 883, 1052
Shaffer, C. F., 944, 957, 998
Shah, V. K., 830, 1052
Shaner, G. A., 264, 993
Shanes, A. M., 211, 1053
Shannon, L. M., 225, 226, 227, 228, 232,
1053, 1069
Shapiro, D. M., 288, 505, 538, 569, 570,
577, 1004, 1010, 1053
Sharon, N., 326, 868, 1032, 1053
Sharp, A. G., 213, 747, 1065
Sharpensteen, H. H., 750, 1010
Shaw, E., 281, 282, 1053
Shaw, P. D., 453, 1053
Shaw, W. H. R., 64, 603, 610, 976,
1026, 1053
Shcherbakova, L. I., 774, 976, 1044
Sheets, R. F., 902, 905, 1053
Sheinfeld, S., 458, 1031
Sheinin, R., 418, 1053
Shemin, D., 160, 1053, 1068
Shepherd, D. M., 310, 353, 1034, 1048
Sherman, I. R., 273, 1053
Sherman, R., 367, 1014
Shetlar, M. R., 838, 1032
Shibasaki, I., 633, 851, 1053
Shichi, H., 553, 1053
Shideman, F. E., 32, 204, 205, 212, 620,
622, 623, 624, 625, 626, 627, 628, 629,
630, 631, 1004, 1047, 1052, 1053, 1058,
1065, 1068
Shifrin, S., 500, 784, 1053
1114
AUTHOR INDEX
Shigeura, H..T., 467, 1053
Shiio, I., 154, 880, 1053, 1061
Shilo, M., 601, 1053
Shils, M. E., 505, 538, 569, 570, 1053
Shimazono, H., 63, 1053
Shimi, I. R., 91, 678, 1053
Shimizu, T., 179, 1059
Shimomura, O., 676, 1053
Shine, H. J., 372, 1009
Shinohara, K., 747, 1054
Shiraki, M., 551, 555, 1022
Shive, W., 354, 588, 1001, 1048
Shkol'nik, M. Y., 15, 1053
Shonk, C. E., 587, 995
Shookhoff, M. W., 5, 1066
Shore, B., 873, 922, 926, 927, 1053
Shore, V., 873, 922, 926, 927, 1053
Shorr, E., 166, 1018
Shrago, E., 846, 851, 1053
Shrivastava, D. L., 692, 831, 987
Shug, A. L., 293, 294, 854, 977, 981,
1053, 1054, 1066
Shukuya, R., 811, 1054
Shull, K. H., 772, 846, 1054
Shulman, A.,531,i05-^
Shwartzman, G., 569, 1054
Sibly, P. M., 834, 1054
Sie, H.-G., 428, 1054
Siebert, G., 707, 844, 852, 1054
Siegel, I. A., 216, 1065
Siegel, L., 804, 1054
Siegel, S. M., 750, 1010
Siegelman, H. W., 173, 182, 1054
Siekevitz, P., 444, 445, 865, 872, 1033,
1054
Sights, W. P., 878, 883, 991
Sih, C. J., 708, 837, 870, 1054
Silber, R., 582, 1054
Silber, R. H., 562, 572, 1045
Silberman, H. R., 281, 1054
Sihprandi, N., 518, 1041
Sillen, L. G., 732, 733, 734, 736, 739,
994, 1018, 1054
Silva, O. L., 693, 1056
Silva, R. B., 745, 754, 760, 1034
Silverman, J. L., 382, 1054
Silverman, M., 582, 1010
Simola, P. E., 109, 1054
Simon, E. W., 19, 28, 60, 182, 837, 855,
870, 1054
Simon, L. N., 474, 1054
Simonds, J. P., 924, 926, 1018, 1054
Simonsen, D. G., 772, 782, 1048
Simonsen, D. H., 50, 1054
Simpson, F. J., 713, 833, 857, 859, 993,
1012, 1054
Simpson, J. R., 450, 1031
Simpson, R. B., 739, 740, 744, 745, 748,
759, 1054
Singer, B., 741, 1054
Singer, T. P., 16, 18, 29, 32, 38, 41, 46,
48, 49, 65, 435, 437, 510, 541, 542,
547, 549, 668, 673, 675, 676, 693, 703,
706, 710, 713, 716, 718, 720, 721, 783,
798, 803, 825, 833, 840, 845, 846, 857,
866, 870, 877, 878, 991, 994, 1006, 1014,
1025, 1036, 1038, 1054, 1065
Singer, Altbeker, R., 532, 992
Sipos, J. C, 744, 1054
Sirsi, M., 490, 847, 1013
Siu, P. M. L., 852, 1054
Sivaramakrishnan, V. M., 522, 1054
Sizer, I. W., 64, 334, 551, 610, 657, 658,
660, 675, 676, 685, 686, 692, 709, 843,
1015, 1023, 1048, 1054, 1055
Sjoerdsma, A., 315, 1042
Skarnes, R. C, 459, 1055
Skeggs, H. R., 264, 993
Skipper, H. E., 585, 1055
Skoda, J., 474, 1055
Skou, J. C, 865, 869, 1055
Slater, E. C, 33, 60, 61, 80, 83, 85, 121,
122, 713, 715, 718, 810, 816, 820, 825,
826, 867, 869, 871, 1032, 1040, 1055
Slater, G. G., 585, 1066
Slater, T. F., 553, 850, 1055
Slaughter, C, 357, 1008
Slechta, L., 476, 1055
Slein, M. W., 376, 1055
Slingerland, D. W., 883, 910, 1055
Sliwinski, R. A., 406, 1047
Sloane, N. H., 356, 1055
Sloboda, A., 504, 505, 1016
Slotin, L., 63, 1007
Sloviter, H. A., 401, 1050
SmaU, P. A., 658, 1045
AUTHOR INDEX
1115
SmaUey, H. M., 231, 997
Smalt, M. A., 744, 870, 1055
Smeby, R. R., 225, 992
Smiley, J. D., 61, 329, 998
Smiley, R. L., 363, 1051
Smillie, R. M., 27, 91, 1055
Smith, A. H., 89, 104, 109, 1010, 1043
Smith, C. G., 201, 413, 577, 968, 1055
Smith, D. E., 956, 972, 1055
Smith, E. E. B., 61, 427, 1038
Smith, E. L., 60, 365, 375, 667, 668, 769,
770, 783, 804, 1008, 1019, 1026, 1055,
1057
Smith, F., 15, 1055
Smith, G. B. L., 362, 1032
Smith, G. N., 231, 997
Smith, H. M., 949, 1055
Smith, J. E., 840, 1055
Smith, J. T., 598, 615, 676, 711, 717,
1016, 1055
Smith, L., 163, 168, 892, 1055
Smith, L. C, 434, 1034
Smith, L. H., Jr., 467, 470, 479, 1055
Smith, M. E., 507, 709, 717, 841, 1055
Smith, O. H., 49, 1044, 1065
Smith, P. F., 592, 1055
Smith, P. H., 953, 1064
Smith, P. J. C, 382, 832, 1055
Smith, S. E., 309. 310, 315, 318, 1055
Smith, W., 550, 1001
Smyrniotis, P. Z., 413, 855, 1021
Smyth, D. H., 77, 87, 94, 95, 176, 265,
387, 991, 1023, 1041, 1056
Sneed, M. C, 736, 1055
Snell, E. E., 358, 452, 475, 539, 543, 544,
561, 564, 565, 569, 575, 576, 578, 833,
1001, 1003, 1020, 1026, 1033, 1034,
1042, 1046, 1064
Snodgrass, P. J., 743, 785, 789, 825, 831,
1055
Snoswell, A. M., 18, 547, 551, 1055
Snow, N. S., 756, 998
Snyder, R., 831, 1055
Snyder, S. H., 693, 1056
Soars, M. H., 582, 589, 1018
Sorbo, B., 803, 1056
Sorbo, B. H., 713, 1056
Sohler, M. R., 835, 1056
Sohonie, K., 833, 1040
Sokoloff, B., 399, 400, 1056
Soldatenkov, S. V., 224, 225, 1056
Solomon, A. K., 435, 1066
Solomon, J. B., 686, 845, 846, 1056
Sols, A., 376, 379, 381, 382, 383, 387,
388, 389, 391, 400, 414, 782, 824, 843,
1001, 1056
Solvonuk, P. F., 825, 836, 854, 1056
Somers, E., 976, 1056
Sondheimer, E., 595, 1056
Soodak, M., 516, 523, 530, 533, 998, 1056
Sophianopoulos, A. J., 610, 1056
Sorkin, E., 518, 530, 1006
Sorm, F., 474, 1055
Sormova, Z., 474, 1055
Sorof, S., 675, 703, 706, 716, 718, 833,
1006
Sorsby, A., 953, 1056
Sourkes, T. L., 308, 325, 544, 816,
1029, 1056, 1067
Southwick, P. L., 518, 531, 1006
Spackman, D. H., 783, 1055
Sparrow, B. W. P., 967, 994
Speakman, J. B., 762, 1019
Speck, J. F., 53, 73, 74, 78, 91, 93, 124,
1056
Spector, A., 662, 1056
Spector, W. G., 663, 909, 914, 1056
Speer, H. L., 169, 1056
Spencer, A. F., 147, 1056
Spencer, B., 427, 444, 684. 1004, 1056
Spencer, D., 615, 1056
Spencer, H. C, 627, 1056
Spencer, R. P., 267, 268, 686, 1056
Spensley, P. C., 453, 459, 1048, 1056
Speyer, J. F., 273, 1056
Spiegelman, S., 326, 351, 875, 1016, 1056
Spiro, R. G., 1056
Spizizen, J., 194, 1056
Spoeri, E., 195, 727, 972, 973, 1033
Spooner, D. F., 168, 179, 693, 882, 1010
Spragg, S. P., 852, 1021
Spatt, N. T., 199, 1056
Spriestersbach, D., 15, 1055
Sprinson, D. B., 413, 1056
SpjTopoules, C. S., 950, 995
Sreenivasan, A., 478, 993
1116
AUTHOR INDEX
Sreenivasaya, M., 15, 1024
Srere, P. A., 855, 1059
Srinivasan, P. R., 413, 969, 1056, 1057
Stachiewicz, E., 435, 437, 1029
Stadtman, E. R., 356, 452, 1000, 1057
Staehelin, M., 318, 1035
Staemmler, M., 924, 1057
Stafford, H. A., 225, 1057
Stafford, J. E., 358, 1002
Stahmann, M. A., 456, 457, 458, 1003,
1027, 1061
Stalder, K., 98, 225, 226, 228, 235, 1057,
1060
Stamer, J. R., 709, 1030
Stanbury, F. A., 730, 735, 962, 990
Standen, O. D., 772, 1041
Stanley, R. G., 27, 74, 80, 82, 1057
Stanley, W. M., 979, 980, 98S
Stansly, P. G., 233, 992
Staple, E., 150, 1066
Stare, F. J., 55, 56, 75, 79, 81, 114, 115,
141, 164, 175, 176, 991, 1011, 1057
Stark, J. B., 15, 1043, 1057
Starr, J. L., 817, 820, 988, 1057
Stauff, J., 760, 1057
Stauffer, J. F., 171, 1018
Steam, A. E., 373, 1057
Stebbins, R. B., 577, 1040
Steberl, E. A., 234, 1057
Stedman, R. L., 74, 77, 632, 1057
Steel, K. J., 971, 972, 974, 1001, 1057
Steel, R., 677, 1047
Steele, A. B., 620, 1057
Steele, R., 401, 988
Stegner, H.-E., 923, 1067
Stein, E., 942, 945, 946, 1027, 1057
Stein, W. D., 906, 990
Steinberg, D., 351, 614, 1057
Steiner, D. F., 391, 1057
Steiner, L. A., 459, 1052
Stengle, J., 388, 401, WL^9
Stent, H. B., 225, 226, 1058
Stephenson, M. L., 156, 887, 1057
Stern, J. R., 80, 228, 233, 234, 235, 708,
751, 774, 837, 1037, 1057
Stetten, D. Jr., 208, 1044
Stevens, C. 0., 709, 1057
Stevenson, T. D., 574, 577, 1051
Stewart, C. J., 392, 393, 394, 403, 404,
996, 999, 1052, 1066
Stewart, H. B., 518, 1057
Steyn-Parve, E. P., 521, 525, 526, 1025,
1027, 1057
Stickland, R. G., 65, 238, 597, 1057
Stjernholm, R., 224, 235, 1057, 1067
Stock, C. C, 576, 1024
Stockell, A., 375, 497, 1057
Stoeken, L. A., 956, 1057
Stoerk, H. C, 566, 576, 577, 587, 995,
1057
Stolen, J. A., 228, 232, 1032
Stolzenbach, F. E., 497, 510, 1024, 1025
Stone, B. A., 132, 1057
Stone, C. A., 315, 1057
Stone, J. E., 478, 1058
Stone, R. W., 26, 36, 1012
Stoneman, F., 626, 1058
Stoppani, A. 0. M., 18, 32, 38, 41, 74,
77, 79, 81, 92, 104, 116, 117, 169, 687,
705, 706, 708, 711, 712, 713, 716, 717,
775, 778, 779, 780, 781, 783, 810, 832,
838, 852, 853, 998, 1003, 1007, 1058
Storey, I. D. E., 713, 817, 859, 860, 1058
Stotz, E., 143, 151, 1036, 1067
Stracher, A., 792, 866, 867, 1058
Straessle, R., 681, 759, 761, 1005, 1021,
1058
Strait, L. A., 694, 1062
Strandskov, F. B., 690, 1068
Straub, F. B., 29, 1058
Straub, R. W., 211, 949, 1014
Strauss, N., 33, 38, 1058
Strayhorn, W. D., 745, 750, 1002
Strecker, H. J., 336, 410, 553, 850, 1012,
1016, 1058
Streicher, J. A., 155, 999
Strelitz, F., 677, 1037
Strength, D. R., 585, 1058
Strickland, K. P., 1058
Strickler, J. C., 206, 1058
Stricks, W., 747, 748, 763, 1027, 1058
Strittmatter, C. F., 547, 711, 787, 847,
1058
Strittmatter, P., 685, 804, 809, 870,
1058
AUTHOR INDEX
1117
Strohman, R. C, 939, 1058
Strominger, J. L., 359, 662, 713, 858,
859, 1020, 1037, 1058
Strong, F. M., 260, 489, 504, 1068
Stulberg, M. P., 812, 1027
Stumpf, P. K., 61, 64. 136, 137, 138, 145,
148, 151, 226, 228, 231, 684, 830, 832,
1012, 1017, 1037, 998, 1021, 1039,
1058
Subrahmanyan, V., 407, 684. 692, 833,
1018
Subramaniam, V., 2, 225, 226, 228,
998, 1058, 1064
Suda, M., 857, 1021
Siipfle, K., 975, 1058
Sugiura, H. T., 462, 1043
Sullivan, L. P., 924, 1062
Sullivan, M., 467, 470, 479, 1055
Sullivan, R. D., 538, 1030
Sullivan, W. J., 929, 1014
Sumizu, K., 859, 1058
Summers, W. A., 576, 1058
Summerson, W. H., 304, 1031
Sund, H., 508, 785, 788, 792, 810, 831,
1064
Sundaram, S., 676, 712, 1058
Sundaram, T. K., 711, 1046, 1058
Surtshin, A., 959, 1058
Sussman, A., 195, 1058
Sussman, M., 875, 1056
Sutherland, V. C, 179. 181, 1006
Sutton, C. R., 780, 783, 810, 814, 1059
Sutton, W. B., 552, 1059
Suzuoki, T., 173, 596, 924, 1059
Suzuoki, Z., 173, 596, 882, 1041, 1059
Svanberg, 0., 813, 837, 1063
Svedberg, A., 928, 997
Svennerholm, L., 76, 81, 152, 1033
Swan, A. A. B., 961, 1010
Swan, J. M., 694, 1059
Swenson, A. D., 643, 780, 788, 803, 833,
1059
Swensson, A., 959, 960, 1059
Swim, H. E., 89, 1003, 1059
Swingle, K. F., 35, 36, 1059
Syrett, P. J., 237, 1037
Szabolcsi, G., 649, 788, 803, 812, 817,
827, 1059
Szego, C. M., 31, 50, 176, 1031
Szeinberg, A., 497, 1036
Szent-Gyorgui, A., 689, 1038
Szep, O., 960, 1059
Szulmajster, J., 52, 60, 74, 81, 989
Tabachnik, M., 855, 1059
Tabor, H., 585, 1059
Tachibana, S., 543, 554, 1059
Tada, M., 537, 1005
Taeger, H., 950, 1015
Taufel, K., 168, 228, 237, 504, 1040
Tager, J. M., 153, 1059
Taggart, J. V., 204, 355, 921, 928, 1002,
1009, 1051
Tahmisian, T. N., 876, 991
Takagaki, G., 126, 127, 135, 153, 176,
1059, 1061
Takagi, Y., 471, 1059
Takahashi, H., 51, 74, 77, 1059
Takahashi, M., 880, 1053
Takahashi, N., 444, 675, 816, 1059
Takamiya, A., 787, 1025
Takebe, I., 444, 1059
Takemori, A. E., 597, 1059
Takemori, S., 842, 1051
Takenaka, Y., 782, 811, 1059
Takeuchi, T., 603, 610, 1059
Talalay, P.. 447, 449, 712, 713, 1032,
1036, 1059
Tamari, M., 285, 992
Tamura, T., 618, 632, 1062
Tanada, T., 909, 1059
Tanaka, K., 747, 973, 1059
Tanaka, M., 542, 1059
Tanaka, R., 518, 522, 523, 1035
Tanaka, S., 179, 443, 859, 1058, 1059
Tanaka, Y., 151, 676, 678, 887, 1059
Tangen, 0., 857, 1008
Tanner, F. W., 972, 983, 984, 985, 1052
Tapley, D. F., 90, 207, 208, 210, 265,
681, 703, 762, 819, 911, 913, 1014,
1018, 1023, 1024, 1041, 1059
Tappel, A. L., 75, 80, 91, 121, 137, 996,
1015
Tarr, H. L. A., 411, 1059
1118
AUTHOR INDEX
Tasaki, I., 950, 995
Tashian, R. E., 329, 1059
Tashiro, M., 340, 1040
Tata, J. R., 555, 778, 1059, 1060
Taube, H., 943, 944, 956, 957, 1031
Tauroq, A., 209, 1060
Taylor, A., 494, 987
Taylor, B. B., 670, 690, 1006
Taylor, E. L., 409, 1060
Taylor, E. S., 660, 845, 1060
Taylor, F. J., 379, 391, 1060
Taylor, H. D., 956, 1060
Taylor, J., 601, 995
Taylor, J. J., 859, 1060
Taylor, K. B., 55, 179, 228, 988
Teal, J. M., 169, 1019
Teitell, L., 228, 993
Tekman, S., 645, 1017
Telegdi, M., 409, 1025
Telkka, A., 925, 1040, 1060
Tenebaum, L. E., 333, 1012
Terner, C, 75, 153, 176, 1060
Terui, G., 633, 851, 1053
Testa, E., 615, 1049
Thannhauser, S. J., 439, 1051
Theis, F. V., 696, 1060
Theorell, H., 508, 544, 688, 779, 780,
784, 785, 1060, 1069
Therattil- Antony, T., 704, 723, 939, 990
Thienes, C. H., 55, 56, 76, 81, 178, 181,
183, 214, 1065
Thiessen, C. P., 678, 992
Thimann, K. V., 156, 196, 887, 966, 968,
1000, 1057, 1060
Thoai, N., 268, 340, 1048
Thoelen, H., 954, 1060
Thomas, C. A., 741, 1066
Thomas, G. M., 780, 798, 865, 1050
Thomas, G. W., 972, 973, 1060
Thomas, K., 98, 225, 226, 228, 235, 1060
Thompson, C. C, 168, 169, 190, 999
Thompson, E. O. P., 770, 1055, 1060
Thompson, J. W., 667, 688, 707, 1014,
1037
Thompson, R. H. S., 854, 1060
Thompson, R. L., 193, 1060
Thompson, T. E., 789, 816, 819, 1048
Thompson, W., 858, 1003, 1060
Thomson, J. F., 34, 38, 240, 447, 1060
Thorn, M. B., 18, 19, 22, 23, 24, 25, 32,
33, 42, 717, 718, 1060
Thome, C. J. R., 807, 846, 1060
Thorne, K. J. I., 886, 1060
Torsell, W., 464, 465, 989
Threefoot, S., 928, 997
Thunberg, T., 2, 32, 35, 37, 40, 175, 185,
1060
Thurlow, S., 280, 1004
Thyagarajan, B. S., 672, 1060
Ticha, M., 389, 1027
Tieckelmann, H., 531, 990
Tietz, A„ 146, 147, 228, 233, 234, 887,
1045, 1060
Tietze, F., 29, 37, 242, 243, 1027, 1060
Tilak, B. D., 415, 1050
Tinacci, F., 218, 219, 1060
T'ing-seng, H., 662, 857, 1060
Tin-Sen', S., 360, 995
Tishler, M., 538, 1006
Tissieres, A., 53, 1017
Titus, E. O., 281, 282, 285, 1005
Tjutjunnikowa, A. W., 835, 999
Tobari, J., 845, 1026
Tobie, E. J., 91, 127, 995
Tocco, D. J., 911, 1051
Todd, A. R., 516, 992
Tokuyama, K., 673, 675, 692, 717, 782,
1060
Tolba, M. K., 911, 973, 978, 1060
Tolbert, N. E., 61, 999
Tolman, L., 586, 1036
Tomchick, R., 843, 996
Tomkins, G. M., 449, 1034
Tong, W., 209, 1060
Tonhazy, N. E., 79, 1060
Tonomura, Y., 816, 820, 866, 867, 868,
939, 940, 1060, 1061
Tooth, B. E., 678, 1048
Topper, Y. J., 385, 1061
Torda, C, 165, 1061
Tosi, L., 54, 114, 174, 552, 708, 870,
994, 1011
Tosteson, D. S., 937, 1066
Totaro, J. A., 315, 316, 317, 318, 1045,
1057
AUTHOR INDEX
1119
Totter, J. R., 585, 845, 1001, 1061
Tower, D. B., 391, 392, 394, 395, 399,
834, 1061
Towsend, E., 325, 1056
Trazler, G., 707, 1054
Traniello, S., 708, 837, 1061
Traub, A., 20, 29, 61, 80, 844, 848, 989,
1013
Trembley, R., 864, 997
Trim, A. R., 741, 1061
Troger, R., 975, 1061
Trudinger, P. A., 321, 323, 1061
Truscoe, R., 177, 209, 1035
Tsao, T.-C, 939, 1061
Tschudy, D. P., 591, 1061
Tsen, C. C, 833, 845, 859, 901, 906,
1061
Tseng, N. S., 513, 1062
Tsuboi, K. K., 452, 711, 713, 788, 839,
842, 855, 1061, 1062
Tsukada, Y., 126, 127, 135, 153, 176,
1059, 1061
Tsukaraoto, H., 196, 1061
Tsunoda, T., 154, 1061
Tsurumaki, T., 985, 1061
Tsuyuki, E., 457, 1061
Tsuyuki, H., 457, 1061
Tubbs, P. K., 61, 552, 1061
Tuck, L. D.. 694, 1062
Tull, F. A., 400, 1006
Tuppy, H., 641, 1067
Turano, C, 808, 827, 1061
Turba, F., 641, 938, 939, 1007, 1029,
1061
Turner, A. W., 707, 1031
Turner, D. H., 831, 1061
Turner, J. F., 831, 853, 1017, 1061
Turner, J. S., 22, 97, 171, 181, 182, 185,
189, 190, 1016, 1061
Turner, W. A., 2, 225, 1061
Turpaev, T. M., 947, 951, 958, 959, 1010,
1042, 1061
Turrian, H., 956, 1061
Turrian, V., 956, 1061
Tuttle, L. P., 459, 993
Tutunji, B., 95, 128, 1007
Tyler, D. B., 133, 1061
Tytell, A. A., 610, 1054
u
Uchida, M., 64, 1043
Udaka, S., 481, 1062
Udenfriend, S., 266, 310, 315, 316, 317,
318, 319, 320, 354, 611, 999, 1002,
1018, 1033, 1042, 1062
tjhlein, E., 760, 1057
Ukita, T., 618, 632, 1062
Ulbricht, T. L. V., 530, 1062
Ullberg, S., 958, 993
Ulmer, D. D., 785, 1032
Ulrich, F., 209, 909, 1062
Ulrich, J. A., 529, 1062
Umbreit, W. W., 168, 350, 564, 566,
1062, 1063
Umeraura, Y., 439, 1062
Ungar, G., 387, 1062
Ungar-Waron, H., 285, 992, 993
Unna, K., 562, 1062
Urata, G., 674, 1062
Uritani, I., 439, 1062
Urivetzky, M., 855, 1062
Usami, S., 78, 79, 86, 349, 1024
Utter, M. F., 676, 853, 1025
Utzinger, G. E., 694, 1062
Vagelos, P. R., 232, 234, 751, 774, 847,
1020, 1062
Vahlhaus, E., 855, 996
Vaidyanathan, C. S., 490, 547, 549,
615, 660, 685, 832, 847, 1013, 1029,
1047
Valentine, R. J., 268, 1056
Vallee, B. L., 743, 746, 770, 780, 781,
785, 789, 806, 818, 825, 827, 831,
1000, 1001, 1019, 1032, 1055, 1062
Van Aken, G. M. F. A., 745, 998
Van Arsdel, P. P., 725, 949, 995, 1062
Van Baerle, R. R., 332, 550, 551, 1013,
1062
Van Bibber, M. J., 475, 992
Vandemark, P. J., 886, 1006
Vandendriessche, L., 462, 693, 1036,
1062
Vander, A. J., 920, 924, 1062
1120
AUTHOR INDEX
Van der Linden, A. C, 5ie, 774, 775, 854,
1043
Van der Schoot, J. B., 320, 1062
Van Duuren, A. J., 15, 1062
Van Eys, J., 434, 497, 498, 508, 513,
784, 804, 842, 1062, 1064
Van Grembergen, G., 54, 173, 1062
Van Heyningen, R., 596, 662, 712, 846,
1062, 1064
Van Oorschot, J. L. P., 597, 1062
Vanov, S., 611, 1062
Van Pilsun, J. F., 285, 1063
Van Rheenen, D. L., 521, 525, 1025, 1027
Van Thoai, N., 779, 780, 1063
Van Vals, G. H., 130, 149, 150, 156, 1063
Van Wagtendonk, W. J., 50, 1054
Vardanis, A., 474, 1063
Varela, 877, 1049
Vargas, R., 918, 1063
Varner, J. E., 269, 336, 662, 841, 1063
Varrone, S., 678, 689, 1047
Vasarhely, F., 470, 1037
Vasilevskis, J., 739, 1008
Vasington, F. D., 209, 909, 910, 1063
Vaslow, F., 373, 1063
Vasta, B. M., 122, 1022
Vaughan, M., 351, 1057
Vavra, J., 334, 1063
Veeger, C, 18, 38, 1003
Veitch, F. P., 238, 1063
Velick, S. F., 334, 766, 770, 785, 786,
787, 802, 824, 870, 1058, 1063
Velluz, L., 518, 1063
Vely, V. G., 26, 989
Venditti, J. M., 496, 1024
Vennesland, B., 63, 226, 470, 1007,
1010, 1063
Ventura, U., 578, 1048
Venturi, V. M., 214, 1063
Vercauteren, R., 676, 1063
Vernberg, W. B., 173, 183, 1063
Vernon, L. P., 557, 710, 847, 849, 1004,
1063, 1066
Vescia, A., 594, 708, 837, 1061, 1063
Vesthing, C. S., 437, 990
Vickery, H. B., 91, 105, 107, 172, 190,
225, 228, 1063
Villee, C. A., 393, 399, 1063
Villela, G. G., 288, 289, 1063
Vilter, R. W., 577, 1040
Vincent, P. C, 902, 906, 908, 1063
Vining, L. C, 547, 549, 1040
Virtanen, A. I., 293, 706, 1029, 1039
Vishniac, W., 675, 1026, 1063
Vishwakarma, P., 207, 1063
Visscher, M. B., 910, 1022
Vitale, J. J., 208, 1063
Vogel, A. I., 3, 1063
Vogel, W., 664, 1045
Vogel, W. H., 831, 1055
Vogler, K. G., 168, 1063
Volk, W. A., 411, 1063
Von Boventer-Heidenhain, B., 195, 1058
von Brand, T., 18, 25, 28, 91, 127, 173,
203, 882, 987, 995
von Bruchhausen, F., 505, 1063
von Euler, H., 61, 500, 504, 685, 813,
837, 987, 1063
von Euler, L., 901, 1038
Von Holt, C, 520, 1063
w
Wachi, T., 64, 1043
Wachstein, M., 926, 927, 1064
Wachter, W., 701, 1037
Wada, E., 225, 1064
Wada, H., 561, 564, 578, 676, 858, 1039,
1064
Waddell, J. G., 564, 1062
Wadkins, C. L., 18, 543, 555, 865, 872,
874, 1031, 1064
Wadso, I., 9, 1005
Wadzinski, I. M., 325, 1009
Waelsch, B., 783, 1038
Waelsch, H., 887, 1000
Wagner, R. H., 817, 818, 820, 1047
Wagreich, H., 298, 299, 301, 1029
Wahl, R., 979, 980, 1064
Wainio, W. W., 60, 61, 550, 1005
Waisman, H. A., 325, 1009
Wajda, I., 363, 364, 994
Wakelin, R. W., 36, 1044
Wakerlin, G. E., 956, 1064
Wakil, S. J., 146, 224, 234, 707, 781, 887,
996, 1014, 1064
AUTHOR INDEX
1121
Walaas, E., 541, 545, 1064
Walaas, O., 541, 545, 1064
Wald, G., 953, 1064
Waldi, I)., 518, 530, 1006
Waley, S. G., 594, 662, 712, 1000, 1062,
1064
Walker, A. D., 77, 78, 1019
Walker, D. G., 389, 1064
Walker, E., 982, 1064
Walker, G. C, 787, 1064
Walker, J., 960, 1014
Walker, L. M., 908, 1048
Walker, M., 428, 1030
Walker, P. G., 419, 420, 1046, 1064
Walker, T. K., 2, 225, 226, 228, 399,
998, 1048, 1068, 1064
Wallace, A., 225, 226, 852, 1021, 1048,
1064
Wallace, D. M., 428, 995
WaUace, R. A., 373, 1064
Wallace, R. H., 74, 77, 190, 228, 999
Wallach, D. P., 358. 1052, 1064
Wallenfels, K., 508, 785, 788, 792, 810,
831, 1064
Wallgren, H., 1064
Walsh, E. O'F., 837, 1064
Walsh, G., 837, 1064
Walter, C, 869, 1064
Walter, P., 508, 1064
Waltman, J. M., 137, 1064
Wanczura, T., 619, 1035
Wang, T. P., 509, 1064
Warburg, 0., 695, 768, 1064
Waring, G. B., 582, 1009
Warkany, J., 953, 1064
Warmke, H. E., 244, 1039
Warner, C, 297, 298, 1064
Warner, R. C, 463, 1064
Warnock, L. G., 434, 1064
Warringa, M. G. P. J., 33, 38, 49, 773,
855, 1064, 1065
Warzecha, K., 814, 861, 1015
Wase, A., 960, 997
Washio, S., 854, 1065
Wasson, G. W., 234, 1057
Watanabe, K., 675, 1065
Watanabe, S., 439, 1022
Waterhouse, D. F., 18, 1065
Watertor, J. L., 197, 1019
Watkins, W. M., 417, 418, 419, 1065
Waton, N. G., 352, 362, 1065
Watson, D. W., 459, 1055
Watt, D. D., 430, 1065
Watts, R. W. E., 165, 1001
Way, J. L., 478, 1002
Weakley, D. R., 412, 1030
Weatherall, M., 901, 905, 1024
Weaver, M. E., 200, 1065
Webb, E. C, 444, 1065
Webb, J. L., 31, 35, 36, 46, 47, 48, 51,
55, 56, 69, 70, 71, 75, 76, 81, 82, 83,
88, 178, 181, 183, 213, 214, 217, 240,
241, 625, 896, 943, 944, 945, 1039,
1065
Webb, J. L. A., 747. 1065
Weber, G., 446, 772, 847, 997
Weber, H. H. 938, 1065
Weber, J. F., 927, 1003
Webster, G. C, 662, 841, 1063
Weed, L. A., 974, 1000
Weed, R. I., 902, 903, 904, 905, 906, 907,
1065
Weeks, J. R., 212, 620, 622, 623, 624,
625, 627, 628, 630, 631, 1052, 1065,
1068
Weeks, T. E., 979, 1032
Wehrli, S., 953, 1070
Weibull, C, 548, 864, 1014
Weichel, E. J., 757, 1033
Weigert, M. G., 281, 282, 285, 1005
Weil-Malherbe, H., 75, 80, 95, 97, 379,
1065
Weiler, P., Jr., 335, 1052
Weill, C. E., 658, 660, 662, 673, 674, 683,
684, 803, 833, 1049, 1065
Wein, J., 38, 40, 42, 240, 1018
Weinbach, E. C, 174, 273, 349, 704, 843,
882, 987, 1010, 1065
Weinberger, R., 926, 927, 993
Weiner, I. M., 917, 923, 924, 929, 931,
932, 933, 934, 935, 1032, 1040, 1065
Weinhouse, S., 137, 141, 149, 151, 152,
178, 273, 274, 1023, 1040, 1065
Weisman, T. H., 195, 727, 972, 973,
1033
Weiss, B., 128, 176, 1065
1122
AUTHOR INDEX
Weiss, L., 614, 1066
Weissbach, A., 413, 855, 1021
Weissbach, H., 310, 103S
Weisz-Tabori, E., 597, 1042
Weitzel, G., 776, 852, 875, 881, 884, 1065
Welch, A. D., 261, 582, 583, 1041, 1065
Welch, K., 209, 1065
Wellman, H., 798, 869, 1030
Wells, H. G., 920, 1065
Wells, I. C, 290, 1065
Welt, L. G., 925, 1065
Wenger, B. S., 773, 817, 1041
Wenger, H. C, 315, 1057
Wenner, C. E., 125, 273, 274, 396, 397,
1065
Wenzel, D. b., 216, 1065
Wenzel, F., 15, 1004
Werkheiser, W. C, 582. 583, 584, 1066
Werkman, C. H., 26, 78, 79, 164, 187,
242, 430, 852. 987, 991, 996, 1065, 1068
Werle, E., 352, 1066
Wessels, J. G. H., 53, 74, 78, 79, 81, 104,
1066
Wessels J. S. C, 445, 548, 864, 891, 892,
1066
Wesson, L. G., 921, 1066
Westcott, W. L., 592, 1003
Westerfeld, W. W., 287, 614, 783, 814,
859, 1004, 1018, 1066
Westennan, M. P., 834, 1006
Westermann, E., 314, 315, 937, 1029,
1066
Westead, E. W., 351, 1066
Westheimer, F. H., 5, 1066
Westlake, D. W. S., 854, 1066
Westling, H., 363, 1032
Westmark, G. W., 325, 1010
Westveer, W. M., 619, 632, 633, 1067
Wetter, L. R., 599, 706, 709, 841, 1011,
1043
Whatley, F. E., 892, 989
Wheatley, A. H. M., 2, 20, 31, 32, 34,
52, 55, 113, 115, 144, 1046
Whetham, M. D., 2, 18, 35, 228, 1046
White, A. G. C, 504, 516, 528, 530, 1068
White, F. G., 847, 1066
White, H. L., 937, 1066
White, H. S., 524, 1052
White, I. G., 77, 1052
Whitehead, B. K., 422, 1019
Whitehouse, M. W., 150, 1066
Whiteley, H. R., 585, 675, 783, 847,
1020, 1033, 1066
Whitley, R. W., 459, 998
Whitmore, F. C., 745, 1066
Whittaker, V. P., 662, 675, 685, 836, 854,
1039. 1060
Whittingham, C. P., 891, 1066
Wick, A. N., 78, 187, 218, 219, 228, 232,
233, 234, 387, 390, 392, 393, 394, 398,
399, 403, 404, 410, 435, 839, 996, 999,
1037, 1040, 1052, 1066
Wickson-Ginzburg, M., 435, 1066
Wiebelhaus, V. D., 377, 1030
Wieland, O., 614, 841, 993, 1066
Wieland, T., 122, 814, 861, 1015
Wien, R., 952, 956, 957, 1066
Wiesmeyer, A., 415, 416, 780, 1066
Wiethoff, E. O., 518, 1066
Wight, K.. 392, 400, 401, 1030
Wigler, P. W., 274, 277, 278, 1066
Wilbrandt, W., 262, 900, 906, 907, 1049,
1066
Wilbur, K. M., 54, 174, 1023
Wilcox, P. E., 739, 759, 1015
Wilcox, S. S., 410, 839, 1037
Wilde, C. E., 836, 1066
Wilde. W. S., 924, 1062
Wilder, V., 878, 883, 991
Wildman, S. G., 58, 170, 191, 297, 995
Wiley, C. E., 461, 462, 712, 1008
Wilkin, G. D., 77, 78, 1019
Wilkinson, J. H., 433, 436, 1006, 1045
Will, J. J., 574, 1040
Will, L. W., 504, 505, 1016
Willard, H. H., 15, 1066
Williams, A. D., 585, 1066
Williams, A. K., 389, 1066
Williams, C. H., Jr., 817, 850, 1066
Williams, C. M., 21, 29, 199, 1050, 1051
Williams, D. C., 428, 995
Williams, D. L., 1031
Williams, G. R., 709, 994
Williams, H. L., 567, 568, 990
Williams, J. N. Jr., 36, 37, 41, 240, 288,
503, 509, 513, 1004, 1007, 1066
AUTHOR INDEX
1123
Williams, K., 428, 444, 684, 995, 1004
Williams, R. H., 391, 1057
Williams, R. J. P., 658, 770, 1062, 1067
Williams, R. T., 225. 427, 630, 990, 1056
Williamson, D. H., 550, 662, 711, 855,
1003, 1021
Williamson, S., 335, 989
Willing, F.. 499, 1067
Wills, E. D., 676, 686, 694, 710, 719.
1067
Wilson, A. N., 518, 1067
Wilson, B. R., 224, 236, 1003
Wilson, G. B., 197, 1067
Wilson, J. B., 168, 1011
Wilson, J. E., 520, 1067
W^ilson, L., 695, 990
Wilson, L. Cx.. 543, 554. 989
Wilson, P. W., 17, 26, 53, 291, 292, 293,
294, 854, 996, 997, 1032, 1038, 1047,
1053, 1066, 1067
Wilson, R. H., 601, 995
Wilson, T. G. G., 292, 294, 404, 1067
Wilson, T. H., 263, 265, 387, 394, 403,
1015, 1067
Wilson, W. L., 965, 1017
Winder, F. G., 77, 78, 1019
Winer, A. D., 433, 435, 436. 783, 787.
845, 846, 1042, 1067
Wingerson. F., 358, 1002
Wingo. W. J.. 328, 1067
Winnick. J.. 156, 684, 685, 687, 1011, 1014
Winestock, C. H., 539. 543, 1067
Winzler, R. J., 585, 1066
Winzor, D. J., 682, 1067
Wiseman, G., 265, 1067
Wiseman, M. H., 544, 1067
Wiskich, J. T., 20, 27, 63, 1067
Wislicenus, J., 1067
Withycombe, W. A., 436, 1006
Witkop, B., 316, 611, 1002, 1062
Witlin, B., 690, 1011
Witter, A., 641, 784, 1067
Witter, R. F., 143, 1067
WMxon, R. L., 708, 1024
Wlodawer, P., 887, 1067
Wockel, W., 923, 1067
Wohl, A., 259, 421, 1067
Wojtczak, L., 887, 1067
Wolbach, R. A., 756, 1048, 1067
Wold, F., 409, 803, 1020, 1067
Wolf, P. A., 619, 632, 633, 1067
Wolfe, J. B., 78. 145, 187, 218, 219, 228,
230, 231, 232, 233, 234, 387, 390, 399,
407, 1040, 1066, 1067
Wolfe, R. G., 802, 1067
Wolfe, S. J., 522, 530, 1067
Wolif, H. G., 165, 1061
Wolff, J. B., 833, 1067
Wood, H. G., 164, 224, 235, 852, 1054,
1057, 1067, 1068
Wood, J. D., 676. 1068
Wood. J. G., 834, 1054
Wood, N. P., 676, 693, 1004
Wood, R. C. 584, 1068
Wood, W. A., 885, 1028
Woodbury, R. A., 921, 1011
Woodruff, L. L., 981, 1068
Woods, D. D., 260, 836, 988, 1068
Woods, L. A., 620, 622, 623, 624, 625,
627, 628, 629, 630, 631, 1052, 1053,
1068
Woodward. G. E., 385, 386, 389. 391,
392, 689, 745, 1001, 1020, 1051, 1066,
1068
Woody, B. R., 26, 1068
Wooldridge, W. R., 2, 21, 26, 35, 36,
40, 62, 152, 237, 1046
Woollen, J. W., 419, 420, 1064
Woolley, D. W., 259, 260, 261, 281, 282,
489, 504, 516, 518, 519, 522, 523, 528,
529, 530. 533, 537, 538, 589, 1053,
1068
Wooltorton, L. S. C, 120, 1023
Worgan, J. T., 427, 1032
Work, E., 261, 593, 708, 1003, 1068
Work, T. S., 261, 1068
Wormser, E. H., 357, 1068
Woronkow, S., 207, 1033
Wortman, B., 707, 816, 1068
Wosilait, W. D., 713, 1068
Wright. C. I., 545, 549, 559, 1068
Wright, E. M., 387, 991
Wright, G. F., 744, 1054
Wright, L. D., 264, 993
Wright, N. C, 658, 1068
Wriston, J. C, Jr., 160, 1068
1124
AUTHOR INDEX
Wu, H. L. C, 600, 1068
Wu, L. C, 27, 1068
Wurtz, E., 538, 1006
Wyatt, A., 574, 1046
Wyatt, H. v., 453, 1068
Wylie, D. W., 611, 612, 1068
Wyman, J., 638, 1005
Wyngaarden, J. B., 281, 282, 467, 474,
480, 1054, 1068
Wyngarden, L., 585, 1059
Wynn, J., 602, 1068
Wyss, 0., 690, 1068
Yoshimatsu, H., 578, 676, 858, 1064
Yoshimura, J., 868, 939, 940, 1061
Young, B. G., 459, 1039
Young, G. A., 358, 1002
Young, L. C. T., 80, 119, 120, 1000
Young, P., 15, 1066
Young, R. H., 225, 226, 227, 228, 232,
1053, 1069
Younger, F., 569, 1048
Yphantis, D. A., 64, 334, 1023
Yushok, W. D., 381, 382, 385, 388, 395,
396, 397, 1033, 1069
Yagi, K., 340, 347, 537, 541, 544, 772,
1005, 1060, 1068, 1069
Yagi, T., 551, 555, 1022
Yall, I.; 237, 1013
Yamada, E. W., 60, 1069
Yamada, H., 543, 554, 1059
Yamada, K., 845, 1069
Yamada, T., 880, 973, 975, 1069
Yamafuji, K., 675, 1065
Yamamura, Y., 26, 36, 62, 63, 148, 228,
855, 1029, 1069
Yamane, T., 739, 741, 1004, 1008, 1069
Yamashita, J., 844, 1069
Yamauchi, M., 224, 1069
Yanagita, T., 880, 973, 975, 1069
Yanari, S., 367, 1069
Yang, W. C. T., 877, 1069
Yanofsky, C, 321, 1012
Yashimatsu, H., 676, 1022
Yates, J. R., 657, 1019
Yeas, M. F., 175, 181, 1069
Yee, R. B., 90, 1069
Yonetani, T., 508, 779, 780, 785, 1069
Yoneya, T., 547, 549, 1069
Yoneyama, T., 163, 888, 1024, 1069
York, J. L., 842, 996
Yoshiba, A., 163, 1069
Yoshida, F., 662, 687, 1069
Yoshida, H., 151, 184, 1069
Yoshihara, I., 194, 999
Yoshikawa, H., 817, 1017
Yoshikawa, M., 859, 1058
Zahl, P. A., 585, 1069
Zakrzewski, S. F., 582, 583, 1069
Zalik, S., 122, 1049
Zalusky, R., 387, 1052
Zambonelli, C, 984, 985, 1069
Zamcheek, N., 208, 1063
Zamecnik, P. C, 156, 887, 1057
Zannoni, V. G., 272, 306, 595, 1029, 1069
Zarnitz, M. L., 810, 1064
Zarudnaya, K., 61, 64, 1058
Zatman, L. J., 485, 487, 490, 496, 498,
504, 1069
Zborowski, J., 887, 1067
Zeliteh, I., 61, 438, 439, 782, 842, 1069
Zeller, A., 335, 1005
Zeller, E. A., 60, 338, 348, 362, 365,
1089, 1070
Zellweger, H., 953, 1070
Zewe, v., 376, 1010
Zhanley, J. C, 854, 1046
Ziegler, D. M., 18, 1070
Ziff, M., 684, 691, 692, 1070
Zimmerman, A. M., 964, 965, 1029, 1070
Zimmerman, S. B., 446, 1070
Zink, M. W., 845, 1051
Zipkin, I., 630, 631, 1070
Zittle, C. A., 475, 1070
Zollner, N., 18, 30, 461, 462, 1033, 1070
Zubrod, C. G., 538, 1030
Zygmunt, W. A., 359, 1070
Zweifach, B. W., 879, 895, 998
SUBJECT INDEX
When a substance is named in connection with an enzyme or process, the effect
of this substance on the enzyme or process, usually an inhibition, is designated.
Accumulation of substances, see specific
substances accumulated
Acetaldehyde, p\Tuvate decarboxylase,
432, 600
Acetaldehyde dehydrogenase, o-iodoso-
benzoate, 705
Acetamide,
A'-acetyl-/S-glucosaminidase, 419
pyruvate decarboxylase, 430
urease, 603
2-Acetamido-2-deoxygalactonolactone,
xV-acetyl-yS-galactosaminidase, 420
A'^-acetyl'/^-glucosaminidase, 419
2-Acetamido-2-deoxygluconolactone,
iV-acetyl-/3-glucosaminidase, 419
carcinostasis, 428
6-Acetaniidolevulinate, aminolevulinate
dehydrase, 592
Acetanilide deacetylase, ferricyanide, 674,
Acetate,
accumulation of, benzoate, 349
iV^-acetyl-/?-glucosaminidase, 419
glutamate decarboxylase, 328
glyoxylate transacetase, 594
incorporation into lipid,
malonate, 146-149
mercurials, 887
propionate, 613-614
kynurenine:a-ketoglutarate transami-
nase, 608
lactate dehydrogenase, 436
metabolism in rabbit, 2-deoxyglucose,
399
metaboUsm of,
fraris-cyclopentane-l,2-dicarbocylate,
241
6-deoxy-6-fluoroglucose, 404
propionate, 613-614
oxidation of, see also Respiration (ace-
tate)
2-deoxyglucose, 397
malonate, 77-78
malonic diethyl ester, 236-237
mercurials, 878, 883
pantoate:/?-alanine ligase, 597
tyrosinase, 300-301
Acetate kinase (acetokinase),
o-iodosobenzoate, 705
mercurials, 830
Acetoacetate,
accumulation of, tartronate, 237
formation of,
from butyrate, 613
from malonate, 234
malonate, 138-144
pathways, 139
)?-hydroxybutyrate dehydrogenase, 594
metabolism of, malonate, 144
Acetoacetate carboxy-lyase, see Acetoace-
tate decarboxylase
Acetoacetate decarboxylase,
acetopyruvate, 591
mercurials, 830
Acetoacetyl-CoA, splitting by mercurials,
751
Acetobacter
ethanol oxidation, mercurials, 898
tartronate occurrence in, 225
Acetobacter melanogenum, pyruvate oxi-
dation,
mercurials, 878
Acetobacter pasteurianutn, succinate oxi-
dation,
malonate, 52
1125
1126
SUBJECT INDEX
Acetobacter xylinum., cellulose synthesis,
malonate, 132
Acetoin, formation from pyruvate,
oxythiamine-PP, 519
phenylpyruvate, 430
Acetokinase, see Acetate kinase
Acetopyruvate,
acetoacetate decarboxylase, 591
Acetylation, folate analogs, 586-587
Acetylcholine,
biosynthesis of, malonate, 165-166
cardiac response to,
malonate, 217
mercurials, 946-947
ganglionic response to, mercurials, 949
intestinal response to, hydrogen pe-
roxide, 696
Acetylcholinesterase, see Cholinesterase
Acetyl-CoA,
formation of, propionate, 613
splitting by mercurials, 751
Acetyl-CoA carboxylase,
acyl-CoA analogs, 614
mercurials, 830
Acetyl-CoA kinase, mercurials, 830
Acetyl-CoA synthetase, propionate, 613
Acetylene-carboxylate hydrase, malon-
ate, 60
Acetylene-dicarboxylate,
intercharge distance, 6
pyruvate oxidation, 240-241
succinate dehydrogenase, 34, 36, 38,
240-241
iV-Acetylgalactosamine,
iV-acetyl-/^-galactosaminidase, 419-420
A'^-acetyl-/3-glucosaminidase, 419
a-galactosidase, 417
/?-galactosidase, 419
iV^-Acetyl-/5-galactosaminidase, analogs,
419-420, 429
iV-Acetylglucosamine,
iV-acetyl-/3-halactosaminidase, 419-420
iV-acetyl-/S-glucosaminidase, 419
a-galactosidase, 417
jS-galactosidase, 418
glucokinase, 390
glucosamine kinase, 593
glucosamine phosphorylation, 382
glycogen formation, 382
hexokinase, 381-382, 390
iV-Acetylglucosamine-6-phosphate, glu-
cose-6-P dehydrogenase, 411
iV-Acetyl-^-glucosaminidase, analogs, 419
429
iV-Acetylglucosaminolactone, iV-acetyl-/S-
glucosaminidase, 429
Acetylindoxyl oxidase, analogs, 591
iV-Acetylisatin, acetylindoxyl oxidase,
591
iV-Acetylleucine, leucine decarboxylase,
352
Acetyllipoate, biosynthesis of,
lipoate analogs, 590
Acetyl-D-phenylalaninamide, chymotryp-
sin, 372
Acetyl-L-phenylalanine, cathepsin C, 375
Acetyl-D-phenylalanine methyl ester, chy-
motrypsin, 372
Acetylphosphatase, o-iodosobenzoate, 705
3-Acetylpyridine,
alcohol dehydrogenase, 498
central nervous system, 499
chick embryo, 494
heart,
conduction, 494
contractility, 499-500
membrane potentials, 499-500
Lactobacillus growth, 494
lethal doses, 499
mechanisms of action, 494
metabolism of, 494-495
iV-methylnicotinamide formation from,
495
NAD levels in tissues, 495-496
NAD metabolism, 489-500
NAD nucleosidase, 491, 498
nicotinamide deamidase, 512
nicotinamide deaminase, 498
nicotinate deficiency, 489
toxicity, 489, 494, 499
4-Acetylpyridine, toxicity, 499
3-Acetylpyridine-NAD,
brain levels of, 499
formation of 496
function as coenzyme, 497
glucose-6-P dehydrogenase, 497
SUBJECT INDEX
1127
NADH:ferricyanide oxidoreductase,
510
nicotinamide deamidase, 512
0-Acetylthiamine,
pyruvate oxidation, 519
structure of, 517
thiamine kinase, 523
iV-Acetylthyroxine, thyroxine deiodinase,
602
Acetyltryptamine, chymotrypsin, 371
Acetyltryptophan,
chymotrypsin, 374
L-tryptophan-sRNA ligase (AMP), 326
Acetyl-D-tryptophanamide, chymotryp-
sin, 371
Acetyl tryptophanates, chymotrypsin, 371,
373
Acetyltryptophanmethylamides, chymo-
trypsin, 371
Acetyl-D-tyrosinamide, chymotrypsin, 371
Acetyl-L-tyrosinate, chymotrypsin, 371
Acetyl-L-tyrosine,
carboxypeptidase, 367
tyrosinase, 304-305
Ace tyl-D- tyrosine ethyl ester, chymo-
trypsin, 371
Acetyl - D - tyrosinehydroxamide, chymo-
trypsin, 371
Acetyl - L - tyrosinemethylamide, chymo-
trypsin, 371
Achro7nobacter, nitrogen fixation,
oxygen, 292
Achromobacter fischeri,
luminescence, mercurials, 888-891
mercurial uptake, 897
respiration (glucose), mercurials, 889-
891
SH groups in, 897
Achromobacter guttatum,
malonate occurrence in, 225
respiration (endogenous), malonate, 168
Acid phosphatase, see Phosphatase (acid)
Aconitase,
frans-aconitate, 272-273
ferricyanide, 674
hydrogen peroxide, 692, 694
y-hydroxy-a-ketoglutarate, 615-616
malonate, 60
mercurials, 830
propane-tricarboxylate, 240
cis-Aconitate,
glutamate decarboxylase, 328
oxalosuccinate decarboxylase, 597
oxidation of, malonate, 79
phosphofructokinase, 385
(rans-Aconitate,
aconitase, 272-273
citrate accumulation, 273-274
fatty acid synthesis, 273
fumarase, 273, 275-276, 279
glutamate decarboxylase, 328
glutamate dehydrogenase, 331
ion transport in barley roots, 274
metabolism of, 274
occurrence in plants, 274
paramecia, 274
respiration, 273-274
Venturia ascosporulation, 195-196
Aconitate hydratase, see Aconitase
Acridines, chymotrypsin, 373
Acriflavine (Trypaflavine, Euflavine),
structure of, 537
Acrodynia,
deoxypyridoxol, 566
mercurials, 953-954
Actin,
association with myosin, o-iodosoben-
zoate, 723
ATP binding, mercurials, 938-940
Ca++ binding,
o-iodosobenzoate, 723
mercurials, 939-940
o-iodosobenzoate, 704-723
polymerization of, mercurials, 938-940
Active transport, see also specific organ-
isms or tissues
analogs, 261-268
malonate, 203-210
mercurials, 205, 907-921, 928
phlorizin, 205
pyridoxal analogs, 574-575
Actomyosin,
hydrogen peroxide, 691
o-iodosobenzoate, 723
mercurials, 938-940
Acylase, mercurials, 831
1128
SUBJECT INDEX
iV-Acylated glucosamines,
carcinostasis, 382
hexokinases, 381-382
Acyl-CoA dehydrogenase, crotonyl-CoA,
591
Acyl 5'-nucleotidase, AMP, 466
Acyl transfer, lipoate analogs, 590
Adenine,
adenosine hydrolase, 466
adenylosuccinate synthetase, 467
D-amino acid oxidase, 545
ATPase, 445
glucose dehydrogenase, 501
inosine phosphorylase, 471
malate dehydrogenase, 509, 513
NAD nucleosidase, 488-490, 492-493
nicotinate deficiency, 513
5'-nucleotidase, 472
pyridoxal kinase, 475, 477
pyrophosphatase, 475
xanthine oxidase, 282
Adenine aminohydrolase, see Adenine
deaminase
Adenine deaminase, analogs, 466
Adenocarcinoma,
dehydrogenase inhibition in vivo, 6-
aminonicotinamide, 505
growth of,
6-aminonicotinamide, 505
riboflavin analogs, 538
pyridoxine levels in, deoxypyridoxol,
568
Adenosinase,
hydrogen peroxide, 692
mercurials, 831
Adenosine,
adenylosuccinate lyase, 466
adenylosuccinate synthetase, 467
alcohol dehydrogenase, 506, 508
D-amino acid oxidase, 545
ATPase, 445
creatine kinase, 446
glutamate dehydrogenase, 508
glutamate semialdehyde reductase, 507
lactate dehydrogenase, 501
malate dehydrogenase, 509, 513
NAD kinase, 509
NADH pyrophosphatase, 506, 511
NAD nucleosidase, 488, 490, 492
5 '-nucleosidase, 472
phosphodiesterase, 473
pyridoxal kinase, 475, 477
pyrophosphatase, 475
thiamine kinase, 475
Adenosine aminohydrolase, see Adeno-
sine deaminase
Adenosine deaminase, analogs, 466, 477
Adenosinediphosphate (ADP),
adenylosuccinate synthetase, 467
alcohol dehydrogenase, 506, 508
D-amino acid oxidase, 545
AMP-ATP transphosphorylase, 446
arginine kinase, 467
ATPase, 444-445
ATP-P, exchange reaction, 444
creatine kinase, 447
deoxycytidylate kinase, 469
fructose- 1,6-disphosphatase, 470
glutamate semialdehyde reductase, 507
glutamine synthetase, 471
GTPase, 446
hexokinase, 383
isocitrate dehydrogenase, 509
ITPase, 446
malate dehydrogenase, 509
NADH:menadione oxidoreductase, 510
NADH oxidase, 506, 511
NADH pyrophosphatase, 506, 511
NAD kinase, 506, 510
NAD:NADP transhydrogenase, 507,
510
NAD nucleosidase, 490, 492-493
phosphodiesterase, 473
phosphoribosyl-PP amidotransferase,
474
pyridoxal kinase, 475, 477
pyrophosphatase, 475
thiamine kinase, 475
UTPase, 446
yeast levels, mercurials, 885
Adenosine hydrolase, adenine, 466
Adenosinemonophosphate (adenosine-5'-
phosphate, adenylate, AMP),
acyl 5 '-nucleotidase, 466
adenylosuccinate lyase, 466
adenylosuccinate synthetase, 467
SUBJECT INDEX
1129
alcohol dehydrogenase, 506, 508
D-amino acid oxidase, 545
arginine kinase, 467
deoxycytidylate deaminase, 469
flavokinase, 545
fructose- 1,6-diphosphatase, 470
glucose-6-P dehydrogenase, 508
glutamate dehydrogenase, 508
glutamate semialdehyde reductase, 507
guanosine-5'-P reductase, 471
isocitrate dehydrogenase, 509
malate dehydrogenase, 509
NADH:menadione oxidoreductase, 510
NADH oxidase, 511
NADH pyrophosphatase, 506, 511
NAD kinase, 509-510
NAD:NADP transhydrogenase, 507,
510
NAD nucleosidase, 492
NADPH:cytochrome c oxidoreductase,
511
NADPH : glutathione oxidoreductase,
512
phosphatase, 439
phosphodiesterase, 473
pyridoxal kinase, 475, 477
pyrophosphatase, 475
ribonuclease, 475
thiamine kinase, 475
yeast levels, mercurials, 885
Adenosinemonophosphate - 3' - phosphate,
phenol sulfokinase, 473
Adenosinemonosulfate, AMP-ATP trans-
phosphorylase, 446
Adenosine-2 '-phosphate, NAD nucleosi-
dase, 490
Adenosine-3 '-phosphate, lactate dehydro-
genase, 501
Adenosine-5'-phosphate, see Adenosine-
monosphosphate
Adenosine-5'-phosphsulfate reductase, io-
dine, 684
Adenosinetriphosphatase (ATPase),
ADP inhibition,
effect of Ca++ and Mg++, 445
effect of pH, 445
analogs, 444-447
Ca++, 453
cystine, 663-664
dehydroacetate, 623
dithioglycolate, 663-664
ferricyanide, 673
hydrogen peroxide, 691-692
iodine, 682, 684
o-iodosobenzoate, 705
mercurials, 789, 863-869, 876, 905, 912
aggregation, 789
Ca++ and Mg++ binding, 868
configurational changes, 868
effects of Ca++ and Mg++, 798
effects of 2,4-dinitrophenol, 869
effects of pH, 792, 867
initial phosphate release, 867-868
kinetics, 869
number of Hg++ ions required, 912
potentiation by substrate, 807
protection by Mg++, 780
rates of SH group reaction, 814
relation to SH groups, 803-804,
806-807, 868-869
stimulation, 815-816, 819-821
porphyrindin, 668
quinacrine, 547-548, 556
Adenosinetriphosphate (ATP),
adenylosuccinate lyase, 466
adenylosuccinate synthetase, 467
D-amino acid oxidase, 545
arginine kinase, 467
aspartate carbamyltransferase, 468
fructose- 1,6-diphosphatase, 470
glutamate semialdehyde reductase, 507
GMP reductase, 471
IMP dehydrogenase, 471
isocitrate dehydrogenase, 509
lactate dehydrogenase, 501
levels in nuclei,
dehydroacetate, 624
malonate, 189
levels in tissues,
aminopterin, 585
2-deoxyglucose, 394-396
o-iodosobenzoate, 721-722
malonate, 157-158
mercurials, 876
quinacrine, 560
levels in tumors,
1130
SUBJECT INDEX
2-deoxyglucose, 395
D-glucosamine, 383
levels in yeast, mercurials, 884-885
malate dehydrogenase, 509, 513
NADH:menadione oxidoreductase, 510
NADH oxidase, 511
NADH pyrophosphatase, 511
NAD:NADP transhydrogenase, 510
NAD nucleosidase, 492
phosphatase, 439
phosphodiesterase, 473
phosphoglucose isomerase, 474
phosphoribosyl - PP amidotransferase,
474
polynucleotide phosphorylase, 474
pyrophosphatase, 475
succinyl-CoA deacylase, 475
Adenovirus type 5, infectivity of,
mercurials, 977
Adenylate, see Adenosinemonophosphate
Adenylate deaminase,
ferricyanide, 674
hydrogen peroxide, 692
mercurials, 774, 831
Adenylate kinase (myokinase, ATP:AMP
phosphotransferase ) ,
adenosinemonosulfate, 446
ADP, 446
mercurials, 772, 847, 860
Adenylmethylenediphosphonate, ATPase,
445-446
Adenylmethylphosphonate, polynucleo-
tide phosphorylase, 474
Adenylosuccinate lyase,
analogs, 466, 481
mercurials, 831
Adenylosuccinate synthetase, analogs,
467
Adipate,
aspartate - a - ketoglutarate transami-
nase, 334
carcinostasis, 201
fumarase, 275
glutamate decarboxylase, 328
glutamate dehydrogenase, 330, 332
intercharge distance, 6
ionization constants, 8
kynurenine:a-ketoglutarate transami-
nase, 595, 607-609
lethal dose, 201
pyridoxamine : oxalacetate transami-
nase, 600
succinate dehydrogenase, 35
urinary citrate, 109
Adipose tissue, seealso Epididymal fat pad
fructose metabolism, 2-deoxyglucose,
391
glucose metabolism,
6-deoxy-6-fluoroglucose, 404
2-deoxyglucose, 391
6-deoxyglucose, 403
glycolysis, 2-deoxyglucose, 392
methylmalonate occurrence in, 224
respiration (endogenous), malonate, 178
respiratory quotient, malonate, 185
ADP, see Adenosinediphosphate
Adrenals,
catecholamine release, mercurials, 947
corticosteroid synthesis, malonate, 150
mercurial levels in, 959
Aedes aegypti,
a -ketoglutarate oxidation, malonate, 80
succinate dehydrogenase, malonate, 29
Aerobacter aerogenes,
growth of, dehydroacetate, 632
malonate metabolism in, 227-229
respiration, mercurials, 984
Aerobacter indologenes, citrate oxidation,
malonate, 78
Aerobic glycolysis, see Glycolysis (aero-
bic)
A-esterase, iodine, 684
Agmatine,
cadaverine oxidation, 363
structure of, 361
Agrobacterium turnefaciens, growth of,
tungstate, 615
Alanine, phosphatase (acid), 441
/?- Alanine, zl i-pyrroline-5-carboxylate de-
hydrogenase, 336
D -Alanine,
L-alanine dehydrogenase, 354
L-amino acid oxidase, 340
Bacillus cereus germination induced by
L-alanine, 270
dipeptidase, 368
SUBJECT INDEX
1131
DL-Alanine,
L-amino acid oxidase, 340
phenylalanine hydroxylase, 354
L-Alanine, arginase, 337
L-Alanine dehydrogenase,
D-alanine, 270
analogs, 354
ferricyanide, 674
o-iodosobenzoate, 705
malonate, 60
mercurials, 831
Alanine:a-ketoglutarate transaminase, see
Transaminases
Alanine:pyriivate transaminase, see Trans-
aminases
Alanine racemase, pyridoxal analogs, 575
D-Alanyl-D-alanine synthetase, D-cyclo-
serine, 360
Alcaligenes faecalis,
growth of, dehydroacetate, 632
succinate oxidation, malonate, 52
Alcohol, see Ethanol
Alcohol dehydrogenase,
6-aminonicotinamide-NAD, 505
o-iodosobenzoate, 705, 714-715, 717
protection by ethanol, 717
protection by NAD, 717
reversal by GSH, 718
titration of, 714-715
mercurials, 784-785, 788-789, 792, 803-
806, 810-812, 825, 831
coenzyme displacement, 784-785
NADH binding, 788
pH effects, 792
protection by ethanol, 779-780
protection by NAD, 779-780
rate of inhibition, 810-812
relation to SH groups, 803-806
reversal with GSH, 825
rotatory dispersion changes, 788
sphtting into subunits, 789
temperature effects, 810-811
NAD analogs, 513-514
nicotinyl-hydrazide-NAD, 497
nucleotides, 506-508
porphyrexide, 668
pyridine derivatives, 498
pyridine-3-sulfonate, 504
quinacrine, 548
thionicotinamide-NAD, 497
Aldehyde dehydrogenase, see also Acetal-
dehyde dehydrogenase
o-iodosobenzoate, 704-706, 717
protection by NAD, 717
mercurials, 780, 832
protection by NAD(P), 780
Aldehyde:NAD oxidoreductase, GSSG,
662
Aldehyde oxidase,
mercurials, 772, 803
relation to SH groups, 803
type of inhibition, 772
quinacrine, 547, 549
Aldolase,
analogs, 407-408
ferricyanide, 673-674, 677
p-fluorophenylalanine incorporation in-
to, 351
hydrogen peroxide, 692
iodine, 682, 684
mercurials, 643, 649, 780, 788, 803,
832-833
protection by fructose- 1,6-diP, 780
rate of SH group reaction, 812
relation to SH groups, 803
susceptibility to trypsin, 788
p-mercuribenzoate, 643, 649
SH groups of,
pk„'s of, 638
titration with p-mercuribenzoate,
643, 649
sulfate, 414
Aldose 1-epimerase, see Mutarotase
Alfalfa, malonate occurrence in, 224
Alginate,
phosphatase (acid), 443, 464
ribonuclease, 462
Aliesterase, quinacrine, 547, 549
A'-Alkyl-2-amino-2-methylpropanediols,
choline oxidase, 290
iV-Alkyl-2-amino-2-methylpropanols, cho-
line oxidase, 290
iV-Alkyl-3-aminopropanols, choline oxi-
dase, 290
iV-Alkylethanolamines, choline oxidase,
290
1132
SUBJECT INDEX
Alkylmalonates, succinate dehydrogen-
ase, 37-38, 40
Alkylnormorphines, morphine iV-deme-
thylase, 590, 604
AlHinase (alHin lyase), o-iodosobenzoate,
706
Alhin lyase, see Alliinase
Allohydroxy - d - proline oxidase, quina-
crine, 547, 549
Allomyces macrogynus,
respiration (endogenous), malonate, 168
succinate dehydrogenase, malonate, 27
Allopurinol (Zyloprim),
urinary excretion of xanthines, 283
xanthine oxidase, 283
Allose, fructokinase, 376-377
Allose kinase, mercurials, 833
Allose-6-phosphate,
hexokinase, 379-380
structure of. 378
Allothreonine aldolase, o-iodosobenzoate,
706
Alloxazine, L-amino acid oxidase, 541
Allylglycine, DL-methionine uptake by
ascites cells, 265
iV^-Allylnormorphine, see Nalorphine
Alternaria solani, growth of,
dehydroacetate, 633
Altrose, fructokinase, 376
Altrose-6-phosphate, hexokinase, 380
Ameboid movement, mercurials, 982
Amethopterin (Methotrexate),
acetyl transfer in liver, 586
dihydro folate reductase, 581-584
folate deficiency, 581
formate incorporation into proteins, 585
purine biosynthesis, 585
resistance to, 583
structure of, 580
sulfonamide acetylation, 586
tetrahydrofolate biosynthesis, 581
tissue distribution, 583
Amidines,
diamine oxidase, 360-365
structures of, 361
Amidophosphoribosyltransferase, see
Phosphoribosylpyrophosphate amido-
transferase
Amidosulfonate, see Sulfamate
Amine oxidase, see Monoamine oxidase
Aminoacetone synthetase, ferricyanide,
674
Amino acid activating enzymes, see spe-
cific L-amino acid:sRNA ligases
Amino acid decarboxylase, see also spe-
cific enzymes
analogs, 317
mercurials, 780
protection by pyridoxal-P, 780
protection by substrates, 780
a-methyl-w -tyrosine in vivo, 315-317
permanganate, 660
D-Amino acid oxidase,
adenine analogs, 545
amino acid analogs, 340-348
analogs, 347-348, 545
competition with FAD, 347-348
competition with substrates, 347-
348
FAD complexes, 347-348
benzoates, 341-348
o-iodosobenzoate, 704, 706, 717-718
protection by FAD, 717
protection by substrates, 717
reversal by cysteine, 718
kojic acid, 349-350
maleate, 343
malonate, 60
mercurials, 780, 804
protection by alanine, 780
protection by FAD, 780
relation to SH groups, 804
p-mercuribenzoate, competitive nature
of inhibition, 771-772
nucleotides, 545
quinacrine, 547, 549, 557, 560
riboflavin analogs, 540-541, 544
L-Amino acid oxidase,
analogs, 338-340
pH effects, 340
benzoate, 348
D-leucine, 268
malonate, 60
D-phenylalanine, 268
quinacrine, 547, 549
riboflavin analogs, 540-542
SUBJECT INDEX
1133
Amino acid-sRNA ligase, stimulation by
mercurials, 816
Amino acids,
active transport of, p>Tidoxal analogs,
574-575
deamination of, D-alanine, 270
liver levels of, mercurials, 954
metabolism of, aminosulfonates, 350
oxidation by hypochlorite. 658
oxidation by permanganate, 657
oxidation of,
kojic acid, 350
malonate, 151-154
synthesis from glucose, 2-deoxyglucose,
399
transport of, analogs, 264-267
Aminoacylase, o-iodosobenzoate, 706
2-Aminoadenine,
adenine deaminase, 466
adenosine deaminase, 466
2-Amino-8-azapurine, conversion to 8-
azaguanine, 281
7n-Aminobenzoate,
glucose dehydrogenase, 501
lactate dehydrogenase, 501
p-Aminobenzoate,
acetylation of, benzoate, 349
D-amino acid oxidase, 341
glucose dehydrogenase, 501
lactate dehydrogenase, 501
transport in Flavobacterium, p-ami-
nosalicylate, 613
tyrosine: a-ketoglutarate transaminase,
306
Aminobenzoates, D-amino acid oxidase,
341, 348
3-o-Aminobenzyl-4-methylthiazole
(ABMT),
structure of, 517
thiaminase, 524-525
Aminobenzylthiazoles, thiaminase, 524-
525
DL-a-Aminobutyrate, homoserine kinase,
357
L-a-Aminobutyrate,
arginase, 337
leucine decarboxylase, 352
y-Aminobutyrate (GABA),
brain levels of,
aminooxyacetate, 358
pj-ridoxal analogs, 573-574
toxopyrimidine, 578
carboxypeptidase, 367
oxidation of, malonate, 154
A ^-pyrroline-5-carboxylate dehydrogen-
ase, 336
y-Aminobutyrate:a-ketoglutarate trans-
aminase, see Transaminases
f-Aminocaproate, carboxypeptidase, 367
a-Amino-/S-chlorobutyrate, valine incor-
poration into protein, 351
1-Aminocyclopentane-carboxylate,
glycine uptake by ascites cells, 265
uptake by ascites cells, 266
uptake by brain, 266
2-Amino-D-glucose, see D-Glucosamine
Aminoguanidine,
cadaverine oxidation, 363
diamine oxidase, 362-363
histamine metabolism in man, 363
histidine decarboxylase, 352
structure of, 361
p-Aminohippurate (PAH),
glycine iV-acylase, 355
renal transport of,
azide, 205
cyanide, 205
dehydroacetate, 205, 625-626
2,4-dinitrophenol, 205
fluoride, 205
fluoroacetate, 205
iodoacetate, 205
malonate, 203-205
mercurials, 205, 920-921, 928
phlorizin, 205
2-Amino-4-hydroxy-6-formylpteridine,
see Pterin-6-aldehyde
2-Amino-4-hydroxy-6-pteridyl aldehyde,
see Pterin-6-aldehyde
a-Aminoisobutyrate, leucine decarboxy-
lase, 352
^-Aminolevulinate dehydrase, analogs,
591-592
<5-Aminolevulinate synthetase, aminoma-
lonate, 238-239
Aminomalonate,
1134
SUBJECT INDEX
5-aminolevulinate sjoithetase, 238-239
condensation with aldehydes, 239
formation in tissues, 239
metabohsm of, 239
2-Amino-iV-methyladenine,
adenine deaminase, 466
inosine phosphorylase, 471
2- Amino - 2 - methyl - 1 - propanol, methio-
nine biosynthesis, 291
6-Aminonicotinamide, 504-505
ADP-ATP levels in tissues, 505
carcinostasis, 505
lactate oxidation, 504
lethal doses, 504
NAD analog of, 504-505
NAD levels in tissues, 505
respiration (endogenous), 504
sulfanilamide acetylase, 601
toxicity, 504
6-Aminonicotinamide-NAD, 504-505
alcohol dehydrogenase, 505
creatine kinase, 505
formation in tissues, 504-505
pyruvate kinase, 505
6-Aminonicotinate,
NAD nucleosidase, 488
toxicity, 505
Aminooxyacetate, 358-359
anticonvulsant activity, 358
central nervous system, 358
GABA : a-ketoglutarate transaminase,
358
GABA levels in brain, 358
oxime formation with pyridoxal-P, 358
pyridoxal deficiency, 358
pyridoxal kinase, 358-359
Aminopeptidases,
analogs, 367-368
cysteine, 663
ferricyanide, 674
GSH, 663
GSSG, 662
mercurials, 787, 833
displacement of Mn++, 787
TO-Aminophenol, D-amino acid oxidase,
348
p-Aminophenol, D-amino acid oxidase,
344
Aminophenols, tyrosinase, 304
p-Aminophenylalanine, L-phenylalanine:
sRNA hgase (AMP), 355
Aminopterin,
acetyl transfer, 586
ATP level in tissues, 585
dihydrofolate reductase, 581-584
folate deficiency, 581
glycine-serine inter con version, 585
NAD level in liver, 585
nucleic acid biosynthesis, 585
purine biosynthesis, 585
respiration, 585
structure of, 580
tetrahydrofolate synthesis, 581
uptake by Bacillus suhtilis, mercurials,
911
uptake by bacteria, 584
6-Aminopurimidine deoxyribonucleoside-
5 '-phosphate deaminase, o-iodosoben-
zoate, 706
Aminoquinolines, chymotrypsin, 373
p- Aminosalicylate,
D-amino acid oxidase, 348
p-aminohippurate transport, 613
protocatechuate transport, 267
sulfanilamide acetylase, 601
a-Aminosulfonates, amino acid metabo-
hsm, 350
Aminotransferases, ses Transaminases
3-Amino-L-tyrosine,
tyrosinase, 304-305
tyrosine: a-ketoglutarate transaminase,
306
f5-Aminovalerate,
carboxypeptidase, 367
A i-pyTroline-5-carboxylate dehydroge-
nase, 336
^-Aminovalerate : a - ketoglutarate trans-
aminase, see Transaminases
Amiphenazole, see 2,4-Diamino-5-phenyl-
thiazole
Ammonium chloride, renal refractoriness
to mercurials, 933-934
Ammonium ion,
L-amino acid oxidase, 338
formation in kidney,
benzamide, 348
SUBJECT INDEX
1135
benzoate, 348
phenylacetate, 348
jS-phenylpropionate, 348
glutaminase, 332
AMP, see Adenosineraonophosphate
Amphetamine (Benzedrine),
dopamine /3-hydroxylase, 320
phenylalanine /S-hydroxylase, 600
Amylamine, diamine oxidase, 361-362
Amylase, succinyl peroxide, 694
a-Amylase,
analogs, 420
bios3mthesis of, D-asparagine, 269
cystine, 662
dehydroacetate, 622
ferricyanide, 674
iodine, 683-684
macroions, 464
mercurials, 792, 795-797, 813, 833
pH effects, 792, 795-797
rate of inhibition, 813
nitrite, 660
permanganate, 659
/S-Amylase,
analogs, 421
cystine, 662
dichromate, 660
ferricyanide, 673-675
GSSG, 662
hydrogen peroxide, 692
iodine, 683-684
o-iodosobenzoate, 703, 706, 716, 718
failure to reverse, 718
kinetics, 716
pH effects, 716
macroions, 464
mercurials, 772, 803, 833
relation to SH groups, 803
type of inhibition, 772
nitrite, 660
permanganate, 659
Amylo- 1 ,6-glucosidase, o-iodosobenzoate,
706, 718
Amylomaltase (maltose 4-glucosyl trans-
ferase),
analogs, 415, 422
mercurials, protection by maltose, 780
Amylopectin sulfate, hyaluronidase, 459
Anaerobic glycolysis, see Glycolysis (an-
aerobic)
Analog inhibition,
annual number of publications, 260
development of concept, 259-261
expression of results, 252-255
kinetics, 248-252
mechanisms of, 246-248
membrane transport, 261-268
substrate isomers, 268-274
Analogs, see also specific substances
definition of different types of, 257
formed by fluorine substitution, 258-259
metabolism to active inhibitor, 247,
251-252, 258
molecular alterations producing, 255-
259
5a-Androstane-3,17-dione, J*-3-ketoste-
roid reductase, 449
Androst - 1 - ene - 3, 17 - dione, A * - 3-ketoste-
roid reductase, 449
Anemonin, 617
1,5-Anhydro-D-glucitol, glucose transport
by intestine, 264
l,5-Anhydro-D-glucitol-6-phosphate,
glucose-6-P dehydrogenase, 379
hexokinase, 377-380
hydrolysis by glucose-6-phosphatase,
379
phosphoglucomutase, 379
phosphoglucose isomerase, 379
respiration (glucose), 379
structure of, 378
Aniline, D-amino acid oxidase, 348
Anodonta eggs, mercurials, 965
Anomers, as analog inhibitors, 271-272
Anthranilate,
derivatives of, tryptophan synthesis,
321
3-hydroxyanthranilate oxidase, 594
kynurenine formamidase, 595
metabolism of, tryptophan analogs, 321
Anthriscus, malonate occurrence in, 225
Antimetabolite, definition of, 246
Antimycin A, gastric acid secretion, 915-
916
Antiseptic mercurials, see also Mercurials
structures of, 970
1136
SUBJECT INDEX
Aorta, respiration (endogenous),
malonate, 179
Apium, malonate occurrence in, 225
Aplysia depilans (sea hare) muscle,
respiration (endogenous), malonate, 174
succinate oxidation, malonate, 54
effect of fumarate on, 114
Apparent inhibitor constant, definition
of, 252
Apple, see also Malus
Apple fruit,
citrate oxidation, malonate, 79
a-ketoglutarate oxidation, malonate, 80
malate oxidation, malonate, 82
respiration (endogenous), malonate, 173
succinate oxidation, malonate, 53
Apple skin, respiration (endogenous),
resorcinol, 296
Apyrase,
o-iodosobenzoate, 707
mercurials, 860
quinacrine, 549
D-Arabinose,
/S-galactosidase, 418
glucose uptake by lymph node, 263
a-glucosidase, 423
a-mannosidase, 422
phosphoarabinose isomerase, 411
uptake by heart, glucose, 263
L-Arabinose, phosphoarabinose isomer-
ase, 411
L-Arabinose dehydrogenase, ferricyanide,
675
Arabinose-5-phosphate, hexokinase, 379
Arabinose-phosphate isomerase, see Phos-
phoarabinose isomerase
Arabitylflavin, phosphorylation of, 539
Arabono-l,4-lactone, a-glucuronidase, 426
Arbacia punctulata aggs,
cleavage,
malonate, 117, 198
mercurials, 963-965
differentiation,
o-iodosobenzoate, 726
malonate, 198
Arbacia punctualata spermatozoa,
motility, mercurials, 964
respiration (endogenous), mercurials,
882
Arcaine,
diamine oxidase, 364
structure of, 361
Arginase (L-arginine ureohydrolase),
analogs, 335-338
permanganate, 660
Argininate, carboxypeptidase B, 367
L-Arginine,
dipeptidase, 367
homoserine kinase, 357
uptake by ascites cells, ornithine, 338
Arginine decarboxylase,
mercurials, protection by substrate,
779-780
toxopyrimidine, 578
Arginine deiminase (L-arginine imino-
hydrolase),
canavanine, 353
mercurials, 833
L-Arginine iminohydrolase, see Arginine
deiminase
Arginine kinase,
o-iodosobenzoate, 707
mercurials, 833
nucleotides, 467
L-Arginine ureohydrolase, see Arginase
Argon, nitrogen fixation, 291
Arsenate,
phosphatases, 439-441
phosphate transport in S. aureus, 267
Arsenite, porphyrin biosynthesis, 162
Arsenite oxidase, o-iodosobenzoate, 707
Arsonoacetate,
intercharge distance, 7
ionization constant, 242
succinate dehydrogenase, 243
Arum maculatum (cuckoo pint) spadix,
cytochrome oxidase,
malonate, 60
mercurials, 836
respiration (endogenous), malonate, 171
182
succinate dehydrogenase,
malonate, 19, 28
mercurials, 855
succinate oxidase, malonate, 19
SUBJECT INDEX
1137
Arylesterase (aryl ester hydrolase), mer-
curials, 834
Arylsulfatase (aryl sulfate sulfohydrolase),
analogs, 443-444
ferricyanide, 675
iodine, 684
mercurials, 816
Arylsulfatase b,
o-iodosobenzoate, 707
mercurials, 816
Ascaridia galli,
a-ketoglutarate oxidation, malonate, 80
respiration (endogenous), malonate, 174
succinate accumulation, malonate, 94
succinate oxidation, malonate, 54
Ascaris muscle,
succinate decarboxylation to propio-
nate, malonate, 165
Ascites carcinoma cells,
acetate oxidation, 2-deoxyglucose, 397
amino acid uptake, analogs, 265-266
arginine uptake, ornithine, 338
citrate levels, 2-deoxyglucose, 399
fructose- 1,6-diP levels, hydrogen per-
oxide, 695
fructose utilization, galactose, 263
glucose utilization, oxamate, 435
glutamate metabolism, malonate, 152
glycine uptake, malonate, 155
glycolysis,
2-deoxyglucose, 392-393
ferricyanide, 677
hydrogen peroxide, 695
malonate, 126, 209
mercurials, 875
oxamate, 434
growth of, 2-deoxyglucose, 400
hydrogen peroxide formation in, 695
K+ uptake, mercurials, 908
malonate inhibition,
citrate accumulation, 104
a-ketoglutarate accumulation. 111
succinate accumulation, 91
NAD levels, hydrogen peroxide, 695
Na+ transport, malonate, 209
oxidative phosphorylation, malonate,
122
phosphate incorporation into ADP-
ATP, 2-deoxyglucose, 395
protein biosynthesis,
o-fluorophenylalanine, 351
malonate, 156
pyridoxine levels, deoxypyridoxol, 568
pyruvate oxidation,
2-deoxyglucose, 397
D-glucosamine, 383
oxamate, 434
pjTuvate utilization, 2-deoxyglucose,
399
respiration (endogenous), malonate, 177
respiration (glucose),
o-iodosobenzoate, 722
malonate, 125, 209
mercurials, 883
transplantability, hydrogen peroxide,
695
valine incorporation into proteins, 2-
deoxyglucose, 399
Asclepain m, iodine, 684
Ascophyllum nodosum,
iodide uptake, malonate, 209
respiration (endogenous), malonate, 169
Ascorbate oxidase,
hydrogen peroxide, 692
malonate, 21
mercurials, protection by amino acids,
778
tetrathionate, 698
Ashbya gossyjni,
acetate oxidation, malonate, 77, 87
citrate accumulation, malonate, 104
respiration (endogenous), malonate, 168
L-Asparaginase (L-asparagine amidohy-
drolase, L-asparagine deamidase),
D-asparagine, 269
hydrogen peroxide, 692
mercurials, 834
D-Asparagine,
a-amylase biosynthesis, 269
L-asparaginase, 269
L-Asparagine deamidase, see Asparaginase
Asparagine:a-ketoglutarate transaminase,
see Transaminases
Asparagine: pyruvate transaminase, see
Transaminases
Aspartase (L-aspartate ammonia-lyase),
1138
SUBJECT INDEX
analogs, 355
iodine, 685
o-iodosobenzoate, 707
mercurials, 834
Aspartate,
cysteine desulfurase, 357
glutamate decarboxylase, 327-328
succinate dehydrogenase, 36
D-Aspartate,
aspartase, 355
L-glutamate dehydrogenase, 332
L- Aspartate,
L-amino acid oxidase, 340
arginase, 337
L-glutamate dehydrogenase, 332
phosphatase (acid), 441
pyridoxamine - oxalacetate transamin-
ase, 600
Aspartate carbamyltransferase,
analogs, 467-469
o-iodosobenzoate, 707
mercurials, 834
protection by substrates, 779-781
stimulation, 816
Aspartate:a-ketoglutarate transaminase,
see Transaminases
Aspartate kinase, analogs, 356
Aspartate transcarbamylase, see Aspar-
tate carbamyltransferase
Aspergillus,
citrate accumulation, hydrogen per-
oxide, 694
succinate accumulation, malonate, 90
Aspergillus glaucus, resistance to mercu-
rials, 983
Aspergillus niger,
citrate accumulation, ferrocyanide, 677
growth of,
dehydroacetate, 632-633
ferrocyanide, 677
malonate, 195
tungstate, 614
malonate formation in, 226
malonate metabolism in, 228
respiration (sucrose), mercurials, 880
sporulation, malonate, 195
succinate dehydrogenase, malonate, 27
Aspergillus oryzae.
2-deoxyglucose utilization, 387, 400
succinate dehydrogenase, malonate, 27
Aspergillus tereus,
glucose utilization, ferrocyanide, 678
itaconate formation, ferrocyanide, 678
Atabrine, see Quinacrine
Atebrin, see Quinacrine
ATP, see Adenosinetriphosphate
ATP-ADP exchange reaction,
ADP, 444
mercurials, 874
ATP:AMP phosphotransferase, see Ade-
nylate kinase
ATP:creatine phosphotransferase, see
Creatine kinase
ATP diphosphohydrolase, see Apyrase
ATP-P, exchange reaction,
ADP, 444
o-iodosobenzoate, 705
mercurials, 872-873
Atrium, see Heart
Australorbis glabratus, respiration (endo-
genous),
iraws-aconitate, 273
malonate, 174
mercurials, 882
Auxin, transport of,
mercurials, 967
Avena sativum coleoptile,
fumarate oxidation, malonate, 81
growth of,
malonate, 196-197
mercurials, 966-968
indoleacetate uptake, mercurials, 911
malonate metabolism in, 228
respiration (endogenous), malonate, 167
169, 182, 184
succinate accumulation, malonate, 93
succinate oxidation, malonate, 53
Avocado,
cis-aconitate oxidation, malonate, 79
citrate oxidation, malonate, 78
fumarate oxidation, malonate, 81
a-ketoglutarate oxidation, malonate, 79
malate oxidation, malonate, 82
malonate formation in, 226
pyruvate oxidation, malonate, 74
succinate accumulation, malonate, 91
SUBJECT INDEX
1139
Avocado mesocarp, fatty acid formation
from acetate,
malonate, 148
2-Azaadenine,
structure of, 280
xanthine oxidase, 280-282
8-Azaguanine,
adenosine deaminase, 466, 477
deamination to 8-azaxanthine, 478
enzyme induction in liver, 478
guanine deficiency, 281
protein biosynthesis, 478
structure of, 280
xanthine oxidase, 281-282, 477
8-Azaguanosinetriphosphate,
competition ^vith GTP, 478
formation from 8-azaguanine, 478
5-Azaorotate, orotate conversion to oro-
tidylate, 480
8-Azapurines, oxidation by xanthine oxi-
dase, 281
Azaserine,
formylglycinaniide ribonucleotide ami-
dotransferase, 333
glutaminase, 356
inosinate biosynthesis, 333
phosphoribosyl-PP amidotransferase,
333
structure of, 333
Azatryptophan,
/S-galactosidase synthesis, 326
E. coli growth, 326
incorporation into enzymes, 326
L-tryptophan:sRNA ligase (AMP), 326
6-Azauracil, carcinostasis, 478
6-Azauridine,
conversion to 6-azauridylate, 480
orotidylate decarboxylase, 472
6-Azauridinediphosphate, polynucleotide
phosphorylase, 474
6-Azauridinemonophosphate,
orotidylate decarboxylase, 472, 478
polynucleotide phosphorylase, 474
8-Azaxanthine,
adenosine deaminase, 466
xanthine oxidase, 282
Azelaeate, kynurenine : a - ketoglutarate
transaminase, 608
Azide, renal transport of PAH, 205
Azotohacter agilis, malonate metabolism
in, 227-228
Azotohacter vinelandii,
growth of, tungstate, 614
nitrogen fixation, gas analogs, 292-294
succinate dehydrogenase, malonate, 26
B
Bacillus anthracis,
growth of, dehydroacetate, 632
mercurial uptake, 975
Bacillus brevis, motility of,
o-iodosobenzoate, 727
malonate, 203
Bacillus cereus,
acetate utilization, malonic diethyl
ester, 237
germination induced by L-alanine, d-
alanine, 270
growth of, dehydroacetate, 632
malonic ethyl esters, 236
Bacillus coagulans, germination of,
quinacrine, 546
Bacillus megaterium, growth of,
dehydroacetate, 632
Bacillus mesentericus, growth of,
dehydroacetate, 632
Bacillus metiens, iodine killing of, 690
Bacillus pumilus, y-aminobutyrate oxida-
tion,
malonate, 154
Bacillus pyocyaneus, o-iodosobenzoate kil-
ling of, 727
Bacillus suhtilis,
aminopterin uptake, mercurials, 911
germination of,
malonate, 195
mercurials, 972
quinacrine, 546
growth of,
dehydroacetate, 632
malonate, 195
trifluorothiamine, 531
succinate oxidation, malonate, 52
Bacteria,
growth of.
1140
SUBJECT INDEX
cyanocobalamin analogs, 589-590
D-cycloserine, 359-360
dehydroacetate, 631-633
desmethyldesthiobiotin, 588
iodine, 690
ion antagonisms, 452
kojic acid, 349
malonate, 195
mercurials, 774, 970-976
pantothenate analogs, 587
pyridine-3-sulfonate, 504
pyridoxal analogs, 575-576
riboflavin analogs, 537-538
thiamine analogs, 528-530
unsaturated lactones, 617-618
infections by, malonate, 221-224
Bacterium lactis, respiration (glucose),
mercurials, 880
Bacterium succinicum ,
acetate oxidation, malonate, 77
pyruvate oxidation, malonate, 74
succinate oxidation, malonate, 51
Balantidium coli,
motility, malonate, 203
respiration (endogenous), malonate, 173
Balanus balanoides, mercurial toxicity,
961-962
Barley, malonate occurrence in, 224
Barley roots,
bromide uptake, malonate, 116-117
ion transport, fraws-aconitate, 274
K+ uptake, malonate, 209
phosphate uptake in mitochondria, ma-
lonate, 122
Rb+ uptake, malonate, 209
respiration (endogenous), malonate,
170, 181, 210
respiration (ion-linked), <raws- aconitate,
273
respiratory quotient, malonate, 185
succinate accumulation, malonate, 91
succinate dehydrogenase, malonate, 27
succinate oxidation, malonate, 51
Beech (Fagus) roots,
respiration (endogenous), malonate, 172
succinate oxidation, malonate, 51
Bees, mannose toxicity, 414
Beets, succinate dehydrogenase,
malonate, 27
Benzamide,
ammonia formation in kidney, 348
chymotrypsin, 372
glucose dehydrogenase, 501-502
lactate dehydrogenase, 501-502
NAD nucleosidase, 488
tyrosinase, 300-301
Benzedrine, see Amphetamine
Benzenesulfonate,
arylsulfatase, 444
glucose dehydrogenase, 501-502
lactate dehydrogenase, 437-438, 501-
502
sulfite oxidase, 451
tyrosinase, 300-301
Benzimidazole, NAD nucleosidase, 492
Benzoate,
acetate accumulation in Proteus, 349
acetoacetate formation from butyrate,
613
amino acid deamination in kidney, 348
D-amino acid oxidase, 340-348
L-amino acid oxidase, 338, 348
p-aminobenzoate acetylation, 349
ammonia formation in kidney, 348
carboxypeptidase, 306
catechol oxidase, 297-302
chymotrypsin, 370, 372
citrate oxidation, 348
crotonate oxidation, 349
dehydroshikimate reductase, 606
dopa decarboxylase, 312
fatty acid oxidation, 349
glucose dehydrogenase, 501
glucose metabolism, 349
glutamate dehydrogenase, 331
D-glutamate oxidase, 349
a-ketoglutarate oxidation, 348
a-ketoisocaproate decarboxylase, 349
kynurenine:a-ketoglutarate transami-
nase, 608-609
lactate dehydrogenase, 501
NADPH dehydrogenase, 349
oxidative phosphorylation, 348
phosphorylation in mitochondria, 349
pyruvate oxidation, 349
respiration (endogenous), 348
SUBJECT INDEX
1141
respiratory quotient, 349
shikimate dehydrogenase, 349
succinate oxidation, 348
tyrosinase, 300-301, 349
yeast growth, 349
Benzoate methyl ester, catechol oxidase,
298
p-Benzoquinone, Fusarium conidial
growth, 660
Benzoyl-L-argininamide, papain, 375
Benzoyl-L-arginine, papain, 375
iV-Benzoyl-a-D-glucosamine, structure of,
378
Benzoyl-D-tryptophanamide, chymotryp-
sin, 371
Benzoyl-L-tyrosinemethylamide, chymo-
trypsin, 371
Benzylmalonate, carboxypeptidase, 367
a-Benzylmalonamide, chymotrypsin, 370
Benzyloxyamine, dopamine/J-hydroxyl-
ase, 320
Benzylsulfate, arylsulfatase, 443
Betaine, thetin.-homocysteine transme-
thylase, 356
Betaine aldehyde dehydrogenase,
o-iodosobenzoate, 707
mercurials, 781
protection by NAD, 781
protection bj' substrate, 781
Betaine:homocysteine methyltransferase,
dimethylglycine, 356
Bicarbonate, renal transport of,
mercurials, 920-921
Bioluminescence, mercurials, 888-891
Biotin,
analogs of,
biotin degradation, 589
biotin oxidase, 589
E. coli growth, 588
fermentation, 588-589
biotin oxidase, 589
intestinal transport of, analogs, 267
structure of, 588
Biotin-diaminecarboxylate, biotin oxi-
dase, 589
Biotinol, biotin oxidase, 589
Biotinol-diamine, biotin oxidase, 589
Biotin oxidase, analogs, 589
Biotinsulfone. biotin oxidase, 589
Bis (p-nitrophenyl) disulfide, determina-
tion of SH groups, 640-641
Blastocladiella emersonii, respiration (en-
dogenous),
malonate, 169
Blastocysts,
glycolysis, 2-deoxyglucose, 393
respiration (endogenous), malonate, 179
respiration (glucose), 2-deoxyglucose,
393
Blebbing of tumor cells, see Sarcoma 37
Blood,
acetoacetate accumulation, malonate,
138, 149
cholesterol level, malonate, 150
coagulation of,
o-iodosobenzoate, 725
malonate, 219
CO2 capacity of, malonate, 219
glucose level, malonate, 149, 219
glycolysis, mercurials, 877
K+ level, malonate, 206
lactate level,
malonate, 219
thiamine analogs, 520
malonate levels in vivo, 100-103
mercurial levels in vivo, 930, 958-960
Na+ level, malonate, 206
pyruvate level,
malonate, 219
pyrithiamine, 520, 527
succinate level, malonate, 100, 102
Blood pressure,
o-iodosobenzoate, 723
malonate, 213
pyrogaUol, 611
Blood vessels, see Vascular smooth muscle
Blowfly, see Calliphora
Bolaform ions, 5, 44
Bone, growth of cultures,
malonate, 199
Bone marrow, nucleic acid biosynthesis,
folate analogs, 585
Borate, phosphatases, 439-440
Borneol-a-glucuronide, a-glucuronidase,
426
1142
SUBJECT INDEX
Botrytis allii, growth of,
dehydroacetate, 632
Botrytis fabae, histidine transport,
analogs, 267
Brain,
^-acetylaspartate formation, malonate,
154
acetylcholine S5m thesis, malonate, 165-
166
ADP level, 2-deoxyglucose, 395
amino acid decarboxylase, a-methyl-m-
tyrosine in vivo, 317
amino acid metabolism, malonate, 153
amino acid synthesis from glucose, 2-
deoxyglucose, 399
amino acid transport, analogs, 266-267
y-aminobutyrate level, toxopyrimidine,
578
y-aminobutyrate oxidation, malonate,
154
6-aminonicotinamide-NAD formation
in, 505
ATP level, 2-deoxyglucose, 395
C-l/C-6 ratio,
2-deoxyglucose, 393-394
malonate, 130
catecholamine levels,
a-methyldopa, 317-318
a-methyl-m-tyrosine, 316-318
citrate level, sequential inhibition by
malonate and fluoroacetate, 112
creatine-P level, 2-deoxygluco8e, 395
cycle intermediates levels, 89
dopamine level, a-methyl-w-tyrosine,
316
glucose uptake, 2-deoxyglucose, 394
glucose utilization, malonate, 126-127,
134-135
glutamate decarboxylase in vivo,
deoxypyridoxol, 569-570
toxopyrimidine, 578
glutamate formation from glucose, ma-
lonate, 153
glutamate uptake, malonate, 153
glutamine level, 2-deoxyglucose, 399
glycolysis (aerobic), malonate, 127-128,
134-135
glycolysis (anaerobic),
dehydroacetate, 624
D-glucosone, 385
a-ketoglutarate oxidation, malonate,
80-81, 84
K+ uptake,
2-deoxyglucose, 399
malonate, 153
lactate oxidation,
nicotinamide, 500
nicotinate, 500
malonate decarboxylation in, 232
malonate level in vivo, 102
mercurial levels, 958-959
NAD level, 6-aminonicotinamide, 505
norepinephrine level,
a-methyl-m-tyrosine, 316
pyrogallol, 611
oxalacetate oxidation, malonate, 82
oxidative phosphorylation, mercurials,
873
phospholipid biosynthesis, malonate,
151
pjrridoxal-P level, deoxypyridoxol, 567-
569
pyrithiamine levels in vivo, 528
pyruvate oxidation, malonate, 75-76,
128
pyruvate utilization,
malonate, 135
oxygen, 658-659
respiration (endogenous),
dehydroacetate, 623-624
malonate, 175-177, 179, 181, 183-
184
respiration (glucose),
hydrogen peroxide, 695
kojic acid, 350
malonate, 124, 127, 133-135
nicotinamide, 500
nicotinate, 500
oxygen, 658-659
quinacrine, 560
tartronate, 238
respiration (glutamate), malonate, 152
serotonin level, a-methyl-m-tyrosine,
316
succinate accumulation, malonate, 95,
97
SUBJECT INDEX
1143
succinate dehydrogenase,
maleate, 36
malonate, 31-32
oxalacetate, 36
succinate levels in vivo, malonate, 102
succinate oxidation,
hydrogen peroxide, 695
malonate, 55
thiamine-PP level, oxythiamine, 526-
527
Branched chain ketonuria (maple sugar
urine disease), role of glutamate de-
carboxylase inhibition, 329
Brevibaderium flavum, respiration (glu-
cose),
mercurials, 880
Bromate, nitrite oxidation, 450
Bromelain,
iodine, 685
mercurials, 792, 810, 813
pH effects, 792
rate of inhibition, 810, 813
Bromide,
^-ketoadipate chlorinase, 453
tyrosinase, 300
uptake by barley roots, malonate, 116-
117
m-Bromobenzoate, glutamate dehydro-
genase, 330
Bromobenzoates, D-amino acid oxidase,
341
5-Bromodeoxycytidine, aspartate carba-
myltransferase, 469
5-Bromofuroate, glutamate dehydrogen-
ase, 330-331
^-Bromopropionate, structure of, 41
Bromopyruvate,
glycerate dehydrogenase, 430
lactate dehydrogenase, 437
Bronchopneumonia virus, inactivation by
mercurials, 977
Brucella abortus, respiration (endogenous),
malonate, 168
Bunias orientalis, malonate occurrence in,
225
Bush bean, see Phaseolus vulgaris
2,3-Butanedione, pyruvate decarboxy-
lase, 431
1 ,4-Butanediphosphonate,
ionization constants, 242
succinate dehydrogenase, 243
a-ButylglucopjTanoside, a-glucosidase,
423
2'-Butylthiamine, thiamine kinase, 523
Butynamine demethylase, 2,4-dichloro-6-
phenylphenoxyethylamine, 592
Butyrate,
D-amino acid oxidase, 343
conversion to acetoacetate, analogs,
613
glutamate decarboxylase, 328
homoserine kinase, 357
kynurenine:a-ketoglutarate transami-
nase, 608
lactate dehydrogenase, 436
leucine decarboxylase, 352
tyrosinase, 300
Butyryl-CoA dehydrogenase, mercurials,
781
protection by FAD, 781
protection by substrate, 781
Cadaverine,
kynureninase, 595
oxidation of, analogs, 363
structure of, 361
Cadaverine oxidase, see Diamine oxidase
Cadmium, complexes with di- and tri-
carboxylates, 12
Caffeate (3,4-dihydroxycinnamate),
acetylindoxyl oxidase, 591
dopa decarboxylase, 314
glutamate decarboxylase, 314
histidine decarboxylase, 314
indoleacetate oxidase, 595
peroxidase, 599
succinate dehydrogenase, 314
tyrosinase, 314
tyrosine decarboxylase, 314
Caffeine,
D-araino acid oxidase, 545
NAD nucleosidase, 492
Calcium,
ATPase, 453
intestinal transport of.
1144
SUBJECT INDEX
malonate, 208
mercurials, 909, 913-914
oxidative phosphorylation, 453
phosphorylase b kinase, 453
uptake by mitochondria, mercurials,
909
urinary excretion, mercurials, 921
Caldariomyces fumago, respiration (glu-
cose),
malonate, 125
Calliphora erythrocephala,
a-ketoglutarate oxidation, malonate,
80, 84
oxidative phosphorylation, malonate,
121
Camellia pollen, malonate metabolism in,
228
Canavanine, arginine deiminase, 353
Candida albicans, growth of,
dehydroacetate, 632
Candida utilis, resistance to mercurials,
983-984
Cape barley, see Hordeum vulgare
Caprate (decanoate), kynurenine: a-keto-
glutarate transaminase, 595, 608
Caproate (hexanoate),
glutamate decarboxylase, 328
kynurenine:a-ketoglutarate transami-
nase, 608
Caprylate (octanoate),
kynurenine: a-ketoglutarate transami-
nase, 608
lactate dehydrogenase, 436
Carbamate kinase, see Carbamyl-phos-
phate synthetase
Carbamyl-ADP phosphotransferase, mer-
curials, 834
Carbamyl-phosphate : L-aspartate carba-
myltransferase, see Ornithine carbamyl-
transferase
Carbamyl-phosphate synthetase, mercu-
rial, 781, 804
protection by acetylglutamate, 781
relation to SH groups, 804
Carbobenzoxy-L-glutamate, papain, 375
Carbohydrate transport, see also specific
sugars
analogs, 262-264
Carbon dioxide, fixation of,
mercurials, 892
Carbon dioxide activating enzyme, mer-
curials, 834
Carbonic anhydrase,
ferricyanide, 675
hydrogen peroxide, 692
iodine, 685
mercurials, 834-835
p-mercuribenzoate, competitive nature
of inhibition, 771-772
Carbon monoxide, nitrogen fixation, 291-
292
Carboxylate group, interaction energy,
276
Carboxylesterase, see Aliesterase
Carboxymethylcellulose, chymotrypsin,
457
Carboxypeptidase,
active center of, 365-366
analogs, 365-367
malonate, 60
mercurials, 770, 781,818
protection by Zn++, 781
stimulation, 818
Carboxypeptidase A, mercurials,
failure of Zn++ to reverse, 827
Carcinoma, see also Ascites carcinoma
cells. Walker carcinoma
growth of, 2-deoxyglucose, 400
succinate dehydrogenase, malonate, 32
Carcinostasis, see Tumors, growth of
Carcinus maenas, oxidative phosphoryla-
tion,
malonate, 121
Caries, from dehydroacetate, 630-631
Carnation seeds, growth of,
mercurials, 965
Carotid artery, contractility,
malonate, 212
Carotid body, excitability,
malonate, 211-212
Carp liver,
fatty acid oxidation, malonate, 137
a-ketoglutarate oxidation, malonate, 80
oxidative phosphorylation, malonate,
121
pyruvate oxidation, malonate, 75
SUBJECT INDEX
1145
succinate accumulation, malonate, 91
Carrots,
glucose uptake, 2 -deoxy glucose, 394
isocitrate oxidation, malonate, 79
malate oxidation, malonate, 82
pyruvate oxidation, malonate, 74
respiratory quotient, malonate, 185
succinate accumulation, malonate, 97
succinate oxidation, malonate, 22
Catalase,
cystine, 662-663
dehydroacetate, 623
induction of, 8-azaguanine, 478
mercurials, 835
rate of inhibition, 810
relation to SH groups, 803
monoethylperoxide, 592, 694
Catechol,
acetylindoxyl oxidase, 591
D -amino acid oxidase, 344
dehydroshikimate reductase, 593, 605
histidine decarboxylase, 352
Catecholamines, see also specific amines
metabolism of, pyrogallol, 611-612
release of,
o-iodosobenzoate, 725
mercurials, 947
a-methyldopa, 315-320
a-methyl-TO-tyrosine, 315-316
Catechol-0-methyl transferase,
effect of inhibition on catecholamine
metabolism, 611-612
pyrogallol, 592, 611-612
Catechol oxidase,
analogs, 296-298
benzoate,
derivatives of, 297-302
pH effects, 298-299
cinnamate, 298
dihydroxymaleate, 297
quinacrine, 547, 549
Cathepsin,
iodine, 685, 688
o-iodosobenzoate, 707
Cathepsin C, analogs, 375
Cauliflower buds,
citrate oxidation, malonate, 78, 87
a-ketoglutarate oxidation, malonate, 79
Cecropia silkworn, spermatid meiosis in,
malonate, 199
Cell division, see also specific organisms or
tissues
2-deoxyglucose, 400
heparin, 462
o-iodosobenzoate, 726-727
malonate, 197-200
mercurials, 963-970
Cellobiase, mercurials, 860
Cellobiose,
)S-glucosidase, 417
a-mannosidase, 422
Cellulose polysulfatase, ferricyanide, 675
Cellulose, biosynthesis of,
malonate, 132
Central nervous system, see Brain
Cephalosporin C, penicillinase, 599
C-esterase, mercurials, 816
Chaetonium globosum, growth of,
dehydroacetate, 633
Chaetopterus eggs, cleavage,
malonate, 198
mercurials, 965
Chelidonate, D-amino acid oxidase, 342
Chick embryo, see also Embryogenesis
development and differentiation,
3-acetylpyridine, 494
malonate, 199
glucose uptake by heart, 2-deoxyglu-
cose, 394
growth of heart fibroblasts, 2-deoxyglu-
cose, 400
protein biosynthesis in cultures, ma-
lonate, 156
respiration (endogenous) of condyles,
malonate, 175
respiration (endogenous) of yolk sac,
malonate, 175
succinate oxidation in cartilage, ma-
lonate, 54
Chicory,
respiration (endogenous) malonate, 173,
182-183
respiratory quotient, malonate, 185
Chitin disulfate, ^-fructofuranosidase, 465
Chitin sulfates, hyaluronidase, 459
Chlorate,
1146
SUBJECT INDEX
nitrite oxidation, 450
Nitrobacter growth, 450
Chlorella,
hydrogen production, mercurials, 891
photosynthesis, malonate, 163
primary photogenic agent, mercurials,
892
succinate oxidation, malonate, 51
Chlorella pyrenoidosa, malonate metabo-
lism in, 228
Chlorella vulgaris,
respiration (endogenous)
malonate, 169
malonic diethyl ester, 237
mercurials, 881
respiration (glucose), mercurials, 881
Chloride,
intestinal transport of, mercurials, 910
tyrosinase, 300-301
Chlormerodrin (Neohydrin), see also Mer-
curials, kidney
structure of, 917
Chloroacetate,
pantoate:/S-alanine ligase, 597
tyrosinase, 300
Chloroacetyl-D-tyrosinamide, chymotryp-
sin, 371
Chloroacetyl-L-tyrosinate, chymotrypsin,
371
m-Chlorobenzoate, glutamate dehydro-
genase, 330
Chlorobenzoates,
D-amino acid oxidase, 341
catechol oxidase, 298-299
6-Chloro-2,8-dihydroxypurine,
urate metabolism in vivo, 285
uricase, 284-285
5-Chlorofuroate, glutamate dehydroge-
nase, 330
Chlorogenate,
indoleacetate oxidase, 595
peroxidase, 599
(5-Chlorolevulinate, aminolevulinate de-
hydrase, 591
2-Chloromethyl-5-hydroxy-l,4-pyrone, d-
amino acid oxidase, 342
Chlorophyll, mercurials, 891
ChlorophyUase, ferricyanide, 675
a-Chloroproprionate , pantoate :/9- alanine
ligase, 598
/3-Chloropropionate, pantoate : ^ - alanine
ligase, 598
6-Chloropurine,
oxidation to 6-chlorourate, 281, 285
purine metabolism in vivo, 281
xanthine oxidase, 281-282
Chloropyru vate ,
lactate dehydrogenase, 437
pyruvate decarboxylase, 431
4-Chlororesorcinol, tyrosinase, 302, 304
6-Chlororiboflavin analog,
flavokinase, 539
Lactobacillus growth, 537
riboflavin deficiency, 538
7-Chlororiboflavin analog,
Lactobacillus growth, 537
riboflavin deficiency, 538
6-Chlorourate,
formation from 6-chloropurine, 281,
285
uricase, 284-285
xanthine oxidase, 282
Chloroxythiamine, structure of, 517
Cholestane, cholesterol esterase, 592
Cholestan-3-one, cholesterol esterase, 592
Cholesterol, see also Sterols
biosynthesis of,
malonate, 149-151
a-phenylbutyrate, 614
Cholesterol esterase (sterol ester hydro-
lase),
analogs, 592
mercurials, 835
Choline, thetin:homocysteine transmethy-
lase, 356
Choline acetylase,
iodine, 685
o-iodosobenzoate, 707
mercurials, 835
spontaneous reversal of inhibition,
813-814
pantothenate analogs, 587
Choline dehydrogenase, tetrathionate,
698-699
Choline oxidase,
analogs, 290-291
SUBJECT INDEX
1147
0-iodosobenzoate, 707, 717
protection by choline, 717
kojic acid, 350
malonate, 60
oxygen inactivation of, 659
Cholinesterase,
dehydroacetate, 622
ferricyanide, 675
GSSG, 662
iodine, 683, 685
o-iodosobenzoate, 707
mercurials, 772-773, 776-778, 788, 810,
835-836, 937-938
kinetic analysis, 776-778
possible denaturation, 788
rate of inhibition, 810
role in muscle action, 937-938
type of inhibition, 772-773
quinacrine, 549
succinyl peroxide, 694
thiamine analogs, 531-532
Choline sulfa tase,
phosphate, 444
sulfate, 444
sulfite, 444
Chondroitin sulfate,
fumarase, 465
phosphatase (acid), 465
ribonuclease, 462
synthesis of, malonate, 166
ulcer reduction, 458
Chondroitin sulfate B, hyaluronidase, 460
Chorioallantoic membrane, respiration
(endogenous),
malonate, 175
Chorioallantoic membrane (virus-infect-
ed),
glucose uptake, malonate, 126-127
respiration (endogenous), malonate,
126-127
Choroid plexus, iodide uptake,
malonate, 209
Chromatium, NAD iihotoreduction,
mercurials, 891
Chromatophores, migration in culture,
malonate, 203
Chymotrypsin,
acridines, 373
active center of, 374
analogs, 368-374
hydrogen peroxide, 694
iodine, 686
macroions, 457
oxidation of, 657
periodate, 657
proteins, 457
quinolines, 373
D-tryptophanides, 271
D-tyrosinamides, 271
Ciliary body, iodide transport,
malonate, 209
nitrate, 267
Ciliary motility,
malonate, 203
mercurials, 981-982
Cinnamide, D-amino acid oxidase, 343
Cinnamate,
acetoacetate formation from butyrate,
613
D-amino acid oxidase, 342, 346
carboxypeptidase, 367
catechol oxidase, 298
derivatives of, dopa decarboxylase,
311-314
structure of, 296
Citraconate (methylmaleate),
fumarase, 279
glutamate decarboxylase, 328
ionization constants, 8
structure of, 279
Citrate,
accumulation of,
trans-sicomtsite, 273-274
ferrocyanide, 677-678
fluoromalonate, 239
hydrogen peroxide, 694
malonate, 104-110, 223
malonic diethyl ester, 236
mercurials, 927
formation of, malonate, 105, 108
fumarase, 275
glutamate dehydrogenase, 331
level in ascites cells, 2-deoxyglucose,
399
malate dehydrogenase, 596
oxidation of.
1148
SUBJECT INDEX
benzoate, 348
malonate, 78-79, 86-87
mercurials, 878
phosphofructokinase, 385-386
urinary excretion of, malonate, 104,
109-110
Citrate (isocitrate) hydro-lyase, see Aco-
nitase
Citrate synthetase (condensing enzyme,
oxalacetate transacetase),
malonate, 63
mercurials, 836
palmityl-CoA, 614
CitruUine,
arginase, 337
carbamyl - P : ornithine transcarbamyl-
ase, 592
Claviceps purpurea (ergot), succinate de-
hydrogenase,
fumarate K,, 38
malonate K,, 33
Cleavage, see Cell division
Clostridium, nitrogen fixation,
hydrogen, 292
Clostridium histolyticum , growth of,
mercurials, 972
Clostridium kluyveri, malonate metabo-
lism in, 232
Clostridium pasteurianum, nitrogen fixa-
tion,
nitric oxide, 292
Clostridium saccharobutyricum, growth of,
malonate, 195
Clostridium welchii, growth of,
mercurials, 972
Clover, malonate occurrence in, 224, 226
Cloxacillin, penicillinase, 598
Coagulase, mercurials, 860
Cobalt, complexes with di- and tricar-
boxylates, 12
Cocciodioides immitis, spherulation of,
mercurials, 971, 973
Cochliobolus miyabeanus, germination of,
mercurials, 973
Cockroach, see also Periplaneta
Cockroach muscle, succinate oxidation,
malonate, 22
Coenzyme A,
formation of, pathways, 586-587
kidney level, mercurials, 927
mercurial complexes, 750
yeast level, mercurials, 750, 885
Coenzyme A analogs, structures of, 587
Coenzyme analogs, 482-590
recombination technic for demonstrat-
ing inhibition, 250
sites of action, 482-483
Coffee trees, mercurial fungicides causing
Zn-deficiency disease, 966
Coliphage,
mercurial inactivation of, 977, 980
proliferation of,
malonate, 194
mercurials, 977, 980-981
Colpidium, succinate accumulation,
malonate, 91
Colpidium campylum,
pyruvate oxidation, malonate, 74
respiration (endogenous), malonate, 173
Colpidium colpoda,
motility of, mercurials, 982
toxicity of mercurials, 982
Common cold virus, infectivity of,
mercurials, 977
Condensing enzyme, see Citrate synthe-
tase
Configurational changes,
enzymes,
alcohol dehydrogenase, 788-789
penicillinase, 249, 615, 688
phosphoribosyl-ATPpyrophosphory-
lase, 351
succinate dehydrogenase, 46
produced by,
mercurials, 787-790
oxidants, 656
SH reagents, 648-650
proteins, 657, 703-704, 761-762
Contracture, see also specific inhibitors
and muscles
heart,
mercurials, 941-944
porphyrindin, 669
intestinal muscle, o-iodosobenzoate, 724
muscle,
o-iodosobenzoate, 723
SUBJECT INDEX
1149
mercurials, 896, 938
tetrathionate, 699
uterus, o-iodosobenzoate, 724
Copper,
catalysis of SH group oxidation, 658
complexes with di- and tricarboxylates,
12
/^-glucuronidase, 795
toxicity to Nitocra, 962
uptake by liver, mercurials, 910, 913
Coproporphyrinogen oxidase, mercurials,
860
Corn (maize) roots,
respiration (endogenous), malonate, 172
pH effects, 191
respiratory quotient, malonate, 185
Cornea, water transport,
mercurials, 911
Coronary floAv, malonate, 213
Corynebacterium, succinate oxidation,
malonate, 52
Corynebacterium creatinovorans, succinate
accumulation,
malonate, 92
Corynebacterium diphtheriae,
growth of, dehydroacetate, 632
succinate dehydrogenase,
fumarate K,, 38
malonate Kj, 33
Coumalate (a-pyrrone-5-carboxylate), D-
amino acid oxidase, 342
<rans-23-Coumarate, phenylalanine deami-
nase, 355
Coxsackie virus, infectivity of,
mercurials, 977
Crabtree effect,
2-deoxyglucose, 396-398
D-glucosone, 385
oxamate, 435
Crassostrea virginica (oyster) mantle,
respiration (endogenous), malonate, 174
succinate oxidation, malonate, 54
Creatinase, mercurials, 836
Creatine kinase,
adenosine, 446
ADP, 447
6-aminonicotinamide-NAD, 505
iodine, 682, 685
o-iodosobenzoate, 704, 707
malonate, 60
mercurials, 836
nitrate, 446
phosphate, 446
sulfate, 446
tripolyphosphate, 446
Creatine - phosphate ,
muscle homogenate level, o-iodosoben-
zoate, 721
tissue levels, 2-deoxyglucose, 395
Creatinine, renal transport of,
dehydroacetate, 625
Crepis capillaris roots, growth of,
mercurials, 966
2?-Cresol, tyrosine: a-ketoglutarate trans-
aminase, 306
o-Cresotamide, sulfanilamide acetylase,
601
Crithidia fasciculata,
succinate dehydrogenase, malonate, 28
succinate oxidation, malonate, 22-23,
54
Crocker 180 sarcoma,
glycolysis, ferricyanide, 677
respiration (endogenous), malonate, 177
Crotonase, see Enoyl-CoA hydratase
Crotonate (/3-methylacrylate),
D-amino acid oxidase, 343, 346
fumarase, 279
oxidation of, benzoate, 349
structure of, 279, 345
Crotonyl-CoA, acyl-CoA dehydrogenase,
591
Crown galls, growth of,
malonate, 197
Crown gall organism, see Agrobacterium
tumefaciens
Cryptococcus terricolus,
malonate oxidation in, 231
respiration (endogenous), malonate, 231
Cuckoopint, see Arum maculatum
Cupric ions, see Copper
Cyanate, nitrite oxidation, 450-451
Cyanide,
poisoning by, tetrathionate antago-
nism, 696
renal transport of PAH, 205
1150
SUBJECT INDEX
Cyanocobalamin,
analogs of,
bacterial growth, 589-590
cyanocobalamin biosynthesis, 589
diol dehydrase, 590
methionine biosynthesis, 590
biosynthesis of, analogs, 589
Cyclic - 2', 3'- adenosinemonophosphate,
isocitrate dehydrogenase, 509
Cyclic - 3 ', 5 '- adenosinemonophosphate,
phosphofructokinase, 474
Cyclic-2',3'-guanosinemonopho8phate,
deoxycytidylate deaminase, 469
Cyclobutane- 1 , 1 -dicarboxylate,
intercharge distance, 7
succinate dehydrogenase, 37, 40
Cyclohexaneacetate, chymotrypsin, 370
y-CycIohexanebutyrate,
chymotrypsin, 370
kynurenine:a-ketoglutarate transami-
nase, 608-609
Cyclohexanecarboxylate,
kynurenine:a-ketoglutarate transami-
nase, 608-609
tyrosinase, 300-302
Cyclohexane- 1 ,2-dicarboxylate,
intercharge distance, 7
kynurenine:a-ketoglutarate transami-
nase, 608
succinate dehydrogenase, 38, 40
^-Cyclohexanepropionate, chymotrypsin ,
369-370
Cyclohexyl-DL-alanine, phenylalanine hy-
droxylase, 354
Cyclohydrolase, folate analogs, 585
Cyclopentane-l,2-dicarboxylate,
acetate utilization, 241
intercharge distance, 7
pyruvate utilization, 241
succinate dehydrogenase, 37, 40, 241
D-Cycloserine (orientomycin, Oxamycin),
D-alanyl-D-alanine synthetase, 360
L-asparagine:a-ketoglutarate transami-
nase, 360
bacterial growth, 359-360
antagonism by D-alanine, 359
antagonism by alanylalanine, 360
cell wall formation, 359
central nervous system, 359
GABA : a - ketoglutarate transaminase,
359
glutamate decarboxylase, 359
mycobacterial growth, 359
oxime formation with pyridoxal-P, 359
staphylococcal resistance to, 359
structure of, 359
transaminases, 359
L-Cycloserine,
L-alanine:a-ketoglutarate transaminase,
360
L-asparagine:a-ketoglutarate transami-
nase, 360
bacterial growth, antagonism by L-ala-
nine, 359
Cystamine, glucose utilization in erythro-
cytes, 663
Cystamine monosulfoxide, 3-phosphogly-
ceraldehyde dehydrogenase, 663
D-Cysteine, L-alanine dehydrogenase, 354
L-Cysteine,
arginase, 337
homoserine kinase, 357
serine deaminase, 357
tyrosine decarboxylase, 307
Cysteine desulfurase,
analogs, 357
deoxypyridoxol in vivo, 570
malonate, 60
Cystine,
enzyme inhibitions, 661-664
oxidation of protein SH groups, 661,
663
Cytidine,
aspartate carbamyltransferase, 467-468
5 '-nucleosidase, 472
Cytidinediphosphate (CDP),
aspartate carbamyltransferase, 468
NADH oxidase, 511
Cytidinemonophosphate (CMP),
adenylosuccinate synthetase, 467
aspartate carbamyltransferase, 467-468
deoxycytidylate deaminase, 469
pyrophosphatase, 475
ribonuclease, 475
Cytidinetriphosphate (CTP), aspartate
carbamyltransferase, 468
SUBJECT INDEX
1151
Cytochrome bj, o-iodosobenzoate, 708
Cytochrome bj reductase,
iodine, 685
mercurials, coenzyme displacement,
787
Cytochrome c, xanthyl-cytochrome c, 592
Cytochrome c oxidase,
o-iodosobenzoate, 708
malonate, 60
mercurials, 837, 870-872
quinacrine, 550
thiols, 661, 663
Cytochrome c reductase,
mercurials, relation to SH groups, 804,
809
quinacrine, 547, 549
Cytochrome c-554 reductase, quinacrine,
549
Cytosine, D-amino acid oxidase, 545
D
Dahlia leaves, CO2 photochemical fixation
mercurials, 892
dAMP, see Deoxyadenosinemonophos-
phate
Daptazole, see 2,4-Diamino-5-phenylthia-
zole
Daraprim, see Pyrimethamine
dCDP, see Deoxycytidinediphosphate
dCMP, see Deoxycytidinemonophosphate
dCTP, see Deoxycytidinetriphosphate
DDD,see2,2'-Dihydroxy-6,6'-dinaphthyl-
disulfide
DDT dehydrochlorinase, ferricyanide, 675
Deamino-AMP, NADPHrcytochrome c
oxidoreductase, 511
Deamino-ATP, NAD pyrophosphorylase,
510
2-Deaminofolate, serine biosynthesis, 585
Deamino-NAD,
NADH oxidase, 511
NADPH:cytochrome c oxidoreductase,
511
Deamino-NADP, NADPH:glutathione
oxidoreductase, 512
Decane - 1,10 - dicarboxylate, kynurenine:
a-ketoglutarate transaminase, 608
Decanoate, see Caprate
Dehydroacetate,
antidotes to, 628-629
ATP level in nuclei, 624
bacterial growth, 617-618, 631-633
binding to plasma proteins, 630
blood levels of, 627-628
cariogenic action, 630-631
central nervous system, 627
chemical properties, 618-620
determination in tissues, 620
diuresis, 625
enzyme inhibitions, 620-623
fungal growth, 632-633
glucose conversion to CO2, 624
glycolysis (anaerobic), 624
growth of microorganisms, pH effects,
633
heart, 625
hydrogenation of, 619
intestine, 624-625
ionization of, 619
keto-enol tautomerism, 618
lethal doses, 627-628
metabolism of, 629-630
oxidative phosphorylation, 623
penicillin levels in blood, 626
permeability to, 625
purification of, 620
reaction with thiols, 621
renal transport of PAH, 205
renal transports, 625-626
respiration (endogenous), 623-624
salivary secretion of, 630
solubility of, 619
structure of, 618-619
succinate oxidase, 620-622
synthesis of, 619-620
tissue distribution of, 629-631
toxicity of, 627-629
urinary excretion of, 629-631
urinary succinate excretion, 628
yeast growth, 632
Dehydroshikimate reductase,
active center of, 607
analogs, 593, 604-606
Dendraster eggs,
cleavage, malonate, 198
1152
SUBJECT INDEX
development, ferricyanide, 678
Deoxyadenosinemonophosphate (dAMP),
adenylosuccinate synthetase, 467
aspartate carbamyltransferase, 469
deoxycytidylate deaminase, 469
phosphodiesterase, 473
DeoxyAMP, see Deoxyadenosinemono-
phosphate
3-Deoxy-D-ara6o - heptonate -7 -phosphate,
2-keto-3-deoxy-D- arabo - heptonate-7- P
synthetase, 413
DeoxyCDP, see Deoxycytidinediphos-
phate
DeoxyCMP, see Deoxycytidinemonophos-
phate
DeoxyCTP, see Deoxycytidinetriphos-
phate
Deoxycytidine, aspartate carbamyltrans-
ferase, 468
Deoxycytidinedisphosphate (dCDP),
hydrolysis of, deoxyCMP and deoxy-
CTP, 446
polynucleotide phosphorylase, 474
Deoxycytidinemonophosphate (dCMP),
aspartate carbamyltransferase, 468
deoxyCDP hydrolysis, 446
polynucleotide phosphorylase, 474
Deoxycytidinetriphosphate (dCTP),
aspartate carbamyltransferase, 468
deoxyCDP hydrolysis, 446
polynucleotide phosphorylase, 474
Deoxycytidylate deaminase,
analogs, 469
mercurials, 837
protection by deoxyCTP, 781
Deoxycytidylate kinase, ADP, 469
6-Deoxy-6-fluoroglucose,
acetate metabolism, 404
fructose fermentation in yeast, 404
glucose fermentation in yeast, 404
glucose oxidation in kidney, 393
glucose utilization, 403-405
hexokinase, 404
intestinal transport of, 404
lactate metabolism, 404
lethal doses, 404-405
membrane transport of hexoses, 404
metabolism of, 404
oxidation by glucose oxidase, 404
2-Deoxygalactose, respiration (galactose),
391-392
4-Deoxygalactose, respiration (galactose),
391-392
2-Deoxygluconate, metabolism of, 389
1-Deoxyglucose,
intestinal transport of, 387
a-methylglucoside uptake, 394
2-Deoxyglucose,
absorption of, 386-387
acetate metabolism in rabbit, 399
acetate oxidation, 397
ADP levels in tissues, 394-398
amino acid sjTithesis from glucose, 399
anaphylactoid reaction, 401
ATP levels in tissues, 394-398
C-l/C-6 ratio in brain, 383-394
carcinostasis, 386, 400-401
cell division, 400
central nervous system, 401
citrate level in ascites cells, 399
CO2 formation from glucose, 393
Crabtree effect, 396-397
creatine-P levels, 395
distribution in tissues, 386-387
DNA level in carcinoma, 399
epinephrine release, 401
Escherichia coli growth, 400
ethanol oxidation, 395-396
fatty acid level in plasma, 399
fatty acid oxidation, 397
fibroblast culture growth, 400
fructose metabolism, 391, 398
fructose uptake, 394
galactose oxidation, 398
/5-galactosidase synthesis, 400
glucokinase, 389-390
glucose membrane transport, 390
glucose metabolism, 391-398
glucose-6-phosphatase, 390
glucose uptake, 393-394
glucose uptake in vivo, 387-388, 401
glutamine level in brain, 399
glycolysis, 391-394
heart, 402-403
hexokinase, 389-390
hyperglycemia, 401
SUBJECT INDEX
1153
IMP level in ascites cells, 395
intestinal transport of, 387
K+ fluxes in atria, 403
K+ uptake by brain, 399
lethal doses, 401
lipogenesis in liver, 399
mannose uptake, 394
metabolism of, 386-389
a-methylglucoside uptake, 394
palmitate oxidation, 397
pentose-P pathway stimulation, 393-
394
phosphorj-lation of, 387-389
epinephrine effect, 387
insulin effect, 387
K^'s for hexokinases, 388
protein biosynthesis, 399
pyruvate decarboxylation, 396
pyruvate oxidation, 392, 397
pjTuvate utilization in ascites cells, 399
resistance of HeLa cells to, 388
respiration (endogenous), 391-392, 396-
397
comparison with glucose, 396-397
respiration (glucose), 391-394, 397
combined with iodoacetate, 397
summary of mechanisms of action on
carbohydrate utilization, 398
toxicity, 401
transport into cerebrospinal fluid, 401
urinary excretion of, 388
utihzation by fungi, 387, 400
virus proliferation, 400
3-Deoxyglucose,
intestinal transport of, 387
a-methylglucoside uptake, 394
6-Deoxygluco8e,
glucose oxidation, 403
intestinal transport of, 403
intestinal transport of hexoses, 264, 403
a-methylglucoside uptake, 394
2-Deoxyglucose-6-phosphate,
formation of, 387-389
glucose transport, 390
glucose-6-P dehydrogenase, 390-391
glycogen synthetase, 391
oxidation of, 388
phosphoglucose isomerase, 390
UDPG:a-l,4-glucan-a-4-glucosyltrans-
ferase, 391
3-Deoxyglucose-6-phosphate,
hexokinase, 380
structure of 378
Deoxyguanosinemonophosphate (dGMP),
adenylosuccinate synthetase, 467
deoxycytidylate deaminase, 469
DeoxypjTidoxine, see DeoxypjTidoxol
Deoxypyridoxol (deoxypyridoxine, des-
oxypyridoxine ) ,
acrodynia, 566
blood cholesterol, 574
blood phospholipids, 574
carcinostasis, 576
cysteine desulfurase, 570
dermatitis, 577
fatty acid biosynthesis, 574
glutamate decarboxylase, 569-570
growth of microorganisms, 575-576
intestinal amino acid transport, 574
intestinal sugar transport, 574
leucocyte count, 577
malignant carcinoid syndrome, 574
phosphorylation of, 564
pyridoxal kinase, 565
pyridoxal-P level in tissues, 566
p>Tidoxamine-P oxidase, 566
p\Tidoxine levels in tissues, 566-568
pyridoxol-P oxidase, 566
serine biosynthesis, 570-571
serotonin metabolism, 574
structure of, 563
toxicity, 562,566-567, 577-578
Toxoplasma infections, 576
transaminases, 569-570
tryptophan metabolism, 572
urinary excretion of xanthurenate, 572
vitamin Bg deficiency, 562, 577-578
Deoxypyridoxol-phosphate,
pyridoxamine-P oxidase, 566
pyridoxol oxidation, 564
pyridoxol-P oxidase, 566
5'-Deox>Tiboflavin, riboflavin biosynthe-
sis, 539
Deoxyribonuclease,
ferricyanide, 675
mercurials, 860
1154
SUBJECT INDEX
RNA, 462
Deoxyribonucleates, see also Nucleic acids
carcinoma levels, 2-deoxyglucose, 399
ribonuclease, 462
Deoxyribose-phosphate aldolase, mercu-
rials, 837
2'-Deoxyribosyl-4-aminopyrimidone-2,5'-
phosphate deaminase, ferri cyanide, 675
Deoxythymidine, aspartate carbamyl-
transferase, 469
Deoxythymidine kinase, deoxyTTP, 470
Deoxythymidinemonophosphate (dTMP),
deoxycytidylate deaminase, 469
Deoxythymidinetriphosphate (dTTP), de-
oxythymidine kinase, 470
Deoxyuridinemonophosphate (dUMP), de-
oxycytidylate deaminase, 469
Desert locust, see also Schristocera
butyrate oxidation in muscle, malonate,
137
Desmethyldesthiobiotin ,
Escherichia coli growth, 588
structure of, 588
Desoxypyridoxine, see DeoxypjTidoxol
Desthiobiotin,
biotin oxidase, 589
structure of, 588
Desulfovibrio desulfuricans,
growth of, mercurials, 972
tetrathionate reduction in, 699
a-l,6-Dextranglucosidase, analogs, 417
Dextrin- 1,6-glucosidase, see Amylo-1,6-
glucosidase
DFPase, o-iodosobenzoate, 711
dGMP, see Deoxyguanosinemonophos-
phate
Diacetylthiamine, thiamine kinase, 523
Dialkylfluorophosphatase, see also DFP-
ase
malonate, role of Mn++ in inhibition, 68
Diamidines,
diamine oxidase, 363-365
structures of, 361
Diamine oxidase (histaminase),
analogs, 360-365
hydrazine, 362
malonate, 60
semicarbazide, 362
2,4-Diamino-6,7-dihydroxypteridine, xan-
thine oxidase, 289
1 ,2-Diamino-4,5-dimethylbenzene, cyano-
cobalamin biosynthesis, 590
2,4-Diamino-7,8-dimethyl-10-ribityl-5,10-
dihydrophenazine, structure of, 537
2,4-Diamino-6-formylpteridine, dihydro-
folate reductase, 583-584
a,£-Diamino-^-hydroxypimelate, diami-
nopimelate decarboxylase, 593
2,4-Diamino-6-hydroxypteridine,dihydro-
folate reductase, 583-584
2,6-Diamino-8-hydroxypurine, xanthine
oxidase, 282
2,4-Diamino-6-methylpteridine, dihydro-
folate reductase, 583-584
2,4-Diamino-5-phenylthiazole (amiphena-
zole, Daptazole), thiamine deficiency,
531
Diaminopimelate decarbocylase,
analogs, 593
o-iodosobenzoate, 708
2,3-Diaminopropionate, aspartate:a-keto-
glutarate transaminase, 355
2,6-Diaminopurine,
dihydrofolate reductase, 583-584
purine metabolism, 480
xanthine oxidase, 282
Diaphragm,
acetate uptake, mercurials, 912
acetate utilization, propionate, 613
amino acid uptake, malonate, 155
carbohydrate uptake, competition be-
tween sugars, 264
C-l/C-6 ratio, malonate, 130
citrate accumulation, malonate, 104
contractility, mercurials, 896
2-deoxyglucose uptake,
glucose, 387
insulin, 387
fructose oxidation, 2-deoxyglucose, 398
galactose oxidation, 2-deoxyglucose,
398
glucose uptake, mercurials, 876, 893-
894
glucose utilization,
6-deoxy-6-fluoroglucose, 404
2-deoxyglucose, 398
SUBJECT INDEX
1155
6-deoxyglucose, 403
mercurials, 884
glycogen level, mercurials, 884
glycolysis, 2-deoxyglucose. 392
malonate decarboxylation in, 232
mercurial levels in vivo, 930
mercurial penetration into, 879
mercuric ion,
complexing material from, 907
uptake, 894-897
respiration (endogenous), malonate,
180-181, 187
respiration (glucose), mercurials, 883,
893-894, 898
xylose uptake, mercurials, 911-912
6-Diazo-5-oxo-L-norleucine (DON ),
formylglycinamide ribonucleotide ami-
dotransferase, 333
glutaminase, 356
glutamine:fructose-6-P transamidase,
356
inosinate biosynthesis, 333
phosphoribosyl-PP amidotransferase,
333
structure of, 333
Dibenzamidines,
diamine oxidase, 364
structures of, 361
Dibenzoylthiamine, thiamine kinase, 523
3,5-Dibromobenzoate, dopa decarboxy-
lase, 312-313
3,5-Dibromot>Tosine, tjTosinera-ketoglu-
tarate trasaminase, 306
2,2' - Dicarboxy - 4,4' - diiodoaminoazaben-
zene, determination of SH groups, 641
Dicarboxylate ions,
chelation with cations, 11-13
intercharge distances, 5-7, 44
ionization of, 7-11
kynurenine:a-ketoglutarate transami-
nase, 608
permeability of erythrocytes to, 187-189
pH on dianion concentration, 191
Dichloroacetate, pantoate : /J - alanine li-
gase, 597
Dichloroarabitylflavin, flavokinase, 539
2,4-Dichlorocinnamate, dopa decarboxy-
lase, 313
2.3 - DichloroisobutjTate, pantothenate
biosynthesis, 588
2.4 - Dichloro - 6 - phenylphenoxyethyla-
nine, butynamine demethylase, 592
a,a-Dichloropropionate, pantoate : ^ - ala-
nine ligase, 598
Dichlororiboflavin analog,
L-amino acid oxidase, 540-541
bacterial growth, 537
phosphorylation of, 539
6,7-Dichloro - 9 - (1 '- d - sorbityl) isoalloxa-
zine, carcinostasis, 538
Dichromate, enzyme inhibition, 660
Diethylethoxymethylenemalonate, carci-
nostasis, 202
Diethyl-L-glutamate, L-glutamate dehy-
drogenase, 331
3,3-Diethylglutarate, kynurenine:a-keto-
glutarate transaminase, 608
6,7-Diethylribotlavin analog,
phosphorylation of, 539
structure of, 536
utilization by Lactobacillus, 539
Diethylstilbestrol, /J-hydroxysteroid de-
hydrogenase, 449
Diethylthetin, thetin:homocysteine trans-
methylase, 356
Difluoromalonamide, succinate dehydro-
genase, 239
Difluoromalonate, succinate dehydroge-
nase, 239
Diguanidines,
diamine oxidase, 363-365
structure of, 361
Dihydrofolate reductase,
amethopterin in vivo, 582-583
analogs, 581-584
mercurials, 816
Dihydroorotase, analogs, 470
Dihydroorotatedehydrogenase, analogs, 70
Dihydroxyacetone, phosphopentose iso-
merase, 411
Dihydroxyacetone-phosphate,
enolase, 409
phosphopentose isomerase, 411
Dihydroxybenzoates,
dehydroshikimate reductase, 593, 605-
606
1156
SUBJECT INDEX
dopa decarboxylase, 312
erythro - 2,3 - Dihydroxybutyrate - phos-
phates, enolase, 410
3,4-Dihydroxycinnamate, see Caffeate
5 - (3,4 - Dihydroxycinnamoyi) - salicylate,
dopa decarboxylase, 312
2,2' - Dihydroxy-6,6' - dinaphthyldisulfide
(DDD), determination of SH groups,
921
3,4-Dihydroxyhydrocinnamate, dopa de-
carboxylase, 313
Dihydroxymaleate, catechol oxidase, 297
2,4-Dihydroxy-6-methylpyrimidine, dihy-
droorotate dehydrogenase, 470
6,7-Dihydroxy-7-n-pentyl-8-(r-ribityl)lu-
mazine, riboflavin synthetase, 543
Dihydroxyphenylalanine, see Dopa
Dihydroxyphenylalanine decarboxylase,
see Dopa decarboxylase
2,5-Dihydroxyphenylpyruvate, p-hydro-
xyphenylpyruvate oxidase, 306
6,7-Dihydroxyriboflavin analog, ribofla-
vin synthesis, 539
Diimidotriphosphate, oxidative phospho-
rylation, 448
Diiodothyronines, thyroxine deiodinase,
603
3,5-Diidotyrosine,
dopa decarboxylase, 308
tyrosine: a-ketoglutarate transaminase,
306
Diisopropylfluorophosphonatase, see DFP
ase
Diisothioureas,
diamine oxidase, 363-365
structures of, 361
a,y-Diketovalerate, lactate dehydroge-
nase, 437
Dimercaptides,ofHg++ and thiols, 746-751
6 - ( 2 , 6 - Dimethoxybenzamido) penicilli-
nate, penicillinase, 599, 615, 688
Dimethoxyphenylalanines, dopa decar-
boxylase, 308
Dimethoxyphenylethylamines, dopa de-
carboxylase, 308
Dimethylacetone, glyoxylase, 594
Dimethylacrylate, d -amino acid oxidase,
343
iV^-Dimethyladenine,
adenine deaminase, 466
inosine phosphorylase, 471
iV-(4 - Dimethylamino - 3,5 - dinitrophenyl)
maleimide, determination of SH groups,
641
5,6-Dimethylbenzimidazole,
cyanocobalamin synthesis from, 589
Lactobacillus growth, 589-590
Dimethylglutarates, kynurenine : a - keto-
glutarate transaminase, 608
Dimethylglycine, betaine : homocysteine
transmethylase, 356
Dimethylguanidine, histidase, 353
4,4- Dimethyl - 17 ^ - hydroxyandrost - 5 -
eno(3,2-c) pyrazole, /?- hydroxy steroid
dehydrogenase, 449
iV-Dimethylmalondiamide, carcinostasis,
202
2,2-Dimethylsuccinate, kynurenine:a-ke-
toglutarate transaminase, 608
2,4 - Dimethyl - cyclo - telluropentane - 3,5-
dione,
bacterial growth, 576
structure of, 563
2,4-Dinitrophenol (DNP),
gastric acid secretion, 915-916
mercurial inhibition of ATPase, 869
mitochondrial swelling, 210
porphyrin synthesis, 162
renal transport of PAH, 205
2,6-Dinitrophenol, D-amino acid oxidase,
348
Dinitrophenol reductase, mercurials, 860
^-2,4 - Dinitrophenylpropionate, chymo-
trypsin, 369-370
Diol dehydrase,
cyanocobalamin, 590
hydroxocobalamin, 590
Dioxindole, acetylindoxyl oxidase, 591
Dipeptidase,
analogs, 367-368
o-iodosobenzoate, 708
mercurials, 837
o-Diphenol oxidase, see Catechol oxidase
Diphenylglycolate, glycolate oxidase, 593
Diphenylphosphate, phosphatase (acid),
441
SUBJECT INDEX
1157
Diphosphite, oxidative phosphorylation,
448
2,3-Diphosphoglycerat€, phosphoribomu-
tase, 413
Diplococcus pneumoniae, infection by,
malonate, 221-222
(a,/3 - Distearoyloxypropyl)dimethyl - {(i'-
hydroxyethyl)ammonium acetate, leci-
thinase A, 595
Disulfide groups, role in reactivity of
enzymes with SH reagents, 644-645
Disulfides,
enzyme inhibition, 661-663
mixed, see Mixed disulfides
oxidation of SH groups, 661, 663-664
reduction by dithiothreitol, 640
5,5'-Dithiobis (2-nitrobenzoate), determi-
nation of SH groups, 641
Dithioglycolate,
ATPase, 663
oxidation of protein SH groups, 661
Dithiothreitol, protein disulfide reduction,
640
6,8-Dithiourate, uricase, 285-286
Diuretics, see Mercurial diuretics
Division, see Cell division
DNA, see DeoxjTibonucleates
DNAase, see Deoxyribonuclease
DON, see 6-diazo-o-oxo-L-norleucine
Dopa,
acetylindoxyl oxidase, 591
histidine decarboxylase, 352
kynureninase, 595
metabolism of, pathways, 307
phenylalanine deaminase, 355
tyrosine: a-ketoglutarate transaminase,
306
Dopa decarboxylase,
analogs, 307-320
deoxypyridoxol in vivo, 596-570
folate analogs, 586
Dopamine,
brain levels, a-methyl-w -tyrosine, 316
dopa decarboxylase, 308
metabolism of, 307
tissue levels of, a-methyldopa, 315-320
Dopamine /3-hydroxylase,
analogs, 320
a-methyldopa, 316
Double bond, interaction energy due to
polarization, 276
DPN, see NAD
DT diaphorase, o-iodosobenzoate, 708
dTMP, see Deoxythymidinemonophos-
phate
dTTP, see Deoxythymidinetriphosphate
dUMP, see DeoxjTiridinemonophosphate
Eagle's KB carcinoma, growth of,
deoxypjTidoxol, 577
mercurials, 968
Earle sarcoma cells, respiration (endoge-
nous),
malonate, 177
Eberthella typhosa, o-iodosobenzoate kil-
ling of, 727
Echinococcus granulosus, respiration (en-
dogenous),
malonate, 173
mercurials, 882
Echinus esculentus eggs, respiration (en-
dogenous),
malonate, 175
mercurials, 882
Echinus miliaris, development of,
mercurials, 964
Echo 7 virus, infectivity of,
mercurials, 976-977
Ectromelia virus, inactivation by mercu-
rials, 977
Ehrlich asctes carcinoma cells, see As-
cites carcinoma cells
Elastase, mercurials, 860
Electrical potentials, see Membrane po-
tentials
Electrocardiogram, see Heart, electrocar-
diogram
Embryogenesis, see also Gastrulation and
specific organisms
deoxypyridoxol, 576
ferricyanide, 678
o-iodosobenzoate, 726-727
malonate, 197-199
mercurials, 963-965
1158
SUBJECT INDEX
porphyrindin, 670
Embryos,
glycolysis, hydrogen paroxide, 695
respiration, mercurials, 882
Enantiomers, as analog inhibitors, 268-
271
Encephalitis virus, see Western equine
encephalitis virus
Encephalomyocarditis virus, inactivation
by mercurials, 977
Endamoeha histolytica, growth of,
malonate, 196
Endogenous respiration, see Respiration
(endogenous)
Endomyces vernalis, pyrithiamine-resist-
ant strain, 529
Enolase (phosphopyruvate hydratase),
analogs, 409-410
mercurials, 768, 789, 803, 810, 837
aggregation, 789
rate of inhibition, 810
relation to SH groups, 803
mercuric ion, crystalline complex with,
768
Enoyl-CoA hydratase (crotonase),
o-iodosobenzoate, 707-708
mercurials, 837
Enteroviruses, adsorption to kidney cells,
mercurials, 981
Enzymes,
biosynthesis of,
D-asparagine, 269
azatryptophan, 326
fluorophenylalanine, 351
5-methyltryptophan, 326
tryptazan, 326
induction of,
8-azaguanine, 478
azatryptophan, 326
malonate, 155-156
mercurials, 888
a-methyltryptophan, 325
Epidermis, mitosis in,
malonate, 199-200
mercurials, 968
Epidermophyton floccosum, respiration
(endogenous),
malonate, 169
Epididymal fat pad, glucose oxidation,
D-glucosamine, 382
Epinephrine, see also Catecholamines
cardiac stimulation,
malonate, 217
mercurials, 947
2-deoxyglucose phosphorylation, 387
dopa decarboxylase, 308
formation of, pathways, 307
release of, 2-deoxyglucose, 401
responses to, pjTogallol, 611
tissue levels of, a-methyldops, 315-320
tyrosi ne : a - ketoglutarate transaminase ,
305-306
urinary excretion, pyrogallol, 612
Epinine, phenylalanine /5-hydroxylase,
600
Equilin dehydrogenase, o-iodosobenzoate,
708
Ergot, see Claviceps
Erythredema, see Acrodynia
Erythrocytes,
amino acid transport, competition by
sugars, 267
carbohydrate uptake, competition be-
tween sugars, 264
glucose uptake, mercurials, 903-905,
911
glucose utilization, cystamine, 663
GSH level, mercurials, 905-906
heme biosynthesis, mercurials, 888
iodine hemolysis, 690
K+ efflux, mercurials, 903-905
K+ influx, mercurials, 908
K+ trasport, malonate, 209
membrane of, mercurials, 906
mercuric ion uptake, 897, 900-907
Na+ transport, malonate, 209
nonelectrolyte transport, iodine, 690
permeability,
mercurials, 900-907
to dicarboxylates, 187-189
porphyrin biosynthesis,
malonate, 159-163
mercurials, 888
protoporphyrin biosynthesis,
arsenite, 162
2,4-dinitrophenol, 162
SUBJECT INDEX
1159
fluoroacetate, 162
malonate, 162
SH groups in, 897
urate transport, hypoxanthine, 267
Erytlu-ocytes (Plasmodium -parasitized),
ATP level, quinacrine, 560
glucose utilization, quinacrine, 560
respiration (glucose),
malonate, 124
quinacrine, 560
succinate accumulation, malonate, 91,
93
Erythrose-4-phosphate, phosphoglucose
isomerase, 407
Escherichia coli,
adaptive enzyme synthesis, malonate,
155
azatryptophan incorporation into en-
zymes of, 326
cycle intermediates concentrations, 89
2 -deoxy glucose uptake, 387
glutamate dehydrogenation, malonate,
152
glycolysis, 2-deoxyglucose, 387
grov/th of,
azatryptophan, 326
biotin analogs, 588
dehydroacetate, 632
2-deoxyglucose, 400
malonate, 195
mercurials, 972
methylindoles, 321, 323
methylthio analog of thiamine, 530
4-methyItryptophan, 323
D-phenylalanine, 268
o-iodosobenzoate killing of, 727
a-ketoglutarate oxidation, malonate, 79
lactate oxidation, malonate, 78
malate oxidation, malonate, 81
malonate metabolism in, 228
mercurial uptake, 974-975
oxidative phosphorylation, malonate,
120
protein biosynthesis, 5-fluorouracil, 479
pyruvate oxidation,
malonate, 74
mercurials, 878
resistance to mercurials, 983-984
succinate dehydrogenase,
adipate, 35
glutarate, 35
malonate, 2, 21, 26, 187
oxalate, 35
tartronate, 36
succinate oxidation, malonate, 52
tryptazan incorporation into enzymes
of, 326
Escherichia coli phage, see Coliphage
Esterase (microsomal), stimulation by
mercurials, 816
Esterases, see individual enzymes
Estradiol,
cholesterol esterase, 592
/9-hydroxysteroid dehydrogenase, 447,
449
Estradiol- 17/?-dehydrogenase, mercurials,
protection by estradiol, 781
protection by NAD, 781
Estra-l,3,5-trienes, ^-hydroxysteroid de-
hydrogenase, 449
Ethane, nitrogen fixation, 291
1,2-Ethenediphosphonate,
ionization constants, 242
succinate dehydrogenase, 243
1 ,2-Ethanedisulfonate,
aspartase, 355
inter charge distance, 7
succinate dehydrogenase, 242-243
Ethanesulfonate, sulfite oxidase, 451
Ethanol, oxidation of,
2-deoxyglucose, 395-396
mercurials, 898
Ethanolamine, choline oxidase, 290
Ethanolamine oxidase, quinacrine, 547,
550
Ethyl- 1 -acetyl - 2 - benzylcarbazate, chy-
motrypsin, 373
2-Ethyl-3-amino-4-ethoxymethyl-5-ami-
nomethylpyridine, pyridoxal kinase,
564
Ethylenediamine, diamine oxidase, 362
Ethyl-D-glutamate,
glutaminase, 333
urinary flow, 333
iV-Ethyl-DL-leucine, D-amino acid oxi-
dase, 340
r
1160
SUBJECT INDEX
Ethylmalonate,
biosynthesis of, 226
carcinostasis, 201
lethal dose, 201
occurrence of, 225
6-Ethyl-8-mercaptooctanoate, acyl trans-
fer, 590
Ethylmercuri-p-toluene sulfonanilide, see
Granosan M
3 - Ethyl-4-methyIthiazole, thiaminase,
524
Ethyloxalacetate, malic enzyme, 597
iV^-Ethyl-DL-phenylalanine, D-amino acid
oxidase, 340
<S-(iV-Ethylsuccinimido)-GSH,glyoxylase,
593
2'-Ethylthiamine, thiamine kinase, 523
a - Ethylthioglucopyranoside, a - gkicosi-
dase, 423
Euglena gracilis,
acetate oxidation, malonate, 77
growth of, pyrithiamine, 529
malonate metaboHsm in, 228
succinate dehydrogenase, malonate, 28
succinate oxidation, malonate, 51, 53,
56
Exopenicillinase, iodine, 688
FAD (flavin adenine dinucleotide),
analogs of, see Riboflavin, analogs of
L-lactate oxidase, 543
lysolecithin oxidase, 543
metabohsm of, pathways, 535
NADH : ferricyanide oxidoreductase,
510
succinate oxidase, 543
FAD pyrophosphorylase,
isoriboflavin, 542
riboflavin, 542
Fagus, see Beech
False feedback inhibition, 321
Fasciola hepatica, respiration (endoge-
nous),
malonate, 173
Fats, see Lipids
Fatty acids,
biosynthesis of,
acyl-CoA analogs, 613-614
deoxypyridoxol, 574
malonate, 146-149
mercurials, 887
a-phenylbutyrate, 614
propionate, 613
formation from malonate, 234
formation from propionate, malonate,
146
kynurenine:a-ketoghitarate transami-
nase, 608-609
pH effects, 609
oxidation of,
benzoate, 349
2-deoxyglucose, 397
malonate, 135-137, 141-143
mercurials, 887
plasma levels, 2-deoxyglucose, 399
Fatty acid synthetase, mercurials,
protection by acetyl-CoA, 781
Fatty acid thiokinase, mercurials, 887
Feedback inhibition,
analogs, 351
anthranilate synthesis, 321
false, 321
glutamate dehydrogenase, 514
pyrimidine metabolism, 478-481
Fermentation, see also substances fer-
mented and Yeast, fermentation
biotin analogs, 588-589
D-glucosone, 384-385
iodine, 689
mercurials, 875
tripolyphosphate, 383
Ferric ions, see also Iron
glutamate dehydrogenase, 863
Ferricyanide, 670-678
Aspergillus growth, 677
chemical properties, 670-671
citrate accumulation, 677-678
enzyme inhibitions, 672-676
glucose oxidation, 677-678
glycolysis, 673, 677
itaconate metabolism, 678
mitochondrial swelling, 678
NADH oxidation by, 673
oxidation of,
SUBJECT INDEX
1161
amino acids, 672
NADH, 673
protein SH groups, 670-672
SH groups, 670-671
oxidation-reduction potential, 670-671
pentose-P pathway, 677
phospholipid biosynthesis, 678
porphyrin biosynthesis, 678
purification of, 671
respiration, 678
sea urchin egg development, 678
yeast growth, 678
Ferrocyanide,
isocitrate dehydrogenase, 677-678
tricarboxylate cycle, 677-678
Fertilization,
o-iodosobenzoate, 726-727
malonate, 198
mercurials, 963-964
Ferulate, peroxidase, 599
Fibroblasts, growth of,
mercurials, 968-969
Fibrosarcoma, growth of,
deoxypyridoxol, 576
Ficin, mercurials,
dimeric complex with Hg++, 770
relation to SH groups, 804
Flagellar motility,
o-iodosobenzoate, 727
malonate, 203
Flavin adenine dinucleotide, see FAD
Flavin mononucleotide, see FIVIN
Flavins,
analogs of, see Riboflavin, analogs of
tissue levels, galactoflavin, 539-540
Flavobacterium,
protocatechuate transport, p-aminosa-
Ucylate, 267
renal transport of PAH, p-aminosalicy-
late, 613
Flavokinase (riboflavin kinase),
analogs, 539
mercurials, 854
permanganate, 660
Flavotin,
structure of, 537
succinate oxidase, 543
Flexner-Jobling tumor,
malonate levels in vivo, 100-102
succinate accumulation, malonate, 100-
103
Fluoride,
/?-ketoadipate chlorinase, 453
phosphorylase, 406
renal transport of PAH, 205
tjTosinase, 300
Fluoride dimer, phosphatase (acid), 441-
442
Fluorine, use in forming analogs, 258-259
Fluoroacetate,
lethal doses, 239
porphyrin biosynthesis, 162
renal transport of PAH, 205
sequential inhibition with malonate,
112
Fluoroacetyl-L-tyrosinate, chymotrypsin,
371
Fluoroamino acids, amino acid and
protein metabolism, 351
Fluorobenzoates, D -amino acid oxidase,
341
5-Fluorocytidine, aspartate carbamyl-
transferase, 467
5-Fluorodeoxycytidine, aspartate carba-
myltransferase, 469
5-Fluorodeoxyuridine, thymidylate syn-
thetase, 476
5-Fluorodeoxyuridinemonophosphate
(FdUMP), thymidylate synthetase, 47( ,
479
/3-Fluoro-DL-malate,
fumarase, 279
malate dehydrogenase, 279
Fluoromalonate,
citrate accumulation, 239
decarboxylation of, 239
lethal doses, 239
succinate dehydrogenase, 239
Fluoromalonic diethyl ester, lethal doses,
239
5-Fluoroorotate,
aspartate carbamyltransferase, 468
conversion to 5-fluoro-UMP, 478
dhydroorotase, 470
UTP biosynthesis, 478-479
Fluorooxalacetate,
1162
SUBJECT INDEX
aspartate : a - ketoglutarate transami-
nase, 334
malate dehydrogenase, 596
transamination to fluoroaspartate, 334
p- Fluoropheny lalanine ,
incorporation into proteins, 351
maltase biosynthesis in yeast, 351
L-phenylalanine:sRNA Ugase (AMP),
354
protein biosynthesis, 351
replacement of phenylalanine in pro-
teins, 351
;5-FluorophenylaIanine, tyrosine transport
by brain, 266
Fluorophosphate, see Monofluorophos-
phate
Fluoropyrimidines, pyridimidine meta-
bohsm, 478-481
5 - Fluor otryptophan ,
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
6-Fluorotryptophan,
anthranilate metabohsm, 321
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
3-Fluorotyrosine,
tyrosinase, 304-305
tyrosinera-ketoglutarate transaminase,
305-306
L-tyrosine:sRNA ligase (AMP), 307
5-Fluorouracil,
abnormal enzymes produced by, 479-
480
protein biosynthesis, 479
thymidylate synthetase, 476
5-Fluorouridine, thymidylate synthetase,
476
5-Fluorouridinemonophosphate (FUMP),
thymidylate synthetase, 476
FMN (flavin mononucleotide),
D-amino acid oxidase, 540-541
glutamate racemase, 542
L-lactate oxidase, 543
NADH:ferricyanideoxidoreductase,510
riboflavin transglucosidase, 543
succinate oxidase, 543
Folate,
analogs, of, 579-586
acetylations, 586
ATP level in tissues, 585
enzyme inhibitions, 586
folate deficiency, 581
folate reduction, 581-584
nucleic acid biosynthesis, 584-585
protein biosynthesis, 585
structures of, 580
metabolic functions of, 579
metabolism of, pathways, 579
photolytic oxidation products as in-
hibitors of xanthine oxidase, 285-287
structure of, 580
Folate reductase, see Dihydrofolate reduc-
tase
Folinate (citrovorum factor)
formation from folate, analogs, 582
structure of, 580
Foot-and-mouth virus, proliferation of,
malonate, 194
Formaldehyde, pyruvate decarboxylase,
432, 600
Formaldehyde polymers, hyaluronidase,
459-461
Formate,
glutamate decarboxylase, 328
kynurenine:a-ketoglutarate transami-
nase, 608
lactate dehydrogenase, 436
tyrosinase, 300
Formate dehydrogenase, hypophosphite,
593
Formate hydrogenlyase,
hypophosphite, 593
o-iodosobenzoate, 708
Formate transacetylase, ferricyanide, 675
Formylglycinamide phosphoriboside syn-
thetase, analogs, 333
9-Formylmethylriboflavin, flavokinase,
539
6-Formylpteridine, see Pterin-6-aldehyde
Form yltetrahydro folate synthetase, ana-
logs, 585
A'^-Formyl-L-tyrosine, tyrosinase, 304-305
N - Formyl - L- tyrosinemethylamide, chy-
motrypsin, 371
Fowl plague virus, infectivity of,
mercurials, 977, 980
SUBJECT INDEX
1163
Frog eggs, protein biosynthesis,
mercurials, 887
^-Fructofuranosidase (invertase),
analogs, 421
dichromate, 660
ferricyanide, 675
hydrogen peroxide, 691-692
iodine, 683, 685, 688
macroions, 465
mercurials, 837-838
increase of inhibition by thiols, 827-
828
pH effects, 791-793
protection by sucrose, 781
rate of inhibition, 811-812
spontaneous reversal, 813
type of inhibition, 772
methylglucoside anomers, 271
oxidation of, 657
periodate, 660
permanganate, 660
succinyl peroxide, 694
/3-Fructofuranosyl-2-deoxyglucose, forma-
tion from 2-deoxyglucose, 389
Fructokinase, see also Hexokinase
analogs, 376
galactose, 376
glucose, 376
o-iodosobenzoate, 708
mannose, 376
mercurials, 837
Fructose,
aldolase, 407
/^-amylase, 421
/3-fructofuranosidase, 421
a-galactosidase, 418
glucosamine phosphorylation 382
glucose uptake by lymph node, 263
a-glucosidase, 416
intestinal transport of, deoxypyridoxol,
574
a-mannosidase, 422
oxidation of, 2-deoxyglucose, 398
phosphopentose isomerase, 411
uptake by lymph node, 2-deoxyglu-
cose, 394
Fructose-l,6-diphosphatase,
induction of, 8-azaguanine, 478
malonate, 60
mercurials, 837
nucleotides, 470
Fructose-l,6-diphosphate,
glucose dehydrogenase, 410
hexokinase, 379
2-keto-3-deoxy-D-ara6o-heptonate-7-P
synthetase, 413
Fructose-phosphate, aldolase, 407
Fructose-6-phosphate,
aldolase, 407
glucose dehydrogenase, 410
hexokinase, 379
phosphodeoxyribomutase, 413
phosphopentose isomerase, 411
Fructose-phosphates, structures of, 378
Fucono-l,5-lactone, /J-galactosidase, 429
Fucose, a-galactosidase, 417
Fucus ceranoides,
iodide uptake, mercurials, 910, 912
respiration (endogenous), mercurials,
881, 912
Fumarase (fumarate hydratase, L-malate
hydro-lyase),
trans-Rconitate, 273
active center of, 274, 278
analogs, 274-279
binding energies of, 275-277
pH effects, 277-279
a-hydroxy-/3-sulfopropionate, 243
o-iodosobenzoate, 708, 717
protection by substrate, 717
macroions, 465
malonate, 60, 65
mercurials, 778, 781, 838
protection by phosphate, 778
protection by substrates, 781
Fumarate,
D-amino acid oxidase, 343
antagonism of malonate inhibitions
112-117, 135
aspartase, 355
glutamate decarboxylase, 328
glutamate dehydrogenase, 330-332
intercharge distance, 6
ionization constants, 8
D-lactate dehydrogenase, 437
malic enzyme, 596
1164
SUBJECT INDEX
oxalacetate decarboxylase, 597
oxidation of, malonate, 81
phosphofructokinase, 385
reduction of, malonate, 48-49
succinate dehydrogenase, 34-35, 38
Fumarate reductase, malonate, 49
Fundulus heteroclitus eggs, cleavage of,
mercurials, 963
Fungi,
growth of,
dehydroacetate, 632-633
2-deoxyglucose, 400
ferrocyanide, 677
malonate, 195-196
oxythiamine, 520, 529
pyrithiamine, 516, 528-529
thiamine analogs, 516, 520, 528-529
tungstate, 614
respiration (glucose), malonate, 133-
134
Furan-2-acrylate, D-amino acid oxidase,
342
Furan-2-carboxylate, D-amino acid oxi-
dase, 242, 346
2-Furoate,
D-araino acid oxidase, 344
glutamate dehydrogenase, 331
structure of, 330
Furyglycine, glycine uptake by ascites
cells, 265
Fusarium, conidial growth,
p-benzoquinone, 660
permanganate, 660
Fusarium decemcellulare, mercurial accu-
mulation in condia, 969
Fusarium graminearum , growth of,
dehydroacetate, 632
GABA, see y-Aminobutyrate
Galactarate (mucate), /3- glucuronidase,
424, 427
Galactoflavin,
carcinostasis, 538
flavin levels in liver, 539-540
flavokinase, 539
glutamate oxidation in vivo, 544
/5-hydroxybutyrate oxidation in vivo,
544
oxidative phosphorylation in liver, 544
riboflavin deficiency, 538
serotonin metabolism, 544
structure of, 536
succinate oxidase, 543
Galactono-l,5-lactone, ^-galactosidase,
429
L-Galactono-y-lactone dehydrogenase,
mercurials, protection by substrate, 781
quinacrine, 547, 550
riboflavin, 540, 542
Galactose,
^-amylase, 421
fructokinase, 376-377
fructose uptake by ascites cells, 263
a-galactosidase, 417-418
/?-galactosidase, 418
glucose uptake by lymph node, 263
a-glucoidase, 416-417
intestinal transport of,
analogs, 263
6-deoxyglucose, 403
mutarotase, 413-414
oxidation of, see also Respiration (ga-
lactose)
2-deoxyglucose, 398
renal transport of, glucose, 262
uptake of, 2-deoxyglucose, 394
Galactose dehydrogenase, ferricyanide,
675
Galactose oxidase, quinacrine, 550
Galactose- 1 -phosphate, UDPglucose py-
rophosphorylase, 603
Galactose-6-phosphate,
hexokinase, 379-380
phosphodeoxyribomutase, 413
a - Galactosidase ,
analogs, 417-418
5-fluorouracil, inactivation in vivo 479-
480
inactive form induced by, 479-480
mercurials, 838
/3-Galactosidase,
analogs, 418-419, 429
hydrogen peroxide, 691-692
iodine, 683, 685
SUBJECT INDEX
1165
mercurials, 838
spontaneous reversal, 814
synthesis of,
azatryptophan, 326
2-deoxyglucose, 400
jralacturonate,
a-glucuronidase, 426
^-glucuronidase, 424, 427
polygalacturonase, 421
Vallate, dehydroshikimate reductase, 593,
605
Ganglia,
acetylcholine response, mercurials, 949
transmission through,
malonate, 211-212
mercurials, 949
Gardner lymphosarcoma, amino acid up-
take,
malonate, 155
Gases, see also specific gases
hydrogenase, 293-294
nitrogen fixation, 291-296
Gasterosteus aculeatus, mercurials,
respiration (endogenous), 882
toxicity, 963
Gastric acid secretion,
antimycin A, 915-916
2,4-dinitrophenol, 915-916
malonate, 187, 208
mercurials, 914-915
Gastric mucosa,
respiration ( endogenous ), malonate,
175, 179
respiration (glucose), mercurials, 883
Gastrula, respiration (endogenous),
mercurials, 882
Gastrulation see also Embryogenesis
malonate, 198-199,
mercurials, 964
GDP, see Guanosinediphosphate
Germination,
Aspergillua spores, malonate, 195
Bacillus cereus, D-alanine, 270
Bacillus subtilis,
malonate, 195
mercurials, 972
quinacrine, 546
fungi, effect on malonate inhibition of
respiration, 133-134
Neurospora ascospores, malonate, 195
Puccinia uredospores, malonate, 195-
196
Glucarate,
/?-glucuronidase, 424, 427
glucuronide synthesis in liver, 428
phosphatase (acid), 442-443
Glucaro-l,4-3,6-dilactone, structure of,
425
Glucaro- 1 ,4-lactone,
/^-glucuronidase, 423-428
in vivo inhibition, 428
glucuronide formation in liver, 428
structure of, 425
Glucaro-3,6-lactone,
/^-glucuronidase, 424, 427
structure of, 425
/S-Glucofuranuronides, structures of, 425
Glucokinase, see also Hexokinase
iV-acetyl-D-glucosamine, 390
2-deoxyglucose, 389-390
D-glucosamine, 390
mannose, 376
Gluconate, /3-glucosidase, 417
Gluconate ethyl ester, ^-glucosidase, 417
D-Gluconate oxidase, riboflavin, 542
Gluconate-6-phosphate dehydrase, mer-
curials, 885
Gluconate - 6 - phosphate dehydrogenase,
mercurials, 838
Gluconic-y-lactone, /?-glucosidase, 417
Gluconokinase, mercurials, 781, 839
protection by ATP, 781
protection by gluconate, 781
Glucono- 1 ,4-lactone,
cellulytic rumen enzymes, 429
a-glucuronidase, 426
/3-glucosidase, 429
isoamylase (debranching), 429
structure of, 425
thioglycosidase, 429
Glucono-l,5-lactone,
a-glucuronidase, 426
/3-gluco8idase, 429
/3-Glucopyranuronides, structures of, 425
Glucosaccharo-l,4-lactone, see Glucaro-
1,4-Iactone
1166
SUBJECT INDEX
D-Glucosamine,
ATP level in ascites cells, 383
glucokinase, 390
glucose oxidation in fat pad, 382
glucose uptake by Scenedesmus, 383
glycogen formation in liver, 382
hexokinases, 381-383, 390
phosphoglucomutase, 382
phosphopentose isomerase, 411
phosphorylase, 382
phosphorylation of,
iV-acetylglucosamine, 382
hexoses, 382
pyruvate oxidation, 383
structure of, 381
UDPglucose-glycogen glucosyltransfe-
rase, 382
UDPglucose pycophosphorylase, 382
D-glucosamine-6-phosphate,
glucose-6-P dehydrogenase, 411
phosphoglucose isomerase, 407
D-Glucosamine-6-phosphate deaminase,
mercurials, 816
D-Glucosamine phosphokinase, iV-acetyl-
glucosamine, 593
Glucose,
aldolase, 407
a-amylase, 420
j3-amylase, 421
arabinose uptake by heart, 263
blood levels of, malonate, 149, 219
2-deoxyglucose uptake, 387, 389
a-l,6-dextranglucosidase, 417
distribution of C" from labeled, ma-
lonate, 130-132
fermentation by yeast,
D-glucosone, 384-385
tripolyphosphate, 383
/3-fructofuranosidase, 421
fructokinase, 376-377
galactose transport by kidney, 262
a-galactosidase, 417
/5-galactosidase, 418
glucosamine phosphorylation 382
glucose-6-phosphatase, 412
a-glucosidase, 416-417, 423
/3-glucosidase, 417
intestinal transport of,
deoxypyridoxol, 574
malonate, 207
mercurials, 916
a-mannosidase, 422
metabolism of,
malonate, 122-135
malonic diethyl ester, 236-237
oxidation of, see also Respiration (glu-
cose)
6-deoxy-6-fluoroglucose, 393
2-deoxyglucose, 393, 398
6-deoxyglucose, 403
D -glucosamine, 382
phosphopentose isomerase, 411
phosphorylase, 405
renal transport of,
dehydroacetate, 625
malonate, 205
mercurials, 920
uptake of,
analogs, 263
6-deoxy-6-fluoroglucose, 404
2-deoxyglucose, 390, 394
mannoheptulose, 376
mercurials, 893-894, 910-912
utilization of,
cystamine, 663
dehydroacetate, 624
ferrocyanide, 678
mercurials, 884, 903-905
oxamate, 435
quinacrine, 560
Glucose dehydrogenase,
analogs, 410, 500-502
benzoate, 501
malonate, 61
mercurials, 838
nicotinamide analogs, 500-502
nucleotides, 501
pyridoxal, 501-502
quinacrine, 550
Glucose - 1 ,6 - diphosphates, hexokinase,
379
Glucose oxidase, pterin-6-aldehyde, 288
Glucose-6-phosphatase,
analogs, 412
2-deoxyglucose, 390
induction of, 8-azaguanine, 478
SUBJECT INDEX
1167
mercurials in vivo, 927
Glucose- 1 -phosphate,
glucose dehydrogenase, 410
hexokinase, 379-380
level in parasitized erythrocytes, qui-
nacrine, 560
phosphatase, 439
phosphorylase, 405
Glucose-6-phosphate,
fructokinase, 381
glucose dehydrogenase, 410
hexokinase, 377, 379, 381
phosphoarabinose isomerase, 411
phosphodeoxyribomutase, 413
phosphoglucomutase, 413
phosphopentose isomerase, 411
phosphorylase, 405
Glucose-6-phosphate dehydrogenase,
3-acetylpyridine-NAD, 497
analogs, 411
l,5-anhydro-D-glucitol-6-P, 379
2-deoxyglucose-6-P, 390-391
o-iodosobenzoate, 608
mercurials, 839
protection by NADP, 781
nicotinamide, 503
nucleotides, 508
quinacrine, 550
Glucose-phosphate isomerase, see Phos-
phoglucose isomerase
Glucose-phosphates, structures of, 378
Glucose respiration, see Respiration (glu-
cose)
a-Glucosidase, see also Maltase
analogs, 416-417, 423
mercurials, 772
/3-Glucosidase,
analogs, 417
mercurials, 839
protection by substrates, 782
D-Glucosone,
Crabtree effect, 385, 397
formation of, 385
glucose fermentation in yeast, 384-
385
glycolysis (anaerobic), 385
hexokinase, 384-385
hyperglycemia, 384
lethal doses, 384
structure of, 383-384
toxicity, 384
L-Glucosone, glucose fermentation in
yeast, 384
Glucuronate,
a-glucuronidase, 426
/^-glucuronidase, 424, 427
hexokinase, 380
phosphatase (acid), 442
structure of, 425
Glucuronate - 1 - phosphate, a - glucuroni-
dase, 426
Glucuronate-6-phosphate, hexokinase,
380
D-Glucurone,
a-glucuronidase, 426
/^-glucuronidase, 424, 427
glucuronide synthesis in liver, 428
structure of, 425
a - Glu curonida se ,
analogs, 426
D-glucurone, 426
a-and /J-glucuronides, 426
/S-Glucuronidase,
analogs, 423-428
copper, 795
inhibition in urinary bladder cancer,
428
malonate, 61
menthyl-a-glucuronide, 272
mercurials, 839
pH effects, 791, 793, 795-797
type of inhibition, 772
silver, 795
Glucuronidases,
heparin, 465
hyaluronate, 465
Glutamate,
cysteine desulfurase, 357
glutaminase, 332
metabolism of,
analogs, 327-336
malonate, 152
pathways, 327
oxidation of,
galactoflavin in vivo, 544
mercurials, 878
1168
SUBJECT INDEX
D-Glutamate,
L-glutamate decaroxylase, 269
L-glutamate dehydrogenase, 330, 332
L-glutamine synthetase, 269, 336
y-glutamyltransferase, 336
L-Ghitamate,
L-amino acid oxidase, 340
D-glutamate oxidase, 336
phosphatase (acid), 441-442
pyridoxamine - oxalacetate transami-
nase, 600
L-Glutamate : ammonia Hgase (ADP), see
Glutamine synthetase
L-Glutamate decarboxylase,
acetate, 328
analogs, 327-329
cafFeate, 314
D- cycloserine, 359
deoxypyridoxol in vivo, 569
D-glutamate, 269
mercurials, 772, 782, 811-812
protection by pyridoxal-P, 782
rate constant for inhibition, 811-812
temperature effects, 811
type of inhibition, 772
permanganate, 660
toxopyrimidine, 578
L-Glutamate dehydrogenase,
analogs, 329-332
ferric ions, 863
o-iodosobenzoate, 708
p-iodosobenzoate, 702
malonate, 61
mercurials, 789, 793, 816, 819, 826,
840-841, 863
pH effects, 793
reversal with GSH, 826
splitting into subunits, 789, 819
stimulation, 816, 819
nicotinamide, 863
nucleotide binding sites, 514
nucleotides, 508, 514
oxygen inactivation of, 659
silver, 863
zinc, 863
D-Glutamate oxidase,
benzoate, 349
L-glutamate, 336
o-iodosobenzoate, 709
mercurials, 840
Glutamate racemase,
FMN, 542, 544
riboflavin, 542, 544
Glutamate semialdehyde reductase,
o-iodosobenzoate, 709, 717
protection by substrate, 717
mercurials, 841
nucleotides, 507
Glutamate transaminases, see also Trans-
aminases
analogs, 334
Glutaminase (L-glutamine amidohydro-
lase),
active center of, 332
analogs, 332-333, 356
mercurials, 841
quinacrine, 550
Glutaminase I, quinacrine, 550
Glutaminase II, quinacrine, 551
L-Glutamine,
brain level, 2-deoxyglucose, 399
L-glutamate dehydrogenase, 331
L-Glutamine amidohydrolase, see Gluta-
minase
Glutamine : fructose - 6 - phosphate trans-
amidase, DON, 356
Glutamine:pyruvate transaminase, see
Transaminases
Glutamine synthetase (L-glutamate:am-
monia ligase),
ADP, 471
analogs, 336
D-glutamate, 269
GSSG, 662
mercurials, 841
methionine sulfoximine, 335
y-Glutamylalanylglycine, glyoxylase, 594
y-Glutamyl-/3-sulfoalanylglycine, glyoxyl-
ase, 594
y-Glutamyltransferase,
analogs, 336
malonate, 61
Glutarate,
aspartate : a - ketoglutarate transami-
nase, 334
carcinostasis, 201
SUBJECT INDEX
1169
cysteine desulfurase, 357
fumarase, 275
glutamate decarboxylase, 328
glutamate dehydrogenase, 330-332
hydroxamic acid formation from, 233
intercharge distance, 5-6
ionic length, 188
ionic volume, 188
ionization constants, 8
kynureninera-ketoglutarate transami-
nase, 607-609
lethal doses, 201
permeability of erythrocytes to, 188
pyridoxamine: oxalacetate transami-
nase, 600
succinate dehydrogenase, 35
urinary citrate, 106, 109
urinary a-ketoglutarate, 110-111
Glutathione,
erythrocyte levels, mercurials, 905-906
glycolysis (aerobic), 637
role in enzyme and metabolic activity,
636-637
tissue levels, tetrathionate, 696, 700
Glutathione (oxidized),
enzyme inhibitions, 662-663
succinate dehydrogenase, temperature
effects, 663-664
Glutathione oxidase, thioglycolate, 593
Glutathione reductase, mercurials, 773,
841
Gly ceraldehy de ,
glucose phosphorylation, 377
glycolysis, 377
Glyceraldehyde-3-pho8phate, see 3- Phos-
phoglyceraldehyde
Glyceraldehyde-3-phosphate dehydrogen-
ase, see 3-Phosphoglyceraldehyde de-
hydrogenase
Glycerate, phosphatase (acid), 441
Glycerate dehydrogenase,
bromopyruvate, 430
mercurials, 817, 841
phenylpyruvate, 430
pyruvate, 430
Glycerate-2,3-diphosphatase,
mercurials, stimulation, 817-818, 820
phosphoglycerates, 413
Glycerate kinase,
o-iodosobenzoate, 709
mercurials, 841
GIycerate-2-phosphatase, mercurials, 817
Glycerate-3-phosphatase, mercurials, 817
Glycerol,
metabolism of, malonate, 164
permeability to, mercurials, 906-907
Glycerol-2-phosphate, enolase, 409
Glycerol-3-phosphate dehydrogenase,
hydrogen peroxide, 693
mercurials, relation to SH groups, 804
Glycerol-3-phosphate oxidase, hydrogen
peroxide, 693
/5-Glycerophosphatase,
dichromate, 660
iodine, 685
mercurials, 860
inhibition in vivo, 926-927
permanganate, 660
/?- Glycerophosphate, phosphatases, 439
a-Glycerophosphate dehydrogenase,
malonate, 61
mercurials, 842
porphyrexide, 668
Glycine,
L-alanine dehydrogenase, 354
L-amino acid oxidase, 340
arginase, 337
dipeptidase, 368
intestinal transport of, analogs, 265
phosphatase (acid), 441
Glycine iV-acylase, hippurates, 355
Glycine:a-ketoglutarate transaminase, see
Transaminases
Glycine methyltransferase, o-iodosoben-
zoate, 709
Glycogen,
formation of,
2-deoxyglucose-6-P, 391
D-glucosamine, 382
level in diaphragm, mercurials, 884
Glycolaldehyde-2-phosphate, formation of
threose-2,4-diphosphate from, 408
Glycolate (hydroxyacetate),
glyoxylate transacetatse, 594
tartronate semialdehyde reductase, 602
1170
SUBJECT INDEX
Glycolate oxidase,
analogs, 438
diphenylglycolate, 593
o-iodosobenzoate, 709
malonate, 61
mercurials, 860
Glycolysis, see also Glycolysis (aerobic)
and Glycolysis (anaerobic)
o-iodosobenzoate, 704
ionic regulation of, 453
malonate, 125-130, 134-135
effect of fumarate, 126
mercurials, 874-877, 884
role of GSH in, 637
tetrathionate, 699
threose-2,4-diphosphate, 409
Glycolysis (aerobic),
2-deoxyglucose, 391-394
ferricyanide, 677
hydrogen peroxide, 695
oxamate, 434
quinacrine, 560
tartronate, 238
Glycolysis (anaerobic),
dehydroacetate, 624
2-deoxyglucose, 391-392
ferricyanide, 673, 677
hydrogen peroxide, 695
iodine, 689
o-iodosobenzoate, 721
macroions, 465
oxalate, 414
oxamate, 434
phosphate, 414
ribonucleonate, 414
sulfate, 414
tartronate, 238
Glycosidase, analogs, 415-429
a,/3-Glycosylphosphatase, mercurials, 842
Glycylglycine, D-amino acid oxidase, 340
Glycylglycine dipeptidase, mercurials, 842
Glycylleucine, D-amino acid oxidase, 340
Glycylphenylalanine, cathepsin C, 375
Glycyltyrosine, carboxypeptidase, 367
Glyoxal, tartronate semialdehyde reduc-
tase, 602
Glyoxalate, see Glyoxylate
Glyoxylase, analogs, 593-594
Glyoxylate,
cycle inhibition by formation of y-
hydroxy-a-ketoglutarate, 615-616
metabolism of, malonate, 165
pyruvate decarboxylase, 431
tartronate semialdehyde reductase, 602
Glyoxylate cycle, role in malonate inhibi-
tion of tricarboxylate cycle, 71-72
Glyoxylate reductase,
analogs, 438
malonate, 61
mercurials, 842
protection by glyoxylate, 782
protection by NADH, 782
Glyoxylate transacetase, analogs, 594
GMP, see Guanosinemonophosphate
Goldfish gills, Na+ transport,
mercurials, 909
Granosan M, toxicity, 954
Growth, see also specific organisms and
tissues
2-deoxyglucose, 400-401
malonate, 196-202
mercurials, 963-970
thiamine analogs, 527
GSH, see Glutathione
GSSG, see Glutathione (oxidized)
GTP, see Guanosinetriphosphate
GTPase, see Guanosinetriphosphatase
Guaiacol, dehydroshikiniate reductase,
593, 605
Guanase (guanine deaminase),
pterin-6-aldehyde, 288
xanthopterin, 288
Guanidine, histidase, 353
Guanidines,
diamine oxidase, 360-365
structures of, 361
Guanidinium ion, resonance structures of,
361
Guanidinovalerate, carboxypeptidase, 367
Guanine,
adenylosuccinate synthetase, 467
ATPase, 445
Guanine deaminase, see Guanase
Guanosine,
adenylosuccinate synthetase, 467
5 '-nucleotidase, 472
SUBJECT INDEX
1171
Guanosinediphosphate (GDP),
adenylosuccinate synthetase, 467
glutamate dehydrogenase, 508
IMP dehydrogenase, 471
isocitrate dehydrogenase, 509
NADH oxidase, 511
polynucleotide phosphorylase, 474
Guanosinemonophosphate (GMP),
adenylosuccinate synthetase, 467
deoxycytidylate deaminase, 469
IMP dehydrogenase, 471
phosphatase, 439
ribonuclease, 475
Guanosinemonophosphate reductase, nu-
cleotides, 471
Guanosinemonophosphate sjTithetase, see
Xanthosine-5 '-phosphate aminase
Guanosine phosphorylase, mercurials, 842
Guanosinetriphosphatase (GTPase),
ADP, 446
IDP, 446
Guanosinetriphosphate (GTP),
asprtate carbamyltransferase, 468
glutamate dehydrogenase, 514
IMP dehydrogenase, 471
pyrophosphatase, 475
Gulonate dehydrogenase, mercurials, 842
Gulose, fructokinase, 376
Gymnodinium nelsoni, respiration (endo-
genous),
malonate, 169
Gynaecotyla adunca, respiration (endo-
genous),
malonate, 173, 183
H
Hadacidin, adenylosuccinate synthetase,
467
Heart,
acetate accumulation, malonate, 140
acetate oxidation, mercurials, 878, 88r
acetate utilization, propionate, 613
acetylcholine response,
malonate, 217
mercurials, 946-947
acetylpyridines, 494, 499-500
aminomalonate decarboxylation in, 239
arabinose uptake, glucose, 263
catecholamine levels in,
a-methyldopa, 317-328
a-methyl-m-tyrosine, 317-318
C-l/C-6 ratio, malonate, 130-131
citrate accumulation,
malonate, 104
mercurials, 927
citrate levels in vivo, sequential inhibi-
tion by malonate and fluoroacetate, 112
citrate oxidation, malonate, 79
conduction,
dehydroacetate, 625
o-iodosobenzoate, 724
mercurials, 942-945
contractility,
2-deoxyglucose, 402-403
o-iodosobenzoate, 723
malonate, 128, 213-215, 217
mercurials, 941-946
porphyrindin, 669
contracture,
mercurials, 941-944
porphyrindin, 669
dehydroacetate, positive inotropic ac-
tion of, 625
electrocardiogram,
malonate, 215
mercurials, 945
epinephrine response,
malonate, 217
mercurials, 947
fumarate oxidation, malonate, 81
glucose metabolism, malonate, 131,
216
glucose uptake, 3-methylglucose, 263
glycolysis (anaerobic), malonate, 128
a-ketoglutarate accumulation, malon-
ate, 111
a-ketoglutarate oxidation,
malonate, 80, 84
mercurials, 879
K+ fluxes, 2-deoxyglucose, 403
K+/Na+ ratio, mercurials, 946
K+ uptake in mitochondria, mercurials,
909
malate oxidation,
malonate, 82
1172
SUBJECT INDEX
mercurials, 879
malonate,
compared with ouabain, 216
decarboxylation of, 232-233
formation in, 226
levels in vivo, 102
metabolism of, 216, 228
pH effects, 191
malonic ethyl ester, 236
malonyl-CoA formation in, 233-234
membrane potentials,
dehydroacetate, 625
2-deoxyglucose, 403
malonate, 214
mercurials, 896, 945-946
mercurials in vivo, 930, 940-941, 943-
945, 959
methylmalonate in, 224
nicotinamide, 500
ouabain stimulation of, 2-deoxyglucose,
402
oxalacetate oxidation, malonate, 82
oxidative phosphorylation,
malonate, 121-122
mercurials, 873-874
permeability to Ca++, mercurials, 947
pyruvate oxidation,
acetylene-dicarboxylate, 241
malonate, 75-77, 216
mercurials, 878-879
propane-tricarboxylate, 241
rate of beating,
dehydroacetate, 625
o-iodosobenzoate, 723
malonate, 128, 213-215
mercurials, 941-944
porphyrindin, 669
thiamine analogs, 527
refractory period, malonate, 214
respiration, kojic acid, 350
respiration (endogenous), malonate,
176-179, 181
respiration (glucose),
l,5-anhydro-D-glucitol-6-P, 379
mercurials, 883, 948
ribose metabolism, malonate, 132
succinate accumulation, malonate, 91,
94-95, 128
succinate dehydrogenase,
alkylmalonates, 37
dicarboxylate ions, 35-39
malonate, 22-24, 31-32, 47-48
succinate levels in vivo, malonate, 100-
102
succinate oxidation, malonate, 55
thiamine-PP level, thiamine analogs,
525-527
tissue culture cells from, mercurials,
968
transamination in, deoxypyridoxol,
569-570
tricarboxylate cycle, mercurials, 877-
879
vagal effects on, mercurials, 946-947
HeLa cells,
2-deoxyglucose-6-P oxidation, 388
glucose uptake, oxamate, 434
glycolysis (aerobic), oxamate, 434
growth of, oxamate, 434
mitosis in, mercurials, 969
Na+ pump, oxamate, 434
resistance to 2-deoxyglucose, 388
RNA biosynthesis, 969
Helianthus annus, malonate occurrence
in, 225
Helianthus tuberosus (Jerusalem arti-
choke), respiration (endogenous),
malonate, 172
Helium, nitrogen fixation, 291
Helix pomatia hepatopancreas,
cycle intermediates oxidations, malon-
ate, 79-81
respiration, malonate, 114-115, 174
succinate oxidation, malonate, 54
Heme, biosynthesis of,
mercurials, 888
Hemicentrotus pulcherrimus eggs, devel-
opment of,
ferricyanide, 678
o-iodosobenzoate, 727
Hemoglobin,
biosynthesis of, a-amino-^-chlorobu-
tyrate, 351
ferricyanide, 670-671
mercurials, 649, 755-757, 760
Hemolysis, mercurials, 900-907
SUBJECT INDEX
1173
Hemophilus parainfluenza e, glutamate
oxidation,
malonate, 152
Hensenula ellipsoidospora, growth of,
malonate, 195
Heparin,
amylases, 464
cell division, 462
/3-fructofuranosidase, 465
fumarase, 465
glucoronidases, 465
hyaluronidase, 459-460
lipoprotein lipase, 463
lysozyme, 459
ribonuclease, 461
trypsin, 456
Hepatomas,
malonate levels in vivo, 102
succinate dehydrogenase, malonate, 30,
50
succinate levels in vivo, malonate, 102
Heptanoate, kynurenine: a-ketoglutarate
transaminase, 608
Herpes virus,
inactivation by mercurials, 978
proliferation of, mercurials, 981
Hevea brasiliensis, malonate,
formation in, 226
occurrence in, 225
Hexanoate, see Caproate
Hexokinase, see also Fructokinase, Glu-
cokinase, and other kinases
iV-acetyl-D-glucosamine, 390
analogs, 376-386
competition between substrates, 376-
377
6-deoxy-6-fluoroglucose, 404
2-deoxyglucose, 389-390
D-glucosamine, 391-383, 390
D-glucosone, 383-385
hexose-phosphates, 377-381
mercurials, 782, 788, 806-807, 824, 842-
843
denaturation, 788
protection by glucose, 782
protection by Zn++, 782
reversal ^\•ith cysteine, 824
titration of SH groups, 806-807
nucleotides, 383
quinacrine, 551
tripolyphosphate, 383
Hexylamine, leucine aminopeptidase, 368
4-Hexylresorcinol, tyrosinase, 304
Hill reaction, mercurials, 891
Hippurate, glycine iV-acylase, 355
Histaminase, see Diamine oxidase
Histamine,
metabolism of, aminoguanidine, 363
release of,
chymotrypsin inhibitors, 374
hydrocinnamate, 374
indole, 374
3-indolepropionate, 374
o-iodosobenzoate, 725
malonate, 212-213
mercurials, 949
skatole, 374
Histidase,
analogs, 353
D-histidine, 269
D-Histidinal, L-histidine formation, 269
Histidine,
diamine oxidase, 365
phosphoribosyl - ATP pyrophosphory-
lase, 351
D-Histidine,
L-amino acid oxidase, 340
histidase, 269
L-Histidine,
arginase, 337
dipeptidase, 367
homoserine kinase, 357
uptake by Botrytis, analogs, 267
Histidine decarboxylase,
analogs, 352-353
caflfeate, 314
permanganate, 660
Histidine hydrazide, histidine decarboxy-
lase, 353
D-Histidinol, L-histidine formation, 269
Homobiotin,
biotin oxidase, 589
structure of, 588
Homocysteine,
serine deaminase, 357
tyrosine decarboxylase, 307
1174
SUBJECT INDEX
Homogentisate, p - hydroxyphenylpyru-
vate oxidase, 306
Homogentisate oxidase,
o-iodosobenzoate, 709, 717
mercurials, relation to Fe++, 771-772,
787
Homogentisicase,
ferricyanide, 673, 675
o-iodosobenzoate, protection by sub-
strate, 717
mercurials, 782
protection by Fe++, 782
protection by homogentisate, 782
Homolog, definition of, 246
Homooxybiotin, yeast fermentation, 588-
589
D-Homoserine, L-threonine synthetase, 357
DL - Homoserine, thetin : homocysteine
transmethylase, 357
L-Homoserine, aspartokinase, 356
Homoserine deaminase,
o-iodosobenzoate, 709
mercurials, 843
Homoserine kinase, analogs, 357
Hordeum vulgar e (cape barley), see also
Barley roots
succinate dehydrogenase, malonate, 27
Hormosira hanlcsii,
K+ uptake, mercurials, 908
respiration (endogenous), mercurials,
881
Horseshoe crab, see Limulus
Hyaluronate,
glucoronidases, 465
lysozyme, 459
ribonuclease, 462
Hyaluronidase, macroions, 459-461
Hydrazine, diamine oxidase, 362
Hydrocinnamate, see also /3-Phenylpro-
pionate
acetoacetate formation from butyrate,
613
D-amino acid oxidase, 342-343
ammonia formation in kidney, 348
carboxypeptidase, 365-366
chymotrypsin, 369-370, 372
dopa decarboxylase, 311
histamine release, 374
Hydrocinnamide, chymotrypsin, 372
Hydrogen,
formation in Chlorella, mercurials, 891
nitrogen fixation, 292-293
physical properties, 295
Hydrogenase,
analogs, 293-294
ferricyanide, 676
iodine, 685
o-iodosobenzoate, 709
malonate, 61
quinacrine, 551
Hydrogen bonding,
reactivity of enzyme SH groups, 644-
646
SH groups, 640
Hydrogenlyase,
hydrogen peroxide, 693
permanganate, 660
Hydrogenomonas facilis, COg fixation,
mercurials, 892
Hydrogen peroxide, see also Peroxides
GSH oxidation by, 694
potentiation of azide toxicity 696
potentiation of toxicity by iodoacetate,
696
Hydroquinone,
D-amido acid oxidase, 344
catechol oxidase, 298
tyrosinase, 304
Hydroquinone monobenzyl ether, see
Monobenzone
Hydroxocobalamin, diol dehydrase, 590
Hydroxyacetate, see Glycolate
D-a-Hydroxy acid dehydrogenase,
analogs, 436-437
malonate, 61, 65
oxalate, 435-437
L-a-Hydroxy acid oxidase,
o-iodosobenzoate, 709
mercurials, 843
quinacrine, 551
L-3-Hydroxyacyl-CoA hydro-lyase, see
Enoyl-CoA hydratase
3 - Hydroxy - N - allylnormophinan, mor-
phine iV-demethylase, 597
2-Hydroxy-6-aminopurine, xanthine oxi-
dase, 282
SUBJECT INDEX
1175
p-Hydroxyamphetamine, phenylalanine
/3-hydroxylase, 599
17/S - Hydroxyandrosta - 1 ,4 - diene - 3 - one,
Zl*-3-ketosteroid reductase, 450
3-Hydroxyanthranilate oxidase,
anthranilate, 594
o-iodosobenzoate, 709
mercurials, 843
protection by Fe++, 782
protection by substrate, 782
type of inhibition, 772
Hydroxyaspartate, aspartate : a - ketoglu-
tarate transaminase, 355
p-Hydroxybenzaldehyde, succinate semi-
aldehyde dehydrogenase, 601
p-Hydroxybenzamide, sulfanilamide ace-
tylase, 601
rw-Hydrobenzoate, dehydroshikimate re-
ductase, 606
o-Hydroxybenzoate, see Salicylate
p-Hydroxybenzoate,
dehydroshikimate reductase, 593, 605
p-hydroxyphenylpyruvate oxidase, 306
tyrosine:a-ketoglutarate transaminase,
305-306
Hydroxybenzoates,
D-amino acid oxidase, 341
catechol oxidase, 297-299
glutamate dehydrogenase, 331
p-Hydroxybenzyloxyamine, dopamine /?-
hydroxylase, 320
2-Hydroxy-5-bromobenzamide, sulfanil-
amide acetylase, 601
/5-Hydroxybutyrate,
conversion to acetoacetate, 6-aminoni-
cotinamide in vivo, 505
oxidation of,
galactoflavin in vivo, 544
iodine, 689
mercurials, 883
thiamine analogs in vivo, 520-521
/3-Hydroxybutyrate dehydrogenase,
analogs, 594
malonate, 61
mercurials, 843
3-Hydroxycinnamate,
dopa decarboxylase, 314
glutamate decarboxylase, 314
histidine decarboxylase, 314
succinate dehydrogenase, 314
tyrosine decarboxylase, 314
a-Hydroxyethanesulfonate, glycolate oxi-
dase, 438
9-(2'-Hydroxyethyl)riboflavin analog,
flavokinase, 539
xanthine oxidase in vivo, 544
/9-Hydroxyglutamate,
glutaminase, 333
glutamine synthetase, 336
Hydroxy group, interaction energy of,
276
p-Hydroxyhippurate, glycine iV-acylase,
355
^-Hydroxyisobutyrate dehydrogenase,
mercurials,
protection by NAD, 782
protection by substrate, 782
a-Hydroxyisocaproamide, leucine amino-
peptidase, 368
a-Hydroxyisocaproate, glutamate decar-
boxylase, 329
a-Hydroxyisovalerate, glutamate decar-
boxylase, 329
y-Hydroxy-a-ketoglutarate,
aconitase, 615-616
formation from glyoxylate and oxala-
cetate, 615
isocitrate dehydrogenase, 615-616
a-ketoglutarate oxidation, 616
respiration (endogenous), 616
succinate oxidation, 616
tricarboxylate cycle, 615-616
}'-Hydroxy - a - ketoglutarate synthetase,
mercurials, 843
Hydroxylamine reductase, mercurials,
787
f5-Hydroxylysine, glutamine synthetase,
336
Hydroxymalonate, see Tartronate
Hydroxymethanesulfonate, glycolate oxi-
dase, 438
2-Hydroxymethylene-17a-methylandro-
stan-17/S-ol-3-one, ^-hydroxysteroid de-
hydrogenase, 449
a-Hydroxy-/3-oxalosuccinate, see Oxalo-
malate
1176
SUBJECT INDEX
Hydroxyphenylacetates,
glutamate decarboxylase, 329
tyrosine:a-ketoglutarate transaminase,
305-306
Hydrox yphenylalanines, tyrosine - a - keto-
glutarate transaminase, 305-306
p-Hydroxyphenyllactate, p-hydroxyphe-
nylpyruvate oxidase, 306
m-Hydroxyphenylpyruvate, p-hydroxy-
phenylpyruvate oxidation, 272, 306,
595
p-Hydroxyphenylpyruvate,
glutamate decarboxylase, 329
oxidation of, m- hydroxy phenylpyru-
vate, 272, 306, 595
peroxidase, 599
pyruvate decarboxylase, 431
Hydroxyphenylpyruvates, dopa decar-
boxylase, 312
p-Hydroxyphenylpyruvate oxidase,
analogs, 305-306
m-hydroxypenylpyruvate, 306, 595
Hydroxyproline,
A i-pyrroline-5-carboxylate dehydroge-
nase, 336, 355
A i-pyrroline-5-carboxylate reductase,
355
/3-Hydroxypropionate-phosphate,cnolase,
410
/5-Hydroxypropionitrile, structure of, 41
Hydroxypurines, keto-enol equilibria, 280
3-Hydroxypyridine, glucose dehydroge-
nase, 501
a-Hydroxy-2-pyridine methanesulfonate,
glycolate levels in tobacco, 439
glycolate oxidase, 438
photosynthesis, 439
structure of, 438
Hydroxypyruvate,
lactate dehydrogenase, 437
pyruvate decarboxylase, 432
Hydroxypyruvate reductase, mercurials,
protection by hydroxypyruvate, 782
protection by NADH, 782
a-Hydroxysteroid dehydrogenase, estra-
trienes, 449
)5-Hydroxysteroid dehydrogenase, ana-
logs, 447, 449, 450
a-Hydroxy-/S-sulfopropionate, fumarase,
243, 275, 277
5-Hydroxytryptamine, see Serotonin
5 - Hydrox y try ptophan ,
histidine decarboxylase, 353
tryptophan pyrrolase, 324-325
L-tryptophan:sRNA ligase (AMP), 326
7-Hydroxytryptophan, tryptophan pyr-
rolase, 324
5-Hydroxytryptophan decarboxylase,
o-iodosobenzoate, 709, 718
a-methyldopa, 309-310
Hymenolepsis diminuta,
malonate metabolism in, 228
respiration (endogenous), malonate, 173
succinate dehydrogenase, malonate, 28
Hypochlorite, amino acid oxidation by,
658
Hypophosphate,
oxidative phosphorylation, 448
succinate dehydrogenase, 243
Hypophosphite,
formate dehydrogenase, 593
formate hydrogenlyase, 593
Hypoxanthine,
D -amino acid oxidase, 545
urate uptake by erythrocytes, 267
Idose, fructokinase, 376
IDP, see Inosinediphosphate
Imidazole,
diamine oxidase, 365
histidase, 353
histidine decarboxylase, 352
A i-pyrroline-5-carboxylate dehydroge-
nase, 355
A i-pyrroline-5-carboxylate reductase,
355
Imidazoleacetate, histidase, 353
Imidazole-iV^-methyltransferase, mercu-
rials, 843
Imidazolonepropionate hydrolase, hydro-
gen peroxide, 693
Imidodipeptidase, see Prolidase
IMP, see Inosinemonophosphate
Indene, tryptophanase, 324
SUBJECT INDEX
1177
Indigo f era endecaphylla, 3-nitropropionate
the toxic principle in, 244
Indole,
acetylindoxyl oxidase, 591
chymotrypsin, 371, 374
histamine release, 374
histidine decarboxylase, 352
tryptophanase, 323-324
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
3-Indoleacrylate, tryptophan pyrrolase,
325
3-Indoleacetate,
D-amino acid oxidase, 342
carboxypeptidase, 366
chymotrypsin, 371
histamine release, 374
tryptophanase, 323
tryptophan pyrrolase, 325
uptake by Avena coleoptile, mercurials,
911
Indoleacetate oxidase, analogs, 595
3-Indolealdehyde, acetylindoxyl oxidase,
591
3-Indolebutyrate,
chymotrypsin, 370-371
tryptophan pyrrolase, 325
2-Indolecarboxylate,
D-amino acid oxidase, 344, 346
glutamate dehydrogenase, 331
3-Indolecarboxylate, glutamate dehydro-
genase, 331
/9-3-Indoleethylamine,
structure of, 324
tryptophanase, 323
3-Indolepropionamide, chymotrypsin, 371
3-Indolepropionate,
carboxypeptidase, 367
chymotrypsin, 369-371, 373
histamine release, 374
tryptophanase, 323
tryptophan pyrrolase, 325
Indolylpyruvate keto-enol tautomerase,
iodine, 686
Indoxyl-sulfate, acetylindoxyl oxidase,
591
Infantile acrodynia, see Acrodynia
Infections, bacterial,
malonate, 221-224
Inflammation,
o-iodosobenzoate, 724
mercurials, 950
Influenza virus,
infectivity of, mercurials, 978, 980-981
proliferation of,
2-deoxyglucose, 400
malonate, 126-127, 193
Inhibition index, 253-254
Inhibitor constants, definition of, 252
Inosine,
5 '-nucleotidase, 472
pyridoxal kinase, 475
Inosinediphosphate (IDP),
ATPase, 445
isocitrate dehydrogenase, 509
Inosine hydrolase, analogs, 471
Inosinemonophosphate (IMP)
adenylosuccinate lyase, 466
D-amino acid oxidase, 545
fructose- 1 ,6-diphosphatase, 470
GMP reductase, 471
level in ascites cells, 2-deoxyglucose,
395
5'-nucleotidase, 471
pyridoxal kinase, 475
thiamine kinase, 475
Inosinemonophosphate dehydrogenase,
analogs, 471, 481
Inosine phosphorylase, analogs, 471
Inosinetriphosphatase (ITPase),
ADP, 446
IDP, 446
mercurials, relation to SH groups, 804
quinacrine, 551
Inosinetriphosphate (ITP)
ATPase, 445
isocitrate dehydrogenase, 509
NADH oxidase, 511
pyridoxal kinase, 475, 477
Inositol dehydrogenase, o-iodosobenzoate,
709
Insulin, 2-deoxyglucose uptake, 387
Interference inhibition, 42
Intestine,
acetylcholine response, hydrogen pero-
xide, 696
1178
SUBJECT INDEX
amino acid transport,
competition between amino acids,
264-265
deoxypyridoxol, 574
biotin transport, analogs, 267
carbohydrate transport, competition
between sugars, 264
Ca++ transport,
malonate, 208
mercurials, 909, 913-914
CI" transport, mercurials, 910
contracture, o-iodosobenzoate, 724
2-deoxyglucose transport, 387
electrical potential across, mercurials,
916-917
galactose transport,
analogs, 263
6-deoxyglucose, 403
mannose, 263
glucose transport,
6-deoxyglucose, 264
deoxypyridoxol, 574
mercurials, 916
glycine transport, analogs, 265
I~ transport, mercurials, 910
mercurial levels in, 930, 959
methionine transport, deoxypyridoxol,
574-575
motility of,
dehydroacetate, 624-625
hydrogen peroxide, 696
o-iodosobenzoate, 724
malonate, 212
mercurials, 948-949
Na+ transport, mercurials, 916, 985
short-circuit current, 2-deoxyglucose,
387
triiodothyroacetate transport, mercu-
rials, 911, 913-914
uracil transport, mercurials, 911
water transport, mercurials, 916, 985
Invertase, see /S-Fructofuranosidase
lodate,
oxidation of nitrite, 450
oxidation of SH groups by, 656
Iodide,
active transports of, malonate, 209
intestinal transport of, mercurials, 910
^-ketoadipate chlorinase, 453
tyrosinase, 300
uptake by ciliary body,
malonate, 209
nitrite, 267
uptake by Fucus, mercurials, 910, 912
uptake by thyroid,
malonate, 209
mercurials, 910
Iodine,
active transports, 690
bacterial growth, 690
chemical properties, 679-680
enzyme inhibitions, 682-688
glycolysis, 689
mitochondrial swelling, 689
NADH oxidation by, 689
oxidation of protein SH groups, 680-
681
oxidative phosphorylation, 688-689
reaction with SH groups, 680
renal SH groups, 689
skin electrical potential, 690
solubility, 679
sulfenyl iodide formation, 681
lodoacetamide,
group attached to enzymes, 649
urease, 643
lodoacetate,
group attached to enzymes, 649
heart, antagonism of malonate effect,
216
kidney, transport of PAH, 205
toxicity, potentiation by hydrogen per-
oxide, 696
7H-Iodobenzoate, glutamate dehydroge-
nase, 330
o-Iodobenzoate,
bacterial growth, 727
formation from o-iodosobenzoate, 702
toxicity, 725
lodobenzoates, D-amino acid oxidase, 341
w;-Iodosobenzoate, 702-703
o-Iodosobenzoate,
ascorbate oxidation by, 703
bacterial growth, 727-728
catecholamine release, 725
central nervous system, 724
SUBJECT INDEX
1179
chemical properties, 701-703
enzyme inhibitions, 704-721
kinetics, 715-716
pH effects, 716-717
protection, 717-718
reversal, 718
variation with substrate, 718-721
glycolysis, 721
heart, 723-725
histamine release, 725
inflammatory activity, 724
intestine, 724
K^ in mitochondria, 722
lethal doses, 725
leucocytosis, 725
muscle, 723
NADH oxidation by, 703
nerve, 724
neuroblastic damage, 724
oxidation of SH groups, 657, 702-703
preparation of, 702
purification of, 702
reaction with protein SH groups, 703-
704
sea urchin egg development, 726-727
solubility, 716
titration of enzyme SH groups, 714-
715
toxicity, 724-726
urease, 643
uterus, 724
viruses, 728
p-Iodosobenzoate,
ascorbate oxidation by, 703
glutamate dehydrogenase, 702
3'-IodothjTonine, thjToxine deiodinase,
603
lodotyrosine, see L-MonoiodotjTosine
o-Iodoxybenzoate,
bacterial growth, 727
chemical properties, 702
Ion antagonisms, 452-453
Ionization constants,
dehydroacetate, 619
dicarboxylic acids, 8
hydrogen paroxide, 690
hypoiodous acid, 679
o-iodobenzoic acid, 702
o-iodosobenzoic acid, 702
mercuric complexes, 736
SH groups, 637-638
Ipomea batatas (sweet potato), oxidative
phosphorylation,
malonate, 120
Iproniazid, structure of, 488
Iron, see also Ferric ions
catalysis of SH oxidation, 658
complexes with di- and tricarboxylates,
12
incorporation into protoporphjTin, mer-
curials, 888
intestinal transport of, malonate, 208
Iron oxidase,
mercurials, 843
quinacrine, 551
Isatin, acetylindoxyl oxidase, 591
Isoamylase, mercurials, 843
Isoamylase (debranching), glucono-1,4-
lactone, 429
IsoamylbutjTate, lipase, 595
Isobologram, malonate and ouabain on
heart, 216
Isobutylamine, leucine decarboxylase, 352
Isobutyrate, glutamate decarboxylase,
328
Isocaproamide, leucine aminopeptidase,
368
Isocaproate,
glutamate decarboxylase. 328
leucine aminopeptidase, 368
Isocitrate,
oxalosuccinate decarboxylase, 597
oxidation of, malonate, 79, 86
phosphofructokinase, 385-386
Isocitrate dehydrogenase,
ferrocyanide, 677-678
y-hydroxy-a-ketoglutarate, 615-616
o-iodosobenzoate, 709
mercurials, 843-844
protection by isocitrate, 782
protection by Mn++, 782
nucleotides, 509, 513
propane-tricarboxylate, 240
Isocitrate lyase,
o-iodosobenzoate, 709
malonate, 61
1180
SUBJECT INDEX
Isohypophosphate, oxidative phosphory-
lation, 448
D-Isoleucine, L-alanine dehydrogenase,
354
L-Isoleucine,
arginase, 337
dipeptidase, 367
homoserine kinase, 357
Isomaltose,
a-dextran-l,6-glucosida8e, 417
a-glucosidase, 416, 423
Isomeric analogs, 268-274
anomeric, 271-272
definition of, 257
enantiomorphic, 268-271
geometric, 272-274
positional, 272
Isomorphic groups, 257
Isoniazid,
cadaverine oxidation, 363
glucose dehydrogenase, 501
NAD nucleosidase, 488, 490-491, 493
structure of, 488
Isoniazid-NAD, NAD nucleosidase, 489
Isonicotinamide, NAD nucleosidase, 488,
493
Isonicotinate,
glucose dehydrogenase, 501
NAD nucleosidase, 491
Isonicotinyl hydrazide, see Isoniazid
Isopenteny 1 - pyrophosphate isomerase ,
mercurials, 887
Isophthalate,
D-amino acid oxidase, 341, 344
aspartate : a - ketoglutarate transami-
nase, 334
glutamate dehydrogenase, 330-331
intercharge distance, 6
ionization constants, 8
kynurenine:a-ketoglutarate transami-
nase, 607-607
Isoporphobilinogen, porphobilinogen de-
aminase, 600
Isoriboflavin,
L-amino acid oxidase, 540-542
FAD pyrophosphorylase, 542
flavokinase, 539
Lactobacillus growth, 537
riboflavin deficiency, 538
structure of, 536
Isostere, definition of, 246
Isosteric groups, 257
Isovalerate,
D-amino acid oxidase, 343
glutamate decarboxylase, 328
leucine decarboxylase, 352
Isoxanthopterin ,
structure of, 287
xanthine oxidase, 289
Isoxanthopterin-6-carboxylate, xanthine
oxidase, 289
Itaconate,
biosjTithesis of, ferrocyanide, 678
fumarase, 279
glutamate decarboxylase, 328
ionization constants, 8
metabolism of, ferrocyanide, 678
structure of, 279
succinate dehydrogenase, 38, 601
ITP, see Inosinetriphosphate
ITPase, see Inosinetriphosphatase
J
Jensen sarcoma,
acetoacetate accumulation, malonate,
138
malonate levels in vivo, 102
respiration (endogenous), malonate, 179
succinate levels in vivo, malonate, 102
Jerusalem artichoke, see Helianthus tube-
rosus
K
/3-Ketoadipate,
aminolevulinate dehydrase, 591
kynurenine:a-ketoglutarate transami-
nase, 595
pyridoxamine : oxalacetate transami-
nase, 600
/3-Ketoadipate chlorinase, halide ions, 453
2 - Keto - 3 - deoxy - D - arabo - heptonate - 7 -
phosphate synthetase, analogs, 413
a-Ketoglutarate,
accumulation of, malonate, 110-111
glutamate dehydrogenase, 330
D-a-hydroxy acid dehydrogenase, 437
SUBJECT INDEX
1181
lactate dehydrogenase, 437
oxidation of,
benzoate, 348
malonate, 72, 79-81, 83-86
mercurials, 879
propane-tricarboxylate, 240
«?eso-tartrate, 432
thiamine analogs in vivo, 520-521
phosphofructokinase, 385
pyridoxamine : oxalacetate transami-
nase, 600
pyruvate decarboxylase, 431
a-Ketoglutarate oxidase,
6-aminocotinamide in vivo, 505
y-hydroxy-a-ketoglutarate, 616
malonate, 61-62, 83-86
mercurials, 844
inhibition in vivo, 926
oxygen inactivation of, 659
a-Ketoglutarate, reductase, mercurials,
844
^-Ketoglutarate, kynureninera-ketogluta-
rate transaminase, 608
2-Keto-L-gulonate, /3-glucuronidase, 424
a-Ketoisocaproamide, leucine aminopepti-
dase, 368
a-Ketoisocaproate, glutamate decarboxy-
lase, 329
a-Ketoisocaproate decarboxylase,
benzoate, 349
mercurials, 844
a-Ketoisovalerate, glutamate decarboxy-
lase, 329
Ketomalonate,
pyridoxamine : oxalacetate transami-
nase, 600
pyruvate decarboxylase, 430-431
zli-3-Ketosteroid reductase, estrone, 449
zl*-3-Ketosteroid reductase, analogs, 449-
450
Kidney,
acetate oxidation, malonate, 78, 232
acetate utilization, propionate, 613
acetoacetate metabolism, malonate,
144
ADP levels in, 2-deoxyglucose, 395
D-alanine oxidation, benzoate, 341
amino acid deamination, benzoate, 348
amino acid decarboxylase in vivo, a-
methyl-wi-tyrosine, 317
amino acid transport, competition be-
tween amino acids, 264
2J-amonohippurate transport,
azide, 205
cyanide, 205
dehydroacetate, 205
2,4-dinitrophenol, 205
fluoride, 205 '
fluoroacetate, 205
iodoacetate, 205
malonate, 205
mercurials, 205
phlorizin, 205
ammonia formation,
benzamide, 348
benzoate, 348
phenylacetate, 348
/3-phenylpropionate, 348
ATP levels in, 2-deoxyglucose, 395
Ca++ uptake in mitochondria, mercu-
rials, 909
C-l/C-6 ratio, malonate, 130
cholesterol biosjTithesis, malonate, 150
citrate accumulation in vivo, malonate,
104-105
citrate levels,
mercurials, 927
sequential inhibition by malonate
and fluoroacetate, 112
coenzyme A level in vivo, mercurials,
927
dehydroacetate, 626
fatty acid oxidation, malonate, 114,
137, 141-143
fluoromalonate decarboxylation, 239
galactose transport, glucose, 262
glucose oxidation,
6-deoxy-6-fluoroglucose, 393, 404
2-deoxyglucose, 393, 403
glucose-6-phosphatase in vivo, mercu-
rials, 927
glucose uptake, malonate, 127
glutamate metabolism, malonate, 152
jff-glycerophosphatase in vivo, mercu-
rials, 926-927
glycolysis, 2-deoxygIuco8e, 392-393
1182
SUBJECT INDEX
glycolysis (aerobic), ferricyanide, 677
GSH levels, tetrathionate, 700
histological changes,
malonate, 219-220
tetrathionate, 700
a-ketoglutarate oxidase in vivo, mer-
curials, 926
a-ketoglutarate oxidation, malonate,
81, 85
K+ transport, malonate, 205-206
malonate,
decarboxylation of, 232
levels in vivo, 100, 102
oxidation of, 233
toxicity of, 2, 219-220
mercurials, see also Mercurials, kidney
levels in vivo, 959-961
resistance to, 985
mitochondria of, cycle intermediates
concentrations in, 88-89
NAD(P) diaphorase in vivo, mercurials,
926
Na+ transport, malonate, 205-206
oxalacetate oxidation, malonate, 82
oxidative phosphorylation, mercurials,
873, 927
phosphatases in vivo, mercurials, 926-
927
protein disulfide reductase in vivo,
mercurials, 926
pyrithiamine levels in vivo, 528
pyruvate oxidation,
malonate, 75
meso-tartrate, 432
respiration,
<ra»5-aconitate, 273
kojic acid, 350
malonate, 219
respiration (acetate),
dehydroacetate, 626
malonate, 206
respiration (endogenous),
benzoate, 348
dehydroacetate, 623-624
malonate, 175-179, 181, 183
mercurials, 833
respiration (glucose), mercurials, 883-
884, 927-928
respiratory quotient, malonate, 185
SH groups in, iodine, 689
sorbitol dehydrogenase in vivo, mercu-
rials, 926
succinate dehydrogenase,
dicarboxylate ions, 35-38
in vivo, mercurials, 925-926
malonate, 29, 31-32
succinate levels in vivo, malonate, 100,
102,
succinate oxidation, malonate, 56
thiamine-PP levels, pyrithiamine, 525
thiols in, mercurials, 931-923
transaminases in vivo, deoxypyridoxol,
570
tubular transports, see specific sub-
stances
Kidney bean leaves, malonate occurence
in, 224, 226
Klebsiella pneumoniae, infection by,
malonate, 221
Kojic acid,
D-amino acid oxidase, 342, 349-350
bacterial growth, 349
central nervous system, 349
choline oxidase, 350
Leptospira growth, 349
lethal doses, 349-350
occurrence of, 349
phagocytosis, 350
L-proline oxidase, 350
structure of, 345, 617
succinate oxidase, 350
tyramine oxidation, 350
urate oxidation, 350
Kynureninase,
analogs, 595
nicotinylalanine, 610
Kynurenine, kynurenine formamidase,
595
Kynurenine formamidase,
analogs, 595
mercurials, 860
Kynurenine hydroxylase, nicotinylala-
nine, 610
Kynurenine : a - ketoglutarate transami-
nase, see Transaminases
SUBJECT INDEX
1183
Lactate,
blood levels of,
malonate, 219
thiamine analogs, 520
enolase, 409
formation of, see Glycolysis
metabolism of, analogs 432-438
oxidation of,
6-aminonicotinamide, 504
malonate, 78
tyrosinase, 300
D-Lactate, phosphatase (acid), 441
L-Lactate,
D-lactate dehydrogenase, 437
phosphatase (acid), 441-443
D-Lactate: cytochrome c oxidoreductase,
analogs, 437
malonate, 62
oxalate, 435
L-Lactate: cytochrome c oxidoreductase,
analogs, 437
D-Lactate dehydrogenase, analogs, 437
L-Lactate dehydrogenase,
acetate, 436
active center of, 434
6-aminonicotinamide in vivo, 505
„/ianalogs, 432-437
benzoate, 501
hydrogen peroxide, 693
iodine, 682-683, 686
o-iodosobenzoate, 710, 717-718
potentiation of inhibition by sub-
strate, 717-718
isoenzymes of, oxalate, 436
malonate, 62, 65
mercurials, 768, 772-773, 786-787, 802,
804, 808-812, 814, 825, 844-845
coenzyme displacement, 786-787
complex with mercuric ion, 768
protection by NADH, 782
rate constant for inhibition, 811
relation to SH groups, 802, 804, 808-
809
reversal with cysteine, 825
spontaneous reversal, 814
type of inhibition, 772-773
nicotinamide analogs, 500-502
nicotinylhydrazide-NAD, 497
oxalate, 435-436
oxamate, 433
porphyrexide, 668
quinacrine, 547, 551
pH effects, 557-558
tartronate, 237-238, 436
thionicotinamide-NAD, 497
D-Lactate oxidase,
quinacrine, 552
riboflavin, 542
L-Lactate oxidase,
FAD, 543
FMN, 543
o-iodosobenzoate, 710
malonate, 62
quinacrine, 552
riboflavin, 542-543
Lactate oxidative decarboxylase, quina-
crine, 552
Lactobacillus, hydrogen peroxide forma-
tion in, 695
Lactobacillus acidophilus, growth of,
dehydroacetate, 632
Lactobacillus arabinosus,
glutamate utilization, analogs, 327
growth of,
ion analogs, 452
5-phosphoribonate, 411
Lactobacillus brevis,
growth of, dehydroacetate, 632
pentose-P utilization, mercurials, 885-
886
Lactobacillus casei,
folate metabolism, analogs, 582
growth of,
3-acetylpyridine, 494
dehydroacetate, 632
mercurials, 972
quinacrine, 546
riboflavin analogs, 537-538
mevalonate incorporation, mercurials,
886
Lactobacillus fermenti, growth of,
dehydroacetate, 632
thiamine analogs, 530
Lactobacillus helveticus, growth of,
deoxypyridoxol, 575
1184
SUBJECT INDEX
co-methylpyridoxol, 575
Lactobacillus lactis, cyanocobalamin ana-
logs, 589-590
Lactobacillus plantarum, growth of,
dehydroacetate, 632
Lactonase-I, mercurials, 845
Lactose,
/3-galactosidase, 418
a-glucosidase, 416
Lecithinase A, analogs, 595
Leguminosae, malonate occurrence in
various species, 225-226
Leptospira icterohaem orrhagiae ,
growth of, kojic acid, 349
respiration (endogenous), malonate, 168
Lethal doses,
3-acetylpyridine, 499
adipate, 201
6-aminonicotinamide, 504
dehydroacetate, 627-628
6-deoxy-6-fluoroglucose, 404-405
2-deoxyglucose, 401
ethylmalonate, 201
fluoromalonate, 239
fluoromalonic diethyl ester, 239
D-glucosone, 384
glutarate, 201
hydrogen peroxide, 696
o-iodosobenzoate, 725
kojic acid, 349-350
malonamide, 201
malonate, 201, 218
mercurials, 955-957
oxythiamine, 530
pyrithiamine, 530
tetrathionate, 700
Lethal synthesis, analog incorporation,
247, 258
DL-Leucinamide, D-amino acid oxidase,
340
L-leucinamide, dipeptidase, 367
D-Leucine, L-amino acid oxidase, 268-
340
L-Leucine,
alanine: a-ketoglutarate transaminase,
334
D-amino acid oxidase, 340
arginase, 337
dipeptidase, 367-368
leucine aminopeptidase, 368
phosphatase (acid), 441
Leucine aminopeptidase, analogs, 368
Leucine decarboxylase,
analogs, 352
mercurials, 783, 810, 814, 845
protection by leucine, 783
protection by pyridoxal-P, 783
rate of inhibition, 810
spontaneous reversal, 814
L-Leucinol, leucine aminopeptidase, 368
Leucocytes,
glycolysis (aerobic), oxamate, 434
migration of, mercurials, 968
phagocytosis, malonate, 203
pyruvate oxidation, malonate, 76
Leucocytosis, o-iodosobenzoate, 725
Leuconostoc mesenteroides, growth of,
5-phosphoribonate, 411
Leucophaea maderae, succinate dehydro-
genase,
malonate, 29
Leucopterin,
structure of, 287
xanthine oxidase, 289
DL-Leucylglycine, D-amino acid oxidase,
340
DL-Leucylglycylglycine, D-amino acid oxi-
dase, 340
Leucyl-L-tyrosine, carboxypeptidase, 367
Leukemia,
deoxypyridoxol use, 576
malonate use, 202
Leukemic cells,
folate metabolism, analogs, 582
glycolysis, 2-deoxylucose, 392
Lilium ,
microspore meiosis, mercurials, 966-967
pollen of, malonate metabolism in, 228
Limulus polyphemus gill cartilage, succi-
nate dehydrogenase,
malonate, 19, 29
Lipase,
analogs, 595-596
dehydroacetate, 623
ferricyanide, 676
iodine, 686
SUBJECT INDEX
1185
o-iodosobenzoate, 710, 718-719
variation of inhibition with subs-
trates, 718-721
malonate, 62
mercurials, 719, 860
porphyrexide, 668
Lipids, see also Fatty acids. Phospholipids,
Sterols
biosynthesis of, mercurials, 886-887
metabolism of, see also Lipogenesis
malonate, 135-151
propionate, 613-614
Lipoamide dehydrogenase, see NADH:
lipoamide oxidoreductase
Lipoate,
analogs of, 590
acetyllipoate biosynthesis, 590
acyl transfer, 590
phosphotransacetylase, 590
pyruvate decarboxylation, 590
pyruvate oxidation, 590
mercurials, complexes with, 750-751
mercurial toxicity, antidote for, 751
pyruvate oxidase, antagonism of mer-
curial inhibition, 751
Lipogenesis, 2-deoxyglucose, 399
Lipoprotein lipase, macroions, 463
Lithium, phosphotransacetylase, 452
Liver,
acetate oxidation, malonate, 78
acetate utihzation, propionate, 613
acetoacetate accumulation, malonate,
138-144
acetoacetate metabolism, malonate,
115, 139-140, 144
ADP levels in, 2-deoxyglucose, 395
D-alanine oxidation, benzoate, 341
amino acid levels in, mercurials, 954
amino acid oxidation, kojic acid, 350
aminomalonate decarboxylation in, 239
arginine conversion from citrulline, ma-
lonate, 157-158
aspartate oxidation, malonate, 152
ATP levels,
aminopterin, 585
2-deoxyglucose, 395
malonate, 157-158
Ca++ uptake by mitochondria, mercu-
rials, 909-910
cholesterol metabolism, malonate, 150
citrate accumulation in vivo, malonate,
104
citrate oxidation, trans-aconitate, 273
Cu++ uptake, mercurials, 910, 913
cycle intermediates concentrations in,
89
damage by mercurials, 954
enzyme induction in, 8-azaguanine, 478
fatty acid biosynthesis,
malonate, 148-149
mercurials, 887
propionate, 613
fatty acid dehydrogenation (anaerobic),
malonate, 136-137
fatty acid oxidation,
malonate, 114, 137, 141-143
folate metabolism, analogs, 582
glucose metabolism, 6-deoxy-6-fluoro-
glucose, 404
glucuronide biosynthesis, analogs, 428
glutamate accumulation, malonate,
153-154
glutamate oxidation,
galactoflavin, 544
malonate, 152
mercurials, 878
glycogen formation, D-glucosamine, 382
glycogen levels in, oxythiamine, 520
glycolysis,
2-deoxyglucose, 383
malonate, 126
glycolysis (aerobic), malonate, 128
GSH levels in, tetrathionate, 700
/5-hydroxybutyrate oxidation,
galactoflavin, 544
mercurials, 883
isocitrate oxidation, malonate, 79, 86
a-ketoglutarate oxidation, malonate, 80
K+ uptake by mitochondria, mercu-
rials, 909, 914
lipogenesis, 2-deoxyglucose, 399
malate oxidation, malonate, 82
malonate,
decarboxylation in, 233
levels in vivo, 100, 102
1186
SUBJECT INDEX
metabolism in, 228, 232-234
mercurial levels, 930, 959-960
methylmalonate formation in, 226
methylmalonyl-CoA and succinyl-CoA
interconversion in, 235
Mg++ uptake by mitochondria, mer-
curials, 909
NAD levels in,
6-aminonicotinamide, 505
aminopterin, 585
Na+ transport by mitochondria, mer-
curials, 909
oxalacetate oxidation, malonate, 82
oxidative phosphorylation,
malonate, 119, 121
mercurials, 873
phenylalanine incorporation into pro-
teins, mercurials, 887
phospholipid biosynthesis, malonate,
151
propionate metabolism, malonate, 145
protein biosynthesis,
p-fluorophenylalanine, 351
malonate, 156,
mercurials, 887
pyridoxine levels in, deoxypyridoxol,
567
pyrithiamine levels in vivo, 528
pyruvate oxidation,
malonate, 75
mercurials, 878
pyrithiamine, 520
quinacrine, 560
respiration,
kojic acid, 350
quinacrine, 559-560
tartronate, 237
respiration (endogenous),
<rans-aconitate, 273
6-aminonicotinamide, 504
dehydroacetate, 623-634, 629
malonate, 176, 183
nicotinamide, 503
respiratory quotient, malonate, 184
squalene biosynthesis, mercurials, 886
succinate accumulation, malonate, 91,
145
succinate dehydrogenase,
malonate, 29-31
phthalate, 37
succinate levels in vim, malonate, 100,
102
succinate oxidase, malonate, 22
thiamine-PP levels in, thiamine ana-
logs, 525-527
transaminase in vivo, deoxypyridoxol,
570
urea formation,
deoxypyridoxol, 573
malonate, 116
Loblolly pine, see Pinus taeda
Locust, see also Schistocera
fat body of,
fatty acid formation from malonate,
234
malonate inhibition of fatty acid
biosynthesis, 147
malonate oxidation in, 233
sarcosomes, a-ketoglutarate accumula-
tion by malonate. 111
Locusta migratoria muscle, respiration
(endogenous),
malonate, 175
Lombricine kinase,
o-iodosobenzoate, 710
mercurials, 845
Lucerne (green alfalfa), malonate occur-
rence in, 225
Luciferase,
ferricyanide, 676
mercurials, 845
Lumichrome, flavokinase, 539
Lumiflavin,
flavokinase, 539
riboflavin transglucosidase, 543
structure of, 536
Luminescence, see Bioluminescence
Lung,
malonate decarboxylation in, 232
malonate levels in vivo, 102
mercurial levels in vivo, 930, 959
pentose-P pathway, malonate, 130
respiration (endogenous), malonate,
176-177
succinate levels in vivo, malonate, 102
SUBJECT INDEX
1187
transaminases in vivo, deoxyp>Tidoxol,
570
Lupine, see also Lupinus
Lupine mitochondria,
a - ketoglutarate oxidation, malonate,
80
oxidative phosphorylation, malonate,
119
succinate oxidation, malonate, 23
Lupinus albus,
oxidative phosphorylation, malonate,
119-120
succinate dehydrogenase, malonate, 33
Lymph node,
glucose uptake, analogs, 263
glycolysis, 2-deoxyglucose, 392
protein biosynthesis, malonate, 156
Lymphoma, protein biosjmthesis, ame-
thopterin, 585
Lymphosarcoma, growth of,
6-aminonicotinamide, 505
deoxypyridoxol, 576
riboflavin analogs, 538
Lysine,
arginase, 335, 337-338
histidase, 353
D-Lysine, D-amino acid oxidase, 340
L-Lysine,
aspartokinase, 356
blood urea, 338
homoserine kinase, 357
intestinal transport of, analogs, 265
leucine decarboxylase, 352
urea formation in vivo, 338
L-Lysine decarboxylase, mercurials, 845
Lysolecithin oxidase, FAD, 543
Lysozyme,
macroions, 459
mercuric ion complex, 770
Lytechinus eggs, cleavage,
malonate, 198
Lyxoflavin,
phosphorylation of, 539
structure of, 536
utilization by Lactobacillus, 539
Lyxoflavin dinucleotide, D-amino acid
oxidase, 544
Lyxose, /S-galactosidase, 418
M
Macroions,
amylases, 464
chymotrypsin, 457
;8-fructofuranosidase, 465
hyaluronidase, 459-461
inhibitions by,
factors determining, 455-456
kinetics of, 454-456
pH effects, 454-457, 461, 464
specificity of, 454
interaction with enzymes, 454-455
lipoprotein lipase, 463
lysozyme, 459
pepsin, 457-458
phosphatase (acid), 464
polynucleotide phosphorylase, 463
ribonuclease, 461-463
trypsin, 456-457
Macrophages, respiration (glucose),
oxamate, 435
Magnesium,
mitochondrial uptake, mercurials, 909
urinary excretion, mercurials, 921
Malate,
fumarase, 275-277
glutamate dehydrogenase, 332
ionization constants, 8
lactate dehydrogenase, 437
oxalacetate decarboxylase, 597
oxidation of,
malonate, 81-82
mercurials, 879
phosphofructokinase, 385
renal excretion of, malonate, 207
succinate dehydrogenase, 36
D- Malate,
phosphatase (acid), 441
D-tartrate dehydrase, 601
weso-tartrate dehydrase, 601
L-Malate,
D-a-hydroxy acid dehydrogenase, 437
phosphatase (acid), 441
D-tartrate dehydrase, 601
tartronate semialdehyde reductase, 602
Malate decarboxylase, malonate, 62
Malate dehydrogenase,
analogs, 596
1188
SUBJECT INDEX
)S-fluoromalate, 279
a-hydroxysulfonate, 438
iodine, 686
D-malate, 268
malonate, 62
mercurials, 845-846
relation to SH groups, 802-804, 807
808
type of inhibition, 772
nicotinamide, 503
nucleotides, 509, 513
porphyrexide, 668
quinacrine, 552
tartronate, 237
Malate dehydrogenase (decarboxylating)
(malic enzyme),
analogs, 596-597
o-iodosobenzoate, 704-710
malonate, 63, 65
mercurials, 846
protection by Mn++, 779, 783
protection by substrates, 779, 783
tartronate, 238
L-Malate hydro-lyase, see Fumarase
Malate synthetase, see Glyoxylate trans-
ace tase
Malate:vitamin K oxidoreductase,
mercurials, 846
quinacrine, 552
Maleate,
alanine; a-ketoglutarate transaminase,
334
D-amino acid oxidase, 343
aspartate : a - ketoglutarate transami-
nase, 334
chelation with cations, 12
fumarase, 275-277
glutamate decarboxylase, 328
intercharge distance, 6
ionic length, 188
ionic volume, 188
ionization constants, 8
permeability of erythrocytes to, 188
phosphatase (acid), 442
succinate dehydrogenase, 34-36
tyrosinase, 301
urinary citrate, 109
urinary a-ketoglutarate, 110
Maleate dehydrogenase, malonate, 63
Malignant carcinoid syndrome, deoxypy-
ridoxol, 574
Malonamide,
lethal dose, 201
tumoristasis, 201
Malonate,
acetate oxidation, 77-78, 232
acetoacetate metabolism, 138-144, 149,
234
acetylaspartate formation, 154
acetylcholine synthesis, 165-166
cis-aconitate oxidation, 79
adaptive enzyme synthesis, 155
amino acid accumulation by, 103
amino acid metabolism, 151-154
y-aminobutyrate oxidation, 154
aspartate : a - ketoglutarate transami-
nase, 334
ATP level in liver homogenate, 157-158
bacterial growth, 195
bacterial infections, 221-224
biosynthesis of, 2, 226-227
blood coagulation 219
blood glucose, 164, 219
blood lactate, 219
blood pressure, 213
blood pyruvate, 219
bond characteristics, 4
Br~ uptake by barley roots, 116-117
carcinostasis, 200-202
cell division, 117, 197-200
cellulose biosynthesis, 132
chelation of cations, 11-14, 66-69
cardiac effects of, 214-216
chemotaxis of leucocytes, 203
cholesterol metabolism, 149-150
chondroitin sulfate synthesis, 166
ciliary activity, 203
citrate accumulation by, 104-110
citrate levels in tissues, 223
citrate oxidation, 78-79, 86-87
coliphage, 194
coronary flow, 213
decarboxylation (nonenzymic) of, 3
decarboxylation in tissues, 232-234
determination of, 13
diuretic action, 221
SUBJECT INDEX
1189
electrocardiogram, 215
embryogenesis, 197-199
fatty acid oxidation, 114, 135-137
fatty acid biosynthesis, 146-149
flagellar activity, 203
fumarase, 275
fumarate antagonism of inhibitions by,
112-117
fumarate oxidation, 81
fungal growth, 195-196
gastric acid secretion, 187, 208
glucose metabolism, 122-135
glutamate decarboxylase, 328
glycerol metabolism, 164
glycolysis, 125-130
glyoxylate metabolism, 165
heart, 213-217
histamine release, 212-213
hydrogen bonding in, 4, 9
hydroxamic acid formation, 233
D-a-hydroxy acid dehydrogenase, 437
incorporation into lipids, 231
intercharge distance, 5-6
intestinal transport of, 207-208
ionic length and volume, 188
ionic species, pH effects, 9-10
ionization of, 5, 8-10
iron incorporation into heme, 163
isocitrate accumulation, 106
isocitrate oxidation, 79, 86
ketogenic activity, 138-144
a-ketoglute^rate oxidation, 72, 79-81,
83-86
kynurenine: a-ketoglutarate transami-
nase, 608
lactate formation, 126-128
lactate oxidation, 78
lethal doses, 201, 218
lipid metabolism, 135-151
malate oxidation, 81-82
metabolism of, 1-2, 105, 107, 138, 216,
224-235
methylmalonate accumulation by, 145
mitochondrial swelling, 210-211
mitosis, 197-200
muscle, 212
nephrotoxicity, 219-221
nerve conduction and potentials, 211-
212
occurrence of, 2, 224-226
oxalacetate oxidation, 66-68, 82
oxidative phosphorylation, 118-122
partition ratios, 189
pentose-P pathway, 130-132
permeability to, 51, 56, 167, 186-192
phagocytosis, 203, 223
pH effects on inhibitions by, 181-182,
189-192, 195-197
phosphatase (acid), 442
phospholipid metabolism, 151
photosynthesis, 163-164
plant growth, 196-197
plasma K+ and Na+, 206
plasma pH, 206
porphyrin biosynthesis, 158-163
propionate metabolism, 144-146
protein biosynthesis, 155-157
purification of, 3
pyruvate oxidation, 73-77
reduction of Ca++ and Mg++ concen-
trations, 13-14
renal excretion of malate, 207
renal transports, 203-207
respiration (endogenous), 166-186
effect of functional activity, 183-184
effect of ions and buffer, 183
effect of tissue age, 182-183
factors determining inhibition, 166-
167, 180
kinetics, 180-182
significance of inhibition, 186
respiration (glucose), 123-125, 127, 133-
135
respiratory quotient, 184-185
sequential inhibition with fluoroacetate,
112
specificity of inhibition, 58-59, 72, 117-
118
stability of, 3
structure of, 4
succinate accumulation by, 90-104
succinate decarboxylation to propio-
nate, 165
succinate dehydrogenase, 15-50
activation of, 45-46
1190
SUBJECT INDEX
binding energy, 42
competitive nature of inhibition, 21-
25
effect of ATP, 48
effect of Ca++, 46-48
effect of electron acceptor, 18-20
effect of Mg++, 48
effect of osmolarity, 46-47
effect of temperature, 46
evidence for site of inhibition, 20-21
inhibition constants, 25, 33-34
nature of binding, 40-45
variation between species and tissues,
49-50
succinate oxidation in cellular prepa-
rations, 50-58
tissue levels in vivo, 100-102
toxicity, 217-221
tricarboxylate cycle, 69-88
effect of alternate pathways, 71-72
effect of oxalacetate supply, 69-71
effect of succinate accumulation, 70-
71
variation with time, 70
uptake by leaves, 107
urea formation, 157-158
urinary acetone, 138
urinary citrate, 99, 104-110
urinary excretion of, 101
urinary flow, 206
urinary a-ketoglutarate, 99, 110-111
urinary pH, 206
urinary succinate, 98-99, 101
virus proliferation, 192-194
yeast growth, 195
Malonate decarboxylase, 229
Malonate semialdehyde, occurrence of,
225
Malondialdehyde,
structure of, 41
succinate dehydrogenase, 40-41
Malondiamide,
structure of, 41
succinate dehydrogenase, 41
Malonic diethyl ester,
acetate oxidation, 236-237
glucose oxidation, 236-237
mycobacterial infections, 224
succinate dehydrogenase, 236
Malonic esters, ionization of,
constants, 1 1
rates, 11
Malonic ethyl esters.
Bacillus cereus sporulation, 236
carcinostasis, 236
effects on metabolism, 236-237
hydrolysis of, 236
ionization constants, 236
keto-enol tautomerism, 4
mycobacterial infections, 236
permeability to, 11
prolongation of hexobarbital action,
236
Malonyl-CoA-COj exchange enzyme, mer-
curials, 847
Malonyl semialdehyde pantetheine, reac-
tion with jj-mercuribenzoate, 751
Maltase, see also Amylomaltase and Iso-
amylase
o-iodosobenzoate, 710
mercurials, 860
methylglucoside anomers, 271
synthesis of,
p-fluoro phenylalanine, 351
tryptazan, 326
Maltose,
a-amylase, 420
/3-amylase, 421
a-dextran-l,6-glucosdase, 417
)S-galactosidase, 418
^-glucosidase, 417, 423
a-mannosidase, 422
Maltose 4-glucosyltransferase, see Amy-
lomaltase
Malus malus,
citrate oxidation, malonate, 79
succinate oxidation, malonate, 53
Malus sylvestris, see also Apple
oxidative phosphorylation, malonate,
120
Mammary carcinoma, growth of,
deoxypyridoxol, 576-577
malonate and derivatives, 202
Mammary gland,
citrate utilization, ^rans-aconitate, 273
fatty acid biosynthesis,
SUBJECT INDEX
1191
<ran5-aconitate, 273
mercurials, 887
fatty acid formation from acetate or
propionate, malonate, 146-147
glutamate oxidation, malonate, 152
pyruvate oxidation, malonate, 75
respiration (acetate), malonate, 146
respiration (endogenous), malonate, 176
Mandelate, lactate dehydrogenase, 437
Mannarate, /3-glucuronidase, 424
Mannaro -1,4-3,6- dilactone, fi - glucoroni-
dase, 424
Mannitol, a-mannosidase, 422
Mannoheptulose,
fructokinase, 376
glucokinase, 376
glucose uptake into tissues, 376
Manono-l,4-lactone, a-mannosidase, 429
Mannose,
a-amylase, 420
fructokinase, 376
galactose transport, 263
glucokinase, 376
glucose uptake by lymph node, 263
a-glucosidase, 416-417
a-mannosidase, 422
toxicity to bees, 414
uptake by lymph node, 394
Mannose-6-phosphate,
glucose-6-P dehydrogenase, 411
hexokinase, 379-380
phosphopentose isomerase, 411
a-Mannosidase,
analogs, 422, 429
mercurials, 812
Mannuronate,
a-glucuronidase, 426
y3-glucoronidase, 424
Mannurone, jS-glucuronidase, 424
Maple sugar urine disease, see Branched
chain ketonuria
Marigold stem cultures, growth of,
malonate, 197
Marinogamm arus m arinus ,
respiration (endogenous), mercurials,
882
toxicity of mercurials, 861
Melanin, formation of.
analogs, 303-305
Melezitose, a-glucosidase, 416
Melibiose, galactosidases, 418
Membrane potentials, see also specific tis-
sues
heart,
acetylpyridines, 499-500
dehydroacetate, 625
malonate, 214-215
mercurials, 896, 945-946
intestine, mercurials, 916-917
kidney, mercurials, 923, 936
muscle, mercurials, 937-938
nerve,
malonate, 211-212
mercurials, 949-950
oxythiamine, 531
pyrithiamine, 531
skin,
iodine, 690
mercurials, 950
porphyrindin, 669-670
Membrane transport, see also Active trans-
port. Permeability, and the specific or-
ganisms and tissues
analog inhibition of, 261-268
Menadione reductase, see NADH:mena-
dione oxidoreductase
Menthyl-a-glucuronide,
a-glucuronidase, 426
^-glucoronidase, 272
Mepacrine, see Quinacrine
Meralluride, see also Mercurials, kidney
structure of, 917
Merbromin (Mercurochrome), structure
of, 970
Mercaptalbumin, mercurials, 757-761
Mercaptans, see Thiols
Mercaptides,
formation of, 642
of mercurials and thiols, 746-751
Mercaptoacetate, lactate dehydrogenase,
437
a-Mercaptobutyrate, lactate dehydroge-
nase, 437
Mercapto groups, see Sulf hydryl groups
Mercaptomerin, see Mercurin and Mer-
curials, kidney
1192
SUBJECT INDEX
a-Mercaptopropionate, lactate dehydro-
genase, 437
6-Mercaptopurine,
adenosine deaminase, 466
inosinic acid metabolism, 281
metabolic products of, 481
purine metabolism, 480
xanthine oxidase, 282
6-Mercaptopurine nucleotide,
adenylosuccinate lyase, 466, 481
adenylosuccinate synthetase, 467
polynucleotide phosphorylase, 481
Mercaptosuccinate, lactate' dehydroge-
nase, 437
Mercuhydrin, see Meralluride
Mercurialism, see Mercurials, toxicity
Mercurials,
accumulation in kidneys in vivo, 923-924
acetate uptake by diaphragm, 912
acrodynia, 953-954
active transports, 907-917, 936-937
actomyosin, 938-940
affinities for ligands, 732-741, 744
ameboid movement, 982
amino acid complexes with, pH effects,
760-761
amino acid levels in liver, 954
auxin transport, 967
bacterial growth, 970-976
bacterial uptake of, 974-975
bioluminescence, 888-891
catecholamine release by, 947
Ca++ uptake by mitochondria, 909-910
cell membranes, 892-907
central nervous system, 951-952
chemical properties, 730-745
citrate accumulation in vivo, 927
coenzyme A complexes with, 750
COa fixation, 892
colored, histochemical SH determina-
tions, 766-768
comparison of inorganic and organic,
743-745
conditioned reflexes, 985
crystalhne mercuri-enzymes, 768-770
Cu++ uptake by liver, 910, 913
determination of protein SH groups,
762-768
dissociation into Hg++, 930-935
diuretics, see also Mercurials, kidney
release of Hg++, 930-935
structures of, 917
type structure of, 930
electrocardiogram, 945
electron transport, 870-872
embryogenesis, 963-965
enzyme inhibitions, 768-869
coenzyme displacement, 784-787
comparison of mercurials, 860-861
configurational changes, 787-790
effect of buffers, 797-798
effect of ionic strength, 797
effect of pH, 790-797
kinetics, 771-778, 908-815
meaning of K„ 861-862
methods of expressing inhibitions,
861-862
mutual depletion situations, 861-
862
protection, 778-784
rates of, 809-815
reversals, 821-828
type of, 771-778
ethanol oxidation, 898
fatty acid biosynthesis, 887
fatty acid oxidation, 887
functionality, 743
ganglionic transmission, 949
gastric acid secretion, 914-916
glucose uptake, 893-894, 903-905, 910-
911
glycolysis, 874-877
growth stimulation, 967-968, 971
GSH level in erythrocytes, 905-906
heart, 940-948
hemoglobin, 755-757, 760
hemolysis, 900-907
histamine release by, 949
histological changes, 954-955
intestine, 909-917, 948-949
membrane potentials, 916-917
motility, 948-949
transports, 909-911, 913-914, 916-
917
invertebrates, 961-963
K+ efflux,
SUBJECT INDEX
1193
erythrocytes, 903-905
yeast, 898-900
kidney, 917-937
coenzyme A levels, 927
damage to, 924-925
enzyme activities in vivo, 925-927
glomerular filtration, 918
mechanisms involved, 930-937
membrane potentials, 923, 936
relation of structure to action, 930-
935
resistance to toxic effects, 985
scheme of reactions in, 934
SH groups in, 921-923
sites of action, 920-925
summary of actions, 918-920
tubular transports, 205, 918-921, 928,
936
K+ uptake, 908-909, 914
lethal doses, 955-957
lipid biosynthesis, 886-887
lipid solubility of, 743-744
lipoate complexes of, 750-751
mercaptalbumin, 757-761
Mg++ uptake by mitochondria, 909
Mg++ urinary excretion, 921
mitosis, 963-970
muscle, 937-940
Na+ transport, 909
nephrotoxicity, 924-925
nerves, 949-950
neuroblastic damage, 964-965
nuclear damage, 965
organic, chemical properties of, 742-
745
ovabulmin, 753-755, 760
oxidative phosphorylation, 872-874
in vivo, 927
parthogenesis by, 963-964
pentose-P pathway, 885-886
permeability of erythrocytes, 900-907
phosphate uptake, 910, 912-913
phospholipid biosynthesis, 887
photophosphorylation, 892
photosynthesis, 891-892
plant growth, 965-968
porphyrin biosynthesis, 888
proteins,
biosynthesis of, 887-888
configurational changes, 761-762
pH effects, 760-761
reactions with, 751-768
protozoa, 981-982
reaction with disulfides, 750
reaction with NAD, 774
reaction with SH groups, 764-751
pH effects, 749-750
reaction with thiamine, 774
resistance to, 983-985
respiration, 879-886
stimulation of, 879, 881-882, 884
respiration (glucose), 893-894, 898
retina, 952-953
skin, 950
spindle formation, 969
sterol biosynthesis, 886-887
sucrose uptake, 911
thioester splitting by, 751
tissue distributions of, 930, 955, 958-
961
titration of enzyme SH groups, 798-
809
titration of protein SH groups, 762-766
toxicity, 940-941, 944, 950-957
transfer ENA biosynthesis, 820
tricarboxylate cycle, 877-879
tumor uptake of in vivo, 969-970
uracil transport, 911
urinary Ca++ excretion, 921
urinary excretion of, 928-930, 952, 955,
958, 960
viruses, 976-981
water transport by cornea, 911
xylose uptake by diaphragm, 911-912
p-Mercuribenzoate (p-MB), see also Mer-
curials
aldolase, 643, 649
group attached to enzymes, 649
hemoglobin SH groups, 649
lipase, variation of inhibition with sub-
strate, 719
nucleic acid complexes of, 744
3 - phosphoglyceraldehyde dehydroge-
nase, 650
phosphorylase a, 648-649
preparation of solutions of, 743
1194
SUBJECT INDEX
purification, 745
reaction with NAD, 774
release of Hg++, 744-745
solubility, 743
stability, 745
structure of, 742
synthesis of, 745
titration of SH groups, 763-766
urease, 643
Mercuric chloride, see also Mercurials and
Mercuric ion
solubility, 730-731
solubility product, 731
structure of, 730-731
Mercuric ion, see also Mercurials for gen-
eral topics
amine complexes of, 738-740
amino acid complexes of, 737-741, 760-
761
bacterial growth, antagonism by thia-
mine, 774
bond characteristics, 731, 744-745
concentrations in solution, 732-736,
740-741
cysteine complexes of, 739, 748-749
EDTA complexes of, 738
electronegativity, 731
GSH complex of, 748-749
halide complexes of, 730-738
hydration of, 736
hydroxyl ion complexes of, 736
methionine complexes of, 739
nucleic acid complexes of, 741
nucleoside complexes of, 739
nucleotide complexes of, 741
oxidation-reduction potentials, 731
phosphoribosyl - ATP pyrophosphoryl -
ase, on inhibition by histidine, 351
purine complexes of, 739
pyrimidine complexes of, 739
pyrophosphate complex of, 738
thioglycolate complex of, 748-749
uptake by Achromohacter , 897
uptake by diagragm, 894-897
uptake by erythrocytes, 897, 900-907
uptake by yeast, 898-900
4-Mercuri-4'-dimethylaminoazabenzene,
determination of tissue thiols, 767-768
Mercurimalonamide, possible inhibitor of
succinate dehydrogenase, 240
Mercurin (mercaptomerin, Thiomerin), see
also Mercurials, kidney,
structure of, 917
Mercuripapain, 769-770
p-Mercuriphenylazo-/3-naphthol, determi-
nation of tissue thiols, 766-767
4-(p-Mercuriphenylazo)-l-naphthylamine-
7 - sulfonate, determination of tissue
thiols, 768
p-Mercuriphenylsulfonate, see also Mer-
curials and 7;-Mercuribenzoate,
structure of, 742
Mercurochrome, see Merbromin
Mercurophylline, see Mercurials, kidney
Merphenyl, see Phenylmercuric ion
Mersalyl, see also Mercurials, kidney,
cyclic mercaptides, 746-747
renal transport of PAH, 205
structure of, 917
Merthiolate, see Thimerosal
Mesaconate (methylfumarate),
fumarase, 275-277, 279
glutamate decarboxylase, 328
ionization constants, 8
structure of, 279
Meso porphyrin, tryptophan pyrrolase,
603
Mesoxalate, malic enzyme, 597
Metanephrine, urinary excretion of,
pyrogallol, 612
Metaphen, see ISiitromersol
Metaphosphate, phosphatases, 439
Methanediphosphonate, succinate dehy-
drogenase, 243
Methanedisulfonate (methionate),
intercharge distance, 7
succinate dehydrogenase, 243
Methanesulfonate, sulfite oxidase, 451
Methicillin, penicillinase, 598
Methionate, see Methanedisulfonate
L-Methionine,
intestinal transport of D-methionine,
265
synthesis from serine, cyanocobalamin
analogs, 590
Methionine sulfoximine,
SUBJECT INDEX
1195
glutamine synthetase, 335
methionine incorporation into proteins,
335
Methotrexate, see Amethopterin
Methoxyacetate, sarcosine oxidase, 601
Methoxybenzoates,
D-amino acid oxidase, 341, 344
catechol oxidase, 298-299
p-Methoxybenzoyl-D-tryptophanamide,
chymotrypsin, 371
3-Methoxydopamine,
dopamine ^-hydroxylase, 320
phenylalanine /^-hydroxylase, 600
4-Methoxypyridoxal,
central nervous system, 573
embryogenesis, 576
GABA in brain, 573-574
Methylacetate, lipase, 596
/3-Methylacrylate, see Crotonate
2-Methyladenine,
adenine deaminase, 466
inosine phosphorylase, 471
iV-Methyladenine,
adenine deaminase, 466
inosine phosphorylase, 471
a-Methylaspartate, aspartate:a-ketoglu-
tarate transaminase, 334
p-Methylbenzoate, catechol oxidase, 298-
299
Methylbenzoates, D-amino acid oxidase,
341
jS-Methylcysteine, glyoxylase, 594
a-Methyldopa,
blood pressure, 315
bronchoconstriction from S-hydroxy-
tryptophan, 315
cardiac stimulation by dopa, 314-315
catecholamine levels in tissues, 315-320
dopa decarboxylase, 308-320
in vivo inhibition, 314-316
kinetics, 309-310
mechanism of inhibition, 309-310
formation of abnormal amine from, 318
5-hydroxytryptophan decarboxylase,
309-310
in vivo inhibition, 314
miosis, 315
pressor response to dopa, 314
reaction with pyridoxal-P, 309
tissue levels of, 318-320
tyrosine decarboxylase, 309
urinary excretion of amines, 315
A^-Methyldopa, dopa decarboxylase, 308
a-Methyldopamine, phenylalanine /3-hy-
droxylase, 600
7-Methyldulcitylflavin, flavokinase, 539
Methylenesuccinate, see Itaconate
3-Methyl-3-ethylglutarate, kynurenine:a-
ketoglutarate transaminase, 608
7-Methylfolate,
blood pressure, 586
dopa decarboxylase, 586
structure of, 580
tyrosine decarboxylase, 586
/?-Methylfructofuranoside, sucrose trans-
fructosylase, 421
Methylfumarate, see Mesaconate
Methylgalactosides, a-galactosidase, 418
3-Methylglucarate, /S-glucuronidase, 424
3-Methylglucaro-l,4-lactone, /3-glucuroni-
dase, 424
a-Methylglucopyranoside, sucrose trans-
fructosylase, 421
3-0-Methylglucose,
galactose transport, 263
glucose uptake, 263
a - Me thy Iglu coside ,
a-amylase, 420
^-fructofuranosidase, 271
galactose transport, 263
a-glucosidase, 416
maltase, 271
a-mannosidase, 422
phosphorylase, 271-272, 405
transport into E. coli, 2-deoxyglucose,
394
/3-Methylglucoside, a-glucosidase, 416
Methylglucosides, maltose transglucosy-
lase, 415
Methylglucuronates, /^-glucuronidase, 424
Methylglucoronides, /^-glucuronidase, 427
a -Methylglutamate ,
aspartate : a - ketoglutarate transami-
nase, 334
glutamate decarboxylase, 327
glutamine synthetase, 336
1196
SUBJECT INDEX
y-glutamyltransferase, 336
/9-Methylglutamate,
glutaminase, 333
urinary flow, 333
a-Methylglutarate,
kynurenine:a-ketoglutarate transami-
nase, 595, 608
/5-Methylglutarate,
glutamate dehydrogenase, 331
kynurenine:a-ketoglutarate transami-
nase, 608
(S-Methylglutathione, glyoxylase, 594
Methylguanidine,
diamine oxidase, 362
histidase, 353
2-Methylhistine, phosphoribosyl-ATP py-
rophosphorylase, 351
4 - Methyl - 5 - (/5 - hydroxyethyl) thiazole-
NAD, inactivation of dehydrogenases,
514
4 - Methyl - 5 - (/?-hydroxyethyl)thiazole-di-
phosphate, pyruvate decarboxylase, 516
2-Methyl-3-hydroxy-5-hydroxymethylpy-
rimidine, pyridoxal kinase, 565
a-Methyl-3-hydroxyphenylalanine, see a-
Methyl-m-tyrosine
a-Methyl-5-hydroxytryptophan, histidine
decarboxylase, 353
Methylindoles,
tryptophanase, 321
tryptophan synthetase, 321
fi- 1 -Methyl-3-indolylalanine,
structure of, 324
tryptophanase, 323-324
6-Methylisoxanthopterin, xanthine oxi-
dase, 289
6-Methyl-ll-ketoprogesterones, Zl*-3-ke-
tosteroid reductase, 450
Methylmaleate, see Citraconate
Methy Imalonate ,
accumulation of, malonate, 145
biosynthesis of, 226
metabolism of, 234-235
occurrence of, 224-225
renal action, 2
Methylmalonyl-CoA isomerase, intercon-
version of methylmalonate and succi-
nate, 235
yS-Methylmaltoside,
a-glucosidase, 416
maltose transglucosylase, 422
7-Methylmannitylflavin, flavokinase, 539
a-Mannoside, a-mannosidase, 422
^-Methylmannoside, a-glucosidase, 416
Methylmercuric ion (MM), see also Mer-
curials
anion complexes of, 744-745
cysteine complex of, 747
protein complexes of, 748
iV-Methylnicotinamide,
NAD nucleosidase, 493
renal transport of, dehydroacetate, 625
urinary levels of, nicotinylalanine, 610
2-Methylnicotinate, glucose dehydroge-
nase, 501
a-Methylnorepinephrine, formation from
a-methyldopa, 318
5-Methylorotate, dihydroorotate dehydro-
genase, 470
7-Methylpteroate, dopa decarboxylase,
586
iV-Methylpyridoxal, pyridoxal kinase, 564
co-Methylpyridoxal, pyridoxal kinase, 565
w-Methylpyridoxol,
glutamate decarboxylase in brain, 569,
571
growth of microorganisms, 575
Methylquinolines, chymotrypsin, 373
5-Methylresorcinol, see Orcinol
4-Methylthiazoles, thiaminase, 524
D-Methyl-DL-thyroxine, L-thyroxine deio-
dinase, 602
0-Methyltransferase, see Catechol-0-me-
thyl transferase
6 -Methyl tryptazan ,
maltase biosynthesis, 326
L-tryptophan:sRNA ligase (AMP), 326
a-Methyltryptophan, induction of tryp-
tophan pyrrolase, 325
)3-Methyltryptophan,
tryptophan pyrrolase,
L-tryptophan:sRNA ligase (AMP), 326
4 - Methy Itryptophan,
E. coli growth, 323
tryptophan synthetase, 312
SUBJECT INDEX
1197
5-Methyltryptophan,
anthranilate synthesis, 321
maltase biosynthesis, 326
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
6-Methyltryptophan,
maltase biosynthesis, 326
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
a-Methyl-m-t>Tosine,
aminofcacid decarboxylase in vivo, 315-
317
catecholamine levels in brain, 315-316
dopa decarboxylase, 308-309
dopamine /3-hydroxylase, 320
0-Methyl-L-tyrosine, tyrosinase, 305
Methylurates, uricase, 284
Methylurea, urease, 603, 610
7-Methylxanthopterin, xanthine oxidase,
289
Mevalonate, incorporation into sterols,
mercurials, 886-887
Mevalonate dehydrogenase, mercurials,
847
Micrococcus aureus, see Staphylococcus
aureus
Micrococcus lactilyticus, succinate dehy-
drogenase,
malonate, 33, 49
Micrococcus lysodeikticus,
oxidative phosphorylation, mercurials,
873
respiration (endogenous), malonate, 168
succinate dehydrogenase, malonate, 26
Micrococcus pyogenes, see Staphylococcus
aureus
Micrococcus sodonensis, citrate oxidation,
malonate, 78
Microsporum audouini,
growth of, pyrithiamine, 529
respiration (endogenous), malonate, 168
Microsporum canis, respiration (endoge-
nous),
malonate, 169
Milkweed bug, see Oncopeltus
Mitochondria,
Ca++ uptake,
malonate, 209
mercurials, 909-910
K+ uptake,
malonate, 209
mercurials, 909, 914
Mg++ uptake, mercurials, 909
Na+ transport, mercurials, 909
swelling of,
ferricyanide, 678
iodine, 689
malonate, 210-211
Mitomycin, degradation of,
dehydroacetate, 623
Mitosis, see Cell division
Mixed disulfides, 639-640, 661, 663
Molybdate,
cofactor function, 614
uptake of, tungstate, 614
Molybdenum, tissue levels of,
tungstate, 614
Moniezia benedeni, succinate oxidation,
malonate, 54
Monoamine oxidase, mercurials, 816, 847
Monoamidines, structures of, 361
Monobenzone (Benoquin),
melanin formation in skin, 304
tyrosinase, 304
Monoethyl peroxide, catalase, 592, 694
Monofluorophosphate, phosphorylase, 406
L-Monoiodotyrosine, intestinal transport
of,
L-amino acids, 265
malonate, 207-208
Moraxella Iwoffi,
malate oxidation, malonate, 81
oxalacetate oxidation, malonate, 82
Morphine iV^-demethylase, analogs, 590,
597, 604
Mosquito, see Aedes
Mucate, see Galactarate
Mucinase, iodine, 686
Mung bean, see also Phaseolus aureus,
Rb"*" uptake in roots of, mercurials, 909
Muscle, see also Diaphragm
ATP levels in,
aminopterin, 585
o-iodosobenzoate, 721-722
mercurials, 876
citrate oxidation, malonate, 79
1198
SUBJECT INDEX
contractility, mercurials, 896, 937-940
contracture,
o-iodosobenzoate, 723
mercurials, 896, 938
tetrathionate, 699
creatine-P levels in, o-iodosobenzoate,
704
cycle intermediates concentrations, 89
excitability,
o-iodosobenzoate, 723
mercurials, 927
fructose- 1,6-diP levels, mercurials, 876
fumarate oxidation, malonate, 81
glycolysis,
in vivo, malonate, 125-126
mercurials, 875-877
glycolysis (aerobic), malonate, 129-130
glycolysis (anaerobic), o-iodosobenzo-
ate, 721-722
GSH levels, tetrathionate, 700
hydrogen peroxide, 696
malonate levels in vivo, 100-102
malonate metabolism in, 228, 233
membrane potentials,
malonate, 212
mercurials, 937-938
mercurial levels in, 959
Na+ pump, malonate, 212
oxidative phosphorylation, malonate,
119, 121,
pH changes in, mercurials, 876
pyrithiamine levels in vivo, 528
pyruvate oxidation, malonate, 75, 115
respiration, kojic acid, 350
respiration (endogenous),
dehydroacetate, 623-624, 629
malonate, 2, 175, 177-179
respiration (glucose),
malonate on insulin stimulation of,
164
respiratory quotient, malonate, 185
succinate accumulation, malonate, 94,
96
succinate dehydrogenase,
dicarboxylate ions, 35-37
malonate, 29, 31-32
succinate levels in vivo, malonate, 100-
102
succinate oxidation, malonate, 55-56
thiamine-PP levels, thiamine analogs,
525-526
Mussel, see Mytilus
Mutarotase (aldose 1-epimerase),
analogs, 413-414
relation of inhibition to cataract for-
mation, 414
Mycobacteria,
fatty acid biosynthesis, malonate, 148
glycerol oxidation, malonate, 164
growth of, D-cycloserine, 359
malonate formation in, 226
phospholipid biosynthesis,
malonate, 151
mercurials, 887
succinate dehydrogenase, oxalacetate,
36
Mycobacterium phlei,
malonate metabolism in, 228
respiration (endogenous),
malonate, 168, 237
malonic diethyl ester, 237
pyridine-3-sulfonate, 504
respiration (glycerol),
malonic diethyl ester, 237
pyridine-3-sulfonate, 504
respiration (lactate), tartronate, 238
Mycobacterium tuberculosis,
growth of, dehydroacetate, 632
infection by, malonic diethyl ester,
224
malonate metabolism in, 228-229
phospholipid biosynthesis, ferricyanide,
678
respiration, ferricyanide, 678
succinate dehydrogenase, malonate, 26
Mycorrhiza,
respiration (endogenous), malonate, 169
succinate oxidation, malonate, 53
Myelin, mercurial incorporation into, 950
Myoglobin, xanthine: cytochrome c oxi-
doreductase, 603
Myokinase, see Adenylate kinase
Myosin, see also Actomyosin,
association with actin, o-iodosobenzo-
ate, 723
mercurials, 938-940
SUBJECT INDEX
1199
titration of SH groups, tetrathionate,
698
Mytilus edulis, succinate dehydrogenase,
malonate, 33, 38
Myxoma virus, infectivity of,
mercurials, 978
Myxomycetes, see also Physarum
respiration (endogenous), malonate, 172
N
NAD (nicotine adenine dinucleotide),
analogs of,
formation from 3-acetylp>Tidine, 496
formation from 6-aminonicotinamide,
503-505
formation from isoniazid, 496
glutamate semialdehyde reductase, 507
NADH pyrophosphatase, 506, 511
NADPH:glutathione oxidoreductase,
506, 512
NADPHmitrite oxidoreductase, 512
reaction with p-mercuribenzoate, 774
synthesis of, pathways, 486
tissue levels of,
3-acetylpyridine, 495-496
6-aminonicotinamide, 505
a-NAD, NAD:NADP transhydrogenase,
510
NADase, see NAD nucleosidase
NAD glycohydrolase, see NAD nucleosi-
dase
NADH,
NAD kinase, 510
NAD nucleosidase, 491, 493
oxidation by ferricyanide, 673
oxidation of,
o-iodosobenzoate, 703
malonate, 20
NADH:aldehyde oxidoreductase, GSSG,
662
NADH:CoQ oxidoreductase, mercurials,
848
NADH:cytochrome c oxidoreductase,
o-iodosobenzoate, 710
mercurials, 848-849, 870
quinacrine, 552
NADH:DCPIP oxidoreductase, mercu-
rials, 849
NADH dehydrogenase, mercurials, 798,
870, 872
NADH: ferricyanide oxidoreductase,
mercurials, 849
nucleotides, 510
NADH:hydroxylamine oxidoreductase,
quinacrine, 552
NADHrHgOa oxidoreductase, hydrogen
peroxide, 693
NADH:lipoamide oxidoreductase (lipo-
amide dehydrogenase),
o-iodosobenzoate, 708
mercurials, coenzyme displacement,
804
NADH:menadione oxidoreductase,
mercurials, 850
nucleotides, 510
quinacrine, 552-553
NADH:methylene blue oxidoreductase,
quinacrine, 553
NADH:nitrate oxidoreductase, see also
Nitrate reductase,
mercurials, 850
NADH:nitrite oxidoreductase, see also
Nitrite reductase,
quinacrine, 547, 553
NADH oxidase,
ferricyanide, 676
hydrogen peroxide, 693
o-iodosobenzoate, 710-711
malonate, 63
mercurials, 847-848, 872
nicotinamide, 503
nucleotides, 511
quinacrine, 547, 553
NADH pyrophosphatase, nucleotides, 506
511
NADH:quinone oxidoreductase,
ferricyanide, 676
mercurials, 850
pH effects, 793
stimulation, 817
NADHrtetrazolium oxidoreductase,
mercurials, 850
quinacrine, 553
NAD kinase, nucleotides, 509-510
1200
SUBJECT INDEX
NAD:NADP transhydrogenase, nucleo-
tides, 510
NAD nucleosidase (NAD glycohydrolase,
NADase, DPNase),
analogs, 485-493
mercurials, 847
nicotinamide, 485-493
nicotinamide analogs, 504
NADP (nicotinamide adenine dinucleo-
tide phosphate),
glutamate semialdehyde reductase, 507
NAD nucleosidase, 489
NADPH:glutathione oxidoreductase,
512
NAD(P),
coenzyme functions of, analogs, 500-503
glutamate dehydrogenase, 508
NADPH:cytochrome c oxidoreductase,
511
nicotinamide deamidase, 512
NAD(P)diaphorase, mercurials mm'i'o, 926
NADP glycohydrolase, mercurials, 850
NADPH: cytochrome c oxidoreductase,
mercurials, 815, 817, 850-851
nucleotides, 511
NADPH dehydrogenase, benzoate, 349
NADPH diaphorase (old yellow enzyme),
iodine, 688
mercurials, 891
porphyrindin, 668
quinacrine, 548
NAD(P)H diaphorase, quinacrine, 553
NADPH:glutathione oxidoreductase, nu-
cleotides, 506, 512
NADPH:menadione oxidoreductase,
mercurials, 851
quinacrine, 553
NADPH:methemoglobin oxidoreductase,
mercurials, 774, 851
riboflavin, 543
NAD(P)H : methemoglobin oxidoreduc-
tase, quinacrine, 553
NADPHmitrate oxidoreductase, see Ni-
trate reductase
NADPHmitrite oxidoreductase, see also
Nitrite reductase
mercurials, 851
NAD, 512
NADPH oxidase, mercurials, 851
NADPH:trichloroindophenol oxidoreduc-
tase, mercurials, 851
NADP reductase (photosynthetic), mer-
curials, 891
NAD pyrophosphorylase (NMN adenyl-
transferase), deamino-ATP, 510
Nalorphine, morphine iV-demethylase,
590, 597, 604
a-Naphthol, chymotrypsin, 373
Naphthoresorcinol, tyrosinase, 304
Naphthylamines, chymotrypsin, 373
a-Naphthylmethylmalonate, chymotryp-
sin, 370
Naphthylpropionates, chymotrypsin, 369-
370
1-Naphthyl-sulfate, arylsulfatase, 443
2-Naphthyl-sulfate, arylsulfatase, 443
Neisseria gonorrhoeae, a-ketoglutarate
oxidation,
malonate, 79
Newatodirus filicollis,
citrate oxidation, malonate, 79
a-ketoglutarate oxidation, malonate, 80
respiration (endogenous), malonate, 174
succinate accumulation, malonate, 94
succinate oxidation, malonate, 54
Neoplectana glaseri, respiration (endoge-
nous),
malonate, 174
Neohydrin, see Chlormerodrin
Neon, nitrogen fixation, 291
Nephrotoxicity,
malonate, 219-220
mercurials, 924-925, 985
Nerve,
conduction,
o-iodosobenzoate, 724
malonate, 211-212
mercurials, 949-950
membrane potentials,
malonate, 211-212
mercurials, 949-950
thiamine analogs, 531
respiration (endogenous), malonate, 177
Neuraminidase, mercurials, 851
Neuroblasts,
o-iodosobenzoate, 724
SUBJECT INDEX
1201
mercurials, 964-965
porphyrindin, 670
Neuromuscular transmission, thiamine
analogs, 531-532
Neurospora crassa,
2-deoxyglucose utilization by, 387, 400
germination of ascopores, malonate,
195
growth of, oxythiamine, 520, 529
thiosulfate utilization, sulfate, 451-452
Neurospora crassa (poky strain), succinate
oxidation,
malonate, 53
Neutrophiles, phagocytosis by,
malonate, 203
Newcastle virus,
inactivation by mercurials, 978
infectivity of, mercurials, 978
Nicotinamidase, see Nicotinamide deami-
dase
Nicotinamide,
D-amino acid oxidase, 344
analogs of, 484-514
dehydrogenase inactivation by, 503
glucose dehydrogenase, 500-502
glucose-6-P dehydrogenase, 503
glutamate dehydrogenase, 863
heart, 500
lactate dehydrogenase, 500-503
malate dehydrogenase, 503
iV-methylnicotinamide formation from,
495
NAD degradation, 485-493
NADH oxidase, 503
NAD nucleosidase, 485-493
nicotinamide riboside phosphorylase,
487-488
6-phosphogluconate dehydrogenase, 503
respiration (endogenous), 503
respiration (glucose), 500
respiration (lactate), 500
Nicotinamide deamidase (nicotinamide
deaminase, nicotinamidase),
3-acetylp5Tidine, 498
o-iodosobenzoate, 711
mercurials, 783
nucleotides, 512
pyridine-3-sulfonamide, 504
Nicotinamide mononucleotide (NMN),
NAD nucleosidase, 489, 493
NADPH:glutathione oxidoreductase,
512
Nicotinamide mononucleotide adenyl-
transferase, see NAD pyrophosphoryl-
ase
Nicotinamide riboside, NAD nucleosidase,
492
Nicotinamide riboside phosphorylase, ni-
cotinamide, 487-488
Nicotinate,
D-amino acid oxidase, 342, 344, 346
catechol oxidase, 298-299, 301
glucose dehydrogenase, 500-502
glutamate dehydrogenase, 863
lactate dehydrogenase, 500-502
NAD nucleosidase, 490, 493
respiration (glucose), 500
Nicotine, NAD nucleosidase, 491
Nicotinic ethyl ester, NAD nucleosidase,
493
Nicotinic-hydrazide, NAD nucleosidase,
493
Nicotinic-hydrazide-NAD,
alcohol dehydrogenase, 497
lactate dehydrogenase, 497
Nicotinylalanine,
formation from tryptophan, 610
kynureninase, 610
k\Tiurenine hydroxylase, 610
urinary iV-methylnicotinamide, 610
Nicotinyl-D-phenylalaninamide, chymo-
trypsin, 372
Nicotinyl - d - tryptophanamide, chymo-
trypsin, 371
Nicotinyl-D-tyrosinamide, chymotrypsin,
371
Nicotinyl-D-tyrosine ethyl ester, chymo-
trypsin, 371
Nicotinyl-L-tyrosinemethylamine, chymo-
trypsin, 371
Nikethamide (Coramine),
NAD nucleosidase, 491
structure of, 488
Nitocra spinipes,
toxicity of Cu++, 962-963
toxicity of mercurials, 962-963
1202
SUBJECT INDEX
Nitrate,
creatine kinase, 446
I" transport by ciliary body, 267
phosphatase (acid), 441
tyrosinase, 301
Nitrate reductase, see also NAD(P)H:ni-
trate oxidoreductase
o-iodosobenzoate, 711
malonate, 65
quinacrine, 547, 554
tungstate, 615
Nitric oxide,
hydrogenase, 294
nitrogen fixation, 291-292
physical properties, 295
Nitrite, see also Nitrous acid
amylases, 660
oxidation of,
bromate, 450
chlorate, 450
cyanate, 450-451
iodate, 450
phosphate, 450
Nitrite reductase, see also NAD(P)H:ni-
trite oxidoreductase
Nitrite reductase, mercurials, 860
Nitrobacter, growth of,
chlorate, 450
m-Nitrobenzoate,
glutamate dehydrogenase, 330
o- and p-nitrobenzoate metabolism, 612
p - Nitrobenzoate, tjTosine : a - ketogluta-
rate transaminase, 306
Nitrobenzoates,
D-amino acid oxidase, 341, 348
catechol oxidase, 298-299
metabolism of, analogs, 612-613
p-Nitrobenzoyltryptophans, chymotryp-
sin, 374
3-o-Nitrobenzyl-4-methylthiazole, thiami-
nase, 524
5-Nitrofuroate, glutamate dehydrogena-
se, 330
Nitrogen, physical properties, 295
Nitrogenase, analogs, 291-296
Nitrogen fixation, analogs, 291-296
Nitromersol (Metaphen), structure of,
970
o-Nitrophenol,
catechol oxidase, 298
m- and p-nitrobenzoate metabolism,
612
2?-Nitrophenol,
D-amino acid oxidase, 348
catechol oxidase, 298
o- and p-nitrobenzoate metabolism, 612
polyphenol oxidase, 297
respiration (endogenous), 297
p-Nitrophenylacetate, carboxypeptidase,
365
p-Nitrophenylalanine, tyrosine : a - keto-
glutarate transaminase, 306
o-Nitrophenylpyruvate, pyruvate decar-
boxylase, 431
3-Nitropropionate (hiptagenate), succi-
nate dehydrogenase, 244
Nitroreductase, quinacrine, 554
2-Nitroresorcinol, tyrosinase, 304
Nitrosoreductase, quinacrine, 554
3-Nitro-L-tyrosine, tyrosinase, 305
Nitrous acid, see also Nitrite,
enzyme group oxidation, 657
Nitrous oxide,
nitrogen fixation, 291-294
physical properties, 295
NMN, see Nicotinamide mononucleotide
Nocardia corallina,
malonate formation in, 226
malonate occurrence in, 225
nitrobenzoate metabolism, analogs,
612-613
succinate accumulation, malonate 90
Nonanoate, kynurenine: a-ketoglutarate
transaminase, 608-609
Norepinephrine, see also Catecholamines,
biosynthesis of, pathways, 307
brain levels,
dopa decarboxylase inhibitors, 316-
318
a-methyl-m-tyrosine, 316
pyrogallol, 611
heart levels, dopa decarboxylase inhibi-
tors, 316-318
tissue levels, a-methyldopa, 315-320
urinary excretion, pyrogallol, 612
Norleucine, arginase, 337
SUBJECT INDEX
1203
Normetanephrine, urinary excretion of,
pyrogallol, 612
D-Norvaline, L-alanine dehydrogenase,
354
L-Norvaline, arginase, 337
Nuclei,
ATP level, malonate, 189
COa formation from glucose, 2-deoxy-
glucose, 393-394
glucose utilization, dehydroacetate, 624
respiration, malonate, 189
respiration (glucose), dehydroacetate,
624
Nucleic acids, see also Deoxyribonucleates
and Ribonucleates
biosynthesis of, fluorophenylalanines,
478-481
chymotrypsin, 457
fumarase, 465
lysozyme, 459
p-mercuribenzoate complex with, 744
mercuric complexes with, 741
Nucleosidediphosphate : polynucleotide
nucleotidyltransferase, see Polynucleo-
tide phosphorylaes
5 '-Nucleotidase, analogs, 471-472
Nucleotide incorporation enzyme, mer-
curials, 817
Oats, see also Avena,
malonate occurrence in, 224
Octanoate, see Caprylate
Oenanthate, see Heptanoate
Old yellow enzyme, see NADPH diapho-
rase
Oleyl-CoA, acetyl-CoA carboxylase, 614
Oncopeltus fasciatus, succinate dehydro-
genase,
malonate, 29
Onion roots, growth of,
mercurials, 966
Ophthalmate, glyoxylase, 594
Opsanus tau (toadfish) pancreas, respira-
tion (endogenous),
malonate, 175
Opsopyrrole-dicarboxylate, porphobilino-
gen deaminase, 600
Orcinol (5-methylresorcinol),
catechol oxidase, 297-298
structure of, 296
tyrosinase, 304
Ornithine,
arginase, 335, 337-338
arginine uptake in ascites cells, 338
histidase, 353
kynureninase, 595
D-Ornithine, carbamyl-P:ornithine trans-
carbamylase, 592
Ornithine carbamyltransferase (carbamyl-
P:L-aspartate carbamyltransferase),
analogs, 592
o-iodosobenzoate, 711
mercurials, 851
protection by substrates, 783
Ornithine decarboxylase, permanganate,
660
Orotate, dihydroorotase, 470
Orotate transphosphoribosylase, analogs,
473
Orotidylate decarboxylase, analogs, 472-
473, 479
Orthosphosphate, see Phosphate
Orthophosphoric diester phosphohydro-
lase, see Phosphodiesterase
Ova (rabbit), respiration (endogenous),
malonate, 183
Ovarian germinal epithelium, mitosis in,
malonate, 200
Oxacillin, penicillinase, 598
Oxalacetate,
glyoxylate reductase, 438
glyoxylate transacetatse, 594
/9-hydroxybutjrrate dehydrogenase, 594
keto-enol tautomerism of, 39
malate dehydrogenase, 596
malate dehydrogenase (decarboxylat-
ing), 597
oxalosuccinate decarboxylase, 597
oxidation of, malonate, 82
pyruvate decarboxylase, 430
succinate dehydrogenase, 36, 38-42
utilization of, malonate, 66-68
Oxalacetate decarboxylase,
2'-AMP, 507
analogs, 597
1204
SUBJECT INDEX
malonate, 63
Oxalacetate ethyl ester, succinate dehy-
drogenase, 39
Oxalate,
chelation with cations, 12
glutaniate decarboxylase, 328
glycolysis (anaerobic), 414
glyoxylate transacetatase, 594
D-hydroxy acid dehydrogenase, 435-
437
intercharge distance, 6
ionic length and volume, 188
kynurenine:a-ketoglutarate transami-
nase, 608
D-lactate:cytochrome c oxidoreductase,
435
lactate dehydrogenase, 435-436
D -lactate dehydrogenase, 437
permeability of erythrocytes to, 188
phosphatase (acid), 442
pyruvate decarboxylase, 430
succinate dehydrogenase, 35
tartronate semialdehyde reductase, 602
tyrosinase, 300
Oxalate decarboxylase,
malonate, 63
mercurials, 860
Oxalomalate, see also y-Hydroxy-a-keto-
glutarate,
decarboxylation to y-hydroxy-a-keto-
glutarate, 616
Oxalosuccinate decarboxylase,
analogs, 597
malonate, 63
mercurials, 852
Oxamate,
Crabtree effect, 435
glucose utilization, 435
glycolysis (aerobic), 434
glyoxylate reductase, 438
lactate dehydrogenase, 432-434
Na+ fluxes in HeLa cells, 434
pentose-P pathway, 435
pyruvate decarboxylase, 430
pyruvate oxidation, 434
respiration (glucose), 435
structure of, 432
toxicity, 434
Oxanilate, pyruvate decarboxylase, 431
Oxidants, see also specific oxidants
degree of SH group oxidation, 656-657
enzyme inhibitions, mechanisms of,
657-658
oxidation of protein groups, 657-658
Oxidative phosphorylation,
benzoate, 348
Ca++, 453
dehydroacetate, 623
galactoflavin, 544
iodide, 689
iodine, 688-689
o-iodosobenzoate, 722
malonate, 118-122
mercurials, 872-874, 927
quinacrine, 556-557
thiophosphate, 447-448
Oxidized glutathione (GSSG), see Gluta-
thione (oxidized)
u-Oximinoglutarate, glutamate decarbo-
xylase, 327
<5-Oximinolevulinate, aminolevulinate de-
hydrase, 591
2-Oxoglutarate, see a-Ketoglutarate
2-Oxo-4-imidazolidinecaproate, see Des-
methyldesthiobiotin
Oxybiotin, structure of, 588
Oxybiotinsulfonate, yeast fermentation,
588-589
Oxygen,
hydrogenase, 293
inactivation of enzymes, 658-659
nitrogen fixation, 291-294
physical properties, 295
pyruvate oxidase, 659
respiration of brain, 658
SH group oxidation by, 658
uptake, see Respiration
Oxypyrithiamine, structure of, 517
Oxythiamine,
blood lactate, 520
blood pressure, 532
blood pyruvate, 520, 527
central nervous system, 527, 530
glycogen in liver, 520
heart rate in vivo, 527
miosis, 532
SUBJECT INDEX
1205
nerve membrane potentials, 531
Neurospora growth, 520, 529
phosphorylation of, 519
pyruvate accumulation, 520
pjTuvate oxidation in vivo, 529-521
rat growth, 527
structure of, 517
thiaminase, 523-524
thiamine deficiency, 530-532
thiamine kinase, 522-523
thiamine levels in tissues, 525-527
toxicity, 516, 530-531
transketolase in vivo, 522
urinary thiamine, 525
Vibrio growth, 522
Oxythiaminediphosphate,
acetoin formation from pyruvate, 519
pyruvate decarboxylase, 518
transketolase, 519
Oxythiaminetriphosphate,
pyruvate decarboxylase, 518
pyruvate oxidase, 519
Oxyurea, urease, 603, 610
Oyster, see also Crassostrea and Saxostrea
Oyster eggs,
glycolysis, mercurials, 874-875, 884
respiration (endogenous), mercurials,
882, 884
succinate oxidation, malonate, 22
Oyster muscle, succinate oxidation,
malonate, 22
Oyster spermatozoa,
glycolysis, mercurials, 884
respiration (endogenous), mercurials,
882, 884
Palmitate, oxidation of,
2-deoxyglucose, 397
Palmityl-CoA, citrate synthetase, 614
Pancreas, respiration (endogenous),
malonate, 175, 177
Pantetheine, structure of, 587
Pantoate:^-alanine ligase (AMP),
acetate, 597
^-alanine analogs, 597-598
Pantothenate,
analogs of, 586-588
acetyl transfer, 587
bacterial growth, 587
choline acetylase, 587
coenzyme A formation, 586-587
pantothenate biosynthesis, 588
biosynthesis of, analogs, 588
conversion to coenzyme A, 586-587
structure of, 587
Pantoylaminoethanethiol,
coenzyme A formation from pantethe-
ine, 587
structure of, 587
sulfonamide acetylation, 587
Pantoyltaurine,
bacterial growth, 587
choline acetylase, 587
structure of, 587
pApA, phosphodiesterase, 473
Papain,
analogs, 375
ferricyanide, 673, 676
hydrogen peroxide, 691, 693-694
iodine, 682-683, 686
mercurials, 769-770, 804
crystalline mercuric ion complex,
769-770
relation to SH groups, 804
porphyrindin, 667-668
succinyl peroxide, 694
Papilloma, malonate and succinate levels
in vivo, 102
Paracentrotus lividus eggs, development
of,
o-iodosobenzoate, 726-727
Paramecium caudatum,
cycle intermediate oxidations in, ma-
lonate, 79-82
mercurial toxicity, 981-982
motility of,
<rans-aconitate, 274
malonate, 203
mercurials, 981
pyruvate oxidation, malonate, 74
respiration (endogenous),
iraws-aconitate, 273
malonate, 173
succinate dehydrogenase, malonate, 28,
50
1206
SUBJECT INDEX
Parasorbic acid, 617
Paris daisy stem cultures, growth of,
malonate, 197
Pasteurella muUocida,
fumarate oxidation, malonate, 81
glutamate oxidation, malonate, 187
isocitrate oxidation, malonate, 79
pyruvate oxidation, malonate, 74
Patulin (clavacin), 617
Pea leaves,
malonate occurrence in, 224
succinate accumulation, malonate, 91
Peanut cotyledons, fatty acid oxidation,
malonate, 136
Peanut mitochondria,
butyrate oxidation, malonate, 137
malonate metabolism in, 228, 231
phospholipid biosynthesis, malonate,
151
propionate metabolism, malonate, 145
Pea roots, mitosis in,
malonate, 197
Peas, sea also Pisum sativum,
respiration (endogenous), malonate, 170
Pea seedlings,
a-ketoglutarate utilization, malonate, 84
oxidative phosphorylation, malonate,
122
Pea stems, growth of,
mercurials, 966
Peloscolex velutinus, respiration (endo-
genous),
malonate, 174
Penicillic acid, 617
Penicillinase,
analogs, 598-599, 615, 688
configurational changes, analogs, 249
ferricyanide, 676
iodine, 688
o-iodosobenzoate, 711, 717
pH effects, 717
mercurials, pH effects, 793
Penicillin-G, renal transport of,
dehydroacetate, 625
Penicillium chrysogenum,
acetate oxidation,
malonate, 77
malonic diethyl ester, 237
lactate oxidation, malonate, 78
respiration (endogenous), malonate, 169
succinate dehydrogenase,
malonate, 26
oxalacetate, 36
Penicillium citrinum, growth of,
dehydroacetate, 633
Penicillium cyclopium, malonate occur-
rence in, 226, 228
Penicillium digitalum, growth of,
dehydroacetate, 632
Penicillium expansum., growth of,
dehydroacetate, 632
Penicillium funiculosum, malonate oc-
currence in, 225
Penicillium notatum, growth of,
mercurials, 973
resistance to mercurials, 983
Penicillium oxalicum, respiration (glu-
cose),
malonate, 133-134
Penicillium roqueforti, tolerance to mer-
curials, 983
Pentanoate, see Valerate
Pentose-phosphate isomerase, o-iodoso-
benzoate, 711
Pentose-phosphate pathway,
2-deoxyglucose, 393-394
malonate, 130-132
mercurials, 885-886
oxamate, 435
pyrithiamine-resistant S. aureus, 529
quinacrine, 560
Pepsin
dehydroacetate, 622
iodine, 680, 682-683, 688
macroions, 457-458
mercurials, 860
permanganate, 657
Peptidases, mercurials, 860
Periodate,
chymotrypsin, 657
/S-fructofuranosidase, 660
ovalbumin SH groups, 657
protein oxidation by, 657
Periplaneta americana, succinate dehydro-
genase,
malonate, 29
SUBJECT INDEX
1207
Periwinkle stem cultures, growth of,
malonate, 197
Permanganate,
enzyme inhibitions, 659-660
Fiisarium conidial growth, 660
insulin, 657
oxidation,
of amino acids, 657
of casein, 657
of proteins, 657
pepsin, 657
Permeability,
to dicarboxylate ions, 188
to malonate, 186-192
to mercurials, 900-907
Peroxidase,
jj-coumarate analogs, 599
iodine. 682, 686
mercurials, 860
Peroxidases, see also Hydrogen peroxide,
Monoethyl peroxide, and Succinyl per-
oxide,
acetylcholine response, 696
chemical properties, 690-691
enzyme inhibitions, 691-694
glycolysis, 695
intestine, 696
muscle, 696
respiration, 695
toxicity, 696
tumor growth, 695
pH, effects of,
ADP on ATPase, 445
carbobenzoxyglutamate on papain, 375
copper on ^-glucuronidase, 795
dehydroacetate,
on growth of microorganisms, 633
on succinate dehydrogenase, 622
fatty acids on kynurenine transami-
nase, 609
ferricyanide oxidation of hemoglobin,
671
hydrogen peroxide on ATPase, 691
iodine,
bactericidal action, 690
on enzymes, 688
oxidation of cysteine, 680
o-iodosobenzoate,
on enzymes, 716-717
oxidation of SH groups, 657, 702
macroionic inhibitions, 454-457, 461,
464
malonate,
on cardiac metabolism, 214-215
on succinate oxidation, 51, 56
permeability to, 189-192
mercurials,
on ATPase, 867
on enzymes, 790-797
on K+ efflux, 898
on luminescence, 889
reactions with proteins, 760-761
reactions with SH groups, 749-750
titration of 3-phosphoglyceraldehyde
dehydrogenase, 806
toxicity to heart, 944-945
mercuric ion,
activation of glycerate-2,3-diphos-
phatase, 820
formation from organic mercurials,
931-933
metal ion reactions with thiols, 638
methylurea on urease, 610
oxidation of enzyme SH groups, 664
quinacrine on D-amino acid oxidase,
557-558
silver on y3-glucoronidase, 795
Phage (coli), see Coliphage
Phage (staphylococcal), see Staphylococ-
cal phage
Phagocytosis,
o-iodosobenzoate, 728
kojic acid, 350
malonate, 223
Phaseolus aureus, see also Mung bean,
citrate oxidation, malonate, 78
a-ketoglutarate oxidation, malonate, 80
oxidative phosphorylation, malonate,
120
Phaseolus coccineus (runner bean),
malonate occurrence in, 225
respiration, malonate, 225
Phaseolus seeds, glutamate metabolism,
pyrithiamine, 522
Phaseolus vulgaris (bush bean),
malonate metabolism in, 226, 228, 232
1208
SUBJECT INDEX
malonate occurrence in, 225-226
succinate dehydrogenase, malonate, 33
2-Phenantryl-sulfate, arysulfatase, 443
Phenbenicillin, penicillinase, 598
Phenethicillin, penicillinase, 598
Phenol,
D-amino acid oxidase, 348
dehydroshikimate reductase, 606
Phenol-a-glucoside,
/3-glucosidase, 417
taka-/3-glucosidase, 271
Phenol oxidase, see also Catechol oxidase
analogs, 296-302
Phenol red, renal transport of,
dehydroacetate, 626
malonate, 205
mercurials, 921
Phenol sulfokinase, 3'-AMP-5'-P, 473
Phenolsulfonphthalein, renal transport of,
dehydroacetate, 625-626
2-Phenoxyethanol, chymotrypsin, 370
Phenylacetamide, chymotrypsin, 372
Phenylacetate,
D-amino acid oxidase, 342, 346
ammonia formation in kidney, 348
carboxypeptidase, 363-366
catechol oxidase, 298
chymotrypsin, 370, 372
dopa decarboxylase, 312
glutamate decarboxylase, 329
p-hydroxyphenylpuruvate oxidase, 306
tyrosinase, 300-301
Phenyl-iV-acetylglucosaminide, iV-acetyl-
/3-galactosaminidase, 420
Phenylalaninamides, cathepsin C, 375
Phenylalanine,
analogs of, dopa decarboxylase, 308
blood serotonin, 325
brain serotonin, 325
dipeptidase, 367
glutamate decarboxylase, 329
histidine decarboxylase, 353
pyruvate decarboxylase, 430
tryptophan hydroxylase, 325
D -Phenylalanine ,
carboxypeptidase, 365, 367
E. coli growth, 268
L-Phenylalanine,
amino acid transport by brain, 266
arginase, 337
carboxypeptidase, 366
cathepsin C, 375
tyrosinase, 305
tyrosine:a-ketoglutarate transaminase,
306
Phenylalanine activating enzyme, see l-
PhenylalanineisRNAligase (AMP)
Phenylalanine deaminase,
analogs, 355
mercurials, 852
Phenylalanine ^-hydroxylase, analogs,
354, 599-600
L-Phenylalanine:sRNA ligase (AMP), ana-
logs, 354-355
Phenylalanylglycine, cathepcin C, 375
Phenylbutyramide, chymotrypsin, 372
a-Phenylbutyrate,
cholesterol biosynthesis, 614
fatty acid biosynthesis, 614
y-Phenylbutyrate,
carboxypeptidase, 365-366
chymotrypsin, 370, 372
kynurenine:a-ketoglutarate transami-
nase, 608-609
Phenylethylam ine ,
dopa decarboxylase, 308
dopamine ^-hydrolase, 320
phenylalanine ^-hydroxylase, 600
Phenyl-a-glucopyranoside, a-glucosidase,
416, 423
Phenylglucosides,
a-amylase, 420
maltose transglucosylase, 415
Phenylglycine, derivatives of,
dopa decarboxylase, 312
Phenylglyoxylate, glyoxylate reductase,
438
Phenylisocyanate, urease, 649
Phenylketonuria (phenyl pyruvate oligo-
phrenia),
inhibition of amino acid transport into
brain, 266
inhibition of dopa decarboxylase, 314
inhibition of glutamate decarboxylase,
329
SUBJECT INDEX
1209
inhibition of pyruvate metabolism,
429-430
inhibition of tryptophan hydroxylase,
325-326
inhibition of tyrosinase, 305
Phenyllactate,
glutamate decarboxylase, 329
pyruvate decarboxylase, 430
tryptophan hydrolase, 325
Phenylmercuric acetate, see Phenylmer-
curic ion
Phenylmercuric ion (PM), see also Mer-
curials,
amino acid complexes of, 744
3-Phenyl-4-methylthiazole, thiaminase,
524
Phenylphosphate, arylsulfatase, 443
Phenylpropionamide, see Hydrocinnamide
y3-Phenylpropionate, see Hydrocinnamate
Phenylpyruvate,
accumulation in phenylketonuria, 429-
430
acetoin formation from pyruvate, 430
dopa decarboxylase, 312-314
glutamate decarboxylase, 329
glycerate dehydrogenase, 430
33-hydroxyphenylpyruvate oxidase,
305-306
lactate dehydrogenase, 437
melanin formation, 305
metabolism of, 429-430
pyruvate decarboxylase, 431-432
pyruvate metabolism, 430
pyruvate oxidase, 430
tryptophan hydroxylase, 325
tyrosinase, 305
Phenylpyruvate oligophrenia, see Phenyl-
ketonuria
/3-Phenylserine,
phenylalanine deaminase, 355
L-phenylalanine:sRNA ligase (AMP),
354
Phenylsulfate, arylsulfatase, 443
Phloretin-phosphate polyesters, hyaluro-
nidase, 461
Phlorizin,
mitochondrial swelling, 210
renal transport of PAH, 205
Phloroglucinol,
D-araino acid oxidase, 344
catechol oxidase, 297
tyrosinase, 304
Phloroglucinol-phosphate polymer, hya-
luronidase, 461
Phosphatase, see also Phosphatase (acid)
and Phosphatase (akaline),
analogs, 439-443
arsenate, 439-440
borate, 439-440
ferricyanide, 676
iodine, 686
mercurials in vivo, 926-927
oxidation of, 657
permanganate, 660
silicate, 439-440
substrate inhibition, 439
tartrates, 440-442
Phosphatase (acid),
active center of, 440-442
alginate, 464
anionic polymers, 443
o-iodosobenzoate, 711, 718
macroions, 464-465
malonate, 63
mercurials, 772-773, 778, 817
Mg++ analogs, 452-453
polyestradiol- phosphate, 464
polyphloretin-phosphate, 464
tartronate, 238
Phosphatase (alkaline),
5-fluorouracil, formation of abnormal
enzyme, 479
o-iodosobenzoate, 711
mercurials, 772-773, 817, 860, 926
Phosphate,
active transport of, malonate, 209-210
arylsulfatase, 443-444
carbamyl phosphatase, 439
choline sulfatase, 444
creatine kinase, 446
glucose dehydrogenase, 501
glucose-6-P dehydrogenase, 411
glycolysis (anaerobic), 414
nitrite oxidation, 450
phosphatases, 439
phosphodeoxyribomutase, 412
1210
SUBJECT INDEX
3-phosphoglyceraIdehyde dehydroge-
nase, 409
phosphopentose isomerase, 411
pyrophosphatase, 439
renal transport of,
dehydroacetate, 625
malonate, 205
ribulose-P carboxylase, 412
serum level of, malonate, 219
transaldolase, 412
transketolase, 412
triose-P isomerase, 412
uptake by erythrocytes, mercurials,
910
uptake by staphylococci,
analogs, 267
mercurials, 910, 912-913
urinary excretion of, malonate, 206
yeast level of, mercurials, 885
Phosphatidate phosphatase, cystine, 662
3'-Phosphoadenosine-5'-phosphosulfate
reductase, quinacrine, 554
Phosphoarabinose isomerase, analogs, 411
Phosphodeoxyribomutase,
analogs, 413
phosphate, 412
Phosphodiesterase (orthophosphoric di-
ester phosphohydrolase), analogs, 473
Phosphoenolpyruvate carboxylase,
o-iodosobenzoate, 711
mercurials, 852
Phosphoenolpyruvate carboxytransphos-
phorylase, mercurials, 852
Phosphofructokinase,
cycle ingermediates, 385-386
cycHc 3 ,5 -AMP, 474
mercurials, 852
Phosphoglucomutase,
l,5-anhydro-D-glucitol-6-P, 379
D-glucosamine, 382
glucose-6-P, 413
GSSG, 662
o-iodosobenzoate, 711, 716
malonate, 130
mercurials, 804, 810,
rate of inhibition, 810
relation to SH groups, 804
oxygen inactivation of, 659
6-Phosphogluconate, phosphoglucose iso-
merase, 406
Phosphogluconate dehydrogenase,
2'-AMP, 507
o-iodosobenzoate, 711
nicotinamide, 503
Phosphoglucose isomerase,
analogs, 406-407
l,5-anhydro-D-glucitol-6-P, 379
ATP, 474
2-deoxyglucose-6-P, 390
o-iodosobenzoate, 711
mercurials, 839
3-Phosphoglyceraldehyde, anolase, 409
3-Phosphoglyceraldehyde dehydrogenase
(triose-P dehydrogenase),
active center of, 641
6-aminonicotinamide in vivo, 505
analogs, 408-409
cystamine monosulfoxide, 663
ferricyanide, 676-677
p-fluorophenylalanine, replacement of
phenylalanine in, 351
GSSG, 661-662
hydrogen peroxide, 694-695
iodine, 683, 687
o-iodosobenzoate, 704, 714
malonate, 63
mercurials, 650, 766, 770, 776, 783,
785-786, 788, 802, 804, 806, 810, 812,
817, 824, 826-827, 852, 876
coenzyme displacement, 785-786
denaturation, 788
protection, 783
rate of inhibition, 810
relation to SH groups, 802, 804, 812
reversal by cysteine, 824, 826
serum effect, 776
spontaneous reversal, 827
stimulation, 817
titration of SH groups, 806
p-mercuribenzoate, 650, 766, 852
titration of SH groups, 766
mercuric ion, crystalline complex of,
770
oxygen inactivation of, 659
porphyrexide, 668
pterin-6-aldehyde, 288
SUBJECT INDEX
1211
succinyl peroxide, 694
tetrathionate, 699
2-Phosphoglycerate, glycerate-2,3-diphos-
phatase, 413
3 -Phosphogly cerate ,
enolase, 410
glycerate-2,3-diphosphatase, 413
Phosphoglycerate kinase, o-iodosoben-
zoate, 711
Phosphoglycerate mutase, mercurials, 852
Phosphoglycerol dehydrogenase, o-iodoso-
benzoate, 711
Phosphohalidase, see DFPase
Phospholactate, enolase, 410
Phospholipase, mercurials, 860
Phospholipids, biosynthesis of,
malonate, 151
mercurials, 887
Phosphomevalonate kinase,
o-iodosobenzoate, 711
mercurials, 852
Phosphomonoesterase, mercurials, 788
Phosphonoacetate, succinate dehydroge-
nase, 243
/S-Phosphopropionate,
intercharge distance, 7
succinate dehydrogenase, 242-243
Phosphopentose isomerase, see also Phos-
phoarabinose isomerase
analogs, 411
Phosphopyruvate carboxylase, see Phos-
phoenolpyruvate carboxylase
Phosphopyruvate hydratase, see Enolase
Phosphoribomutase, 2,3-diphosphoglyce-
rate, 413
5-Phosphoribonate,
phosphopentose isomerase, 411
phosphoribulokinase, 413
Phosphoribosyl-ATP pyrophosphorylase,
feedback inhibition by histidine, 351
trypsin inactivation of, 351
Phosphoribosylpyrophosphate amido-
transferase,
ADP and ATP, 474
mercurials, 853
Phosphoribulokinase, 5-phosphoribonate,
413
Phosphorylase,
analogs, 405-406
D-glucosamine, 382
mercurials, 648-649, 789, 803-804, 808,
811-813, 852-853
rate of inhibition, 811-812
relation to SH groups, 803-804, 808,
813
splitting into subunits, 789
a-methylglucoside, 271-272
SH groups of, 649
Phosphorylase kinase, Ca++, 453
Phosphorylphosphatase, mercurials. 817
0-Phosphoserine phosphatase,
alanine, 270-271
mercurials, 853
serine, 270
Phosphothreonine, L-threonine synthe-
tase, 357
Phosphotransacetylase,
lipoate analogs, 590
Na+ and Li+, 452
Phosphotransferases, see also Kinases,
analogs, 444-447
Photophosphorylation ,
mercurials, 892
quinacrine, 557
Photosynthesis ,
a - hydroxy - 2 - pyridinemethanesulfo-
nate, 439
malonate, 163-164
mercurials, 891-892
threose-2,4-diphosphate, 409
xylose, 414
Phthalate,
D-amino acid oxidase, 341, 344
aspartate : a - ketoglutarate transami-
nase, 334
glutamate dehydrogenase, 331
intercharge distance, 6
ionization constants, 8
kynurenine:a-ketoglutarate transami-
nase, 607-608
succinate dehydrogenase, 37
tyrosinase, 300
m-Phthalate, see Isophthalate
p-Phthalate see Terephthalate
Phycomyces blakesleeanus,
1212
SUBJECT INDEX
carbohydrate metabolism, thiamine
analogs, 529
resistance to pyrithiamine, 529
Physarum polycephalum , succinate dehy-
drogenase,
malonate, 28
Picolinate, NAD nucleosidase, 488
Pimelate,
aspartate : a - ketoglutarate transami-
nase, 334
glutamate decarboxylase, 328
ionization constants, 8
kynurenine:a-ketoglutarate transami-
nase, 595,608
Pink disease, see Acrodynia
Pinus lambertiana (sugar pine), succinate
dehydrogenase,
malonate, 27
Pinus taeda (loblolly pine) roots, phos-
phate uptake,
malonate, 209
iV-Piperidinomethylnicotinamide, inhibi-
tion of various oxidations, 503
Pisuni sativum, succinate dehydrogenase,
malonate, 27
Pituitary, succinate dehydrogenase,
malonate, 31
Placenta,
acetate oxidation, malonate, 78
lipid biosynthesis, malonate, 147, 149
malonate metabohsm in, 228, 233
pyruvate oxidation, malonate, 77
respiration (endogenous), malonate, 179
succinate oxidation, malonate, 55
Plants, growth of,
malonate, 196-197
mercurials, 965-968
Plasma, see Blood
Plasmodium berghei, glucose utihzation,
quinacrine, 560
Plasmodium gallinaceum, succinate accu-
mulation,
malonate, 93
Plasmodium lophurae, respiration,
quinacrine, 560
Plasmodium vivax,
lactate oxidation, malonate, 78
pyruvate oxidation, malonate, 74
succinate oxidation, 53
Pleuropneumonia virus, infectivity of,
mercurials, 979
Plumaria elegans, mercurial toxicity, 967
PM, see Phenylmercuric ion
Pneumococcus polysaccharide, lysozyme,
459
Pneumonitis virus, proliferation of,
D-cycloserine, 360
malonate, 194
Poliomyelitis virus, infectivity of,
mercurials, 979
Pollen, malonate metabolism in, 228
Polyacrylate,
phosphatase (acid), 465
trypsin, 457
Polyadenylate, polycytidylate phosphory-
lase, 463
Polyalanine, pepsin, 458
Polyaspartate,
pepsin, 458
ribonuclease, 462
Polyestradiol-phosphate, phosphatase
(acid), 464
Polyethylenesulfonate, phosphatase (acid)
464-465
Polygalacturonase, galacturonate, 421
Polyglucose (oxidized), lysozyme, 459
Polyglucose-sulfate,
lipoprotein lipase, 463
lysozyme, 459
ribonuclease, 462
Polyglutamate,
lysozyme, 459
pepsin, 458
ribonuclease, 462
trypsin, 456-457
Polyhydroquinone, phosphatase (acid),
464-465
Polylysine,
lipoprotein lipase, 463
pepsin, 457-458
trypsin, 456-457
Polymeric sulfonates,
lysozyme, 459
ulcer reduction, 458
Polymers, see Macroions
Polymethacrylate, hyaluronidase, 459
SUBJECT INDEX
1213
Polynucleotide nucleotidyltransferase, see
Polynucleotide phosphorylase
Polynucleotide phosphorylase,
analogs, 474
macroions, 463
Polyornithine, pepsin, 457-458
Polyphenol oxidase, see Catechol oxidase,
Phenol oxidase, and Tyrosinase
Polyphloretin-phosphate, hyaluronidase,
461
Polyphloroglucinol-phosphate, hyaluroni-
dase, 461
Polystyrenesulfonate, hyaluronidase, 459
Polyuridylate, polyadenylate phosphory-
lase, 463
Polyvinyl-sulfate, ribonuclease, 463
Polyxenyl-phosphate, phosphatase (acid),
443, 464
Porphobilinogen deaminase, analogs, 600
Porphyra perforata,
K+-Na+ transport, mercurials, 908-909
respiration (endogenous), mercurials,
881
Porphyrexide,
chemical propeties, 664-666
enzyme inhibitions, 667-669
oxidation of amino acids, 666
oxidation of proteins, 666-667
oxidation of SH groups, 666
oxidation-reduction potential, 665
paramagnetism, 664-665
spiro analogs, 666
stability, 665
Porphyridium cruentum, respiration (en-
dogenous),
malonate, 169
Porphyrin, biosynthesis of,
ferricyanide, 678
malonate, 158-163
mercurials, 888
pathways of, 159
Porphyrindin,
chemical properties, 664-666
determination of SH groups, 666-667
enzyme inhibitions, 667-669
heart, 669
neuroblastic damage, 670
oxidation of proteins, 666-667
oxidation of SH groups, 666
oxidation-reduction potential, 665
paramagnetism, 664-665
Sarcoma 37, 670
skin, 669-670
spiro analogs, 666
stability, 665
urease, 643
Potassium,
active uptakes, mercurials, 908-909
barley root uptake, malonate, 209-210
erythrocytic efflux, mercurials, 903-905
mitochondrial uptake,
o-iodosobenzoate, 722
malonate, 209
plasma level, malonate, 206
renal transport,
dehydroacetate, 626
mercurials, 921, 928, 936
renal uptake, malonate, 205-206
yeast efflux, mercurials, 898-900
Potato,
amino acid levels in, malonate, 106
amino acid metabolism, 154
carbohydrate biosynthesis, malonate,
106
citrate formation from glucose, ma-
lonate, 105-106
glucose metabolism, 106, HI, 132-133
lipid biosynthesis, malonate, 149
pentose-P pathway, malonate, 132
respiration (endogenous),
y-hydroxy-a-ketoglutarate, 616
malonate, 172, 182
succinate accumulation, malonate, 91,
106
succinate dehydrogenase, 19, 27
sucrose biosynthesis, malonate, 132
Potato virus X, splitting into subunits by
mercurials, 980
Proflavine,
Lactobacillus growth, 537
structure of, 537
Prolidase (imidodipeptidase), mercurials,
protection by Mn++, 783
D-Proline, Zl^-pyrroline-5-carboxylate re-
ductase, 355
L-Proline,
1214
SUBJECT INDEX
L-amino acid oxidase, 340
arginase, 337
A i-pyTroline-5-carboxylate dehydroge-
nase, 336, 355
A ^ - pyrroline - 5 - carboxylate reductase,
355
Proline oxidase, kojic acid, 350
Propane-tricarboxylate,
aconitase, 240
isocitrate dehydrogenase, 240
a-ketoglutarate oxidation, 240
pyruvate oxidation, 240-241
succinate dehydrogenase, 240-241
PropicilHn, penicilHnase, 598
Propionate,
acetate metabolism, 613-614
acetyl-CoA synthetase, 613
carboxypeptidase, 366
fatty acid synthesis from acetate, 613
formation from lactate, malonate, 165
glutamate decarboxylase, 328
kynurenine: a-ketoglutarate transami-
nase, 608
lactate dehydrogenase, 436
lipid metabolism, 613-614
metabolism of, malonate, 144-145
pantoate:/?-alanine ligase, 598
pyruvate decarboxylase, 430
Propionibacterium, methylmalonate in,
224
Propionibacterium pentosaceum, glycerol
fermentation,
malonate, 164
Propionibecterium shermanii, methylma-
lonyl-CoA and succinyl-CoA intercon-
version in, 235
Propionyl-CoA, fatty acid biosynthesis,
613-614
Propionyl-CoA carboxylase, mercurials,
853
Prostate,
malonate metabolism in, 228
respiration (endogenous), malonate, 179
succinate oxidation, malonate, 55
Protease (Aspergillus),
hydrogen peroxide, 693
iodine, 687
Protease (Etroplus), o-iodosobenzoate, 712
Protease (Rastrelliger), o-iodosobenzoate,
712
Protease (Trifolium), hydrogen peroxide,
693
Protection, see also specific enzymes and
inhibitors,
against o-iodosobenzoate, 717
against mercurials, 778-779, 783-784
against SH reagents, 41-42, 650-651
Proteinase (Aspergillus),
cystine, 662
iodine, 687
Proteinase (Clostridium),
iodine, 687
mercurials, 793
Proteinase (lens),
GSSG, 662
o-iodosobenzoate, 712
Proteinase (mackerel), ferri cyanide, 676
Proteinase (Pseudomonas), permanganate,
660
Proteinase (yeast), mercurials, 791, 793
Protein disulfides reductase,
mercurials, 853, 926
quinacrine, 554
riboflavin, 543
Proteins,
biosynthesis of,
amethopterin, 585
a-amino-/3-chlorobutyrate, 351
jff-azaguanine, 478
2-deoxyglucose, 399
p-fluorophenylalanine, 351
5-fluorouracil, 479
folate analogs, 585
malonate, 155-157
mercurials, 887-888
chymotrypsin inhibition, 457
oxidation of,
ferricyanide, 671-672
porphyrexide, 666-667
porphyrindin, 666-667
tetrathionate, 697-698
reactions with,
iodine, 680-682
o-iodosobenzoate, 703-704
mercurials, 751-768
SUBJECT INDEX
1215
Proteus morganii, infections by,
malonate, 221-222
Proteus vulgaris,
acetate accumulation, benzoate, 349
citrate oxidation, malonate, 78, 86
glucose metabolism, benzoate, 349
growth of, mercurials, 972
isocitrate oxidation, malonate, 79, 86
pyruvate metabolism, benzoate, 349
succinate oxidation, malonate, 52
Protocatechuate, see 3,5-Dihydroxyben-
zoate
Protoporphyrin, biosynthesis of,
arsenite, 162
2,4-dinitrophenol, 162
fluoroacetate, 162
malonate, 162
Pseudomonas,
fumarate oxidation, difluoromalonate,
239
malonate inhibition, effect of drying,
187
malonic semialdehyde occurrence in,
225
succinate oxidation, difluoromalonate,
239
Pseudomonas aeruginosa,
enzyme induction in, mercurials, 888
growth of, dehydroacetate, 632
malonate metabolism in, 228-229
succinate oxidation, malonate, 52
Pseudomonas fluorescens,
growth of, malonate, 195
malonate metabolism in, 228, 230-231
Pseudomonas hydrophila, pentose oxida-
tion,
malonate, 132
Pseudomonas saccharophila,
a-amylase synthesis in, D-asparagine,
269
fumarate oxidation, malonate, 81
pyruvate oxidation, malonate, 74
succinate oxidation, malonate, 52
Psicofuranine, xanthosine-5'-P aminase,
476, 481
9 - D - Psicofuranosyl - 6 - aminopurine, see
Psicofuranine
Psittacosis agent.
inactivation of,
o-iodosobenzoate, 728
mercurials, 979-980
proliferation of,
D-cycloserine, 360
o-iodosobenzoate, 728
malonate, 194
mercurials, 981
Psittacosis-lymphogranuloma virus, in-
fectivity of,
mercurials, 979
Pteridines,
structures of, 287
xanthine oxidase, 285-289
Pteridylaldehyde, see Pterin-6-aldehyde
Pterin, structure of, 287
Pterin-6-aldehyde,
glucose oxidase, 288
guanase, 288
oxidation by xanthine oxidase, 288
3 - phosphoglyceraldehyde dehydroge-
nase, 288
potentiation of 8-azaguanine carcino-
stasis, 288
quinine oxidase, 288
structure of, 287
urate levels in tissues in vivo, 288
uricase, 288
xanthine oxidase, 285-289
Pterin-6-carboxylate, xanthine oxidase,
289
Pteroate, xanthine oxidase, 289
Pteroylaspartate, dopa decarboxylase,
586
Pteroyglutamate, see Folate
pTpTpTpT, phosphodiesterase, 473
Puccinia (stem rust),
germination of uredospores, malonate,
195-196
leaf infections by, oxythiamine, 529
respiration (endogenous), malonate, 169
Pullularia pullulans,
acetate oxidation, malonate, 77
malonate metabolism in, 190, 228
pyruvate oxidation, malonate, 74
Pupillary size, thiamine analogs, 532
Purines,
ionization of, 280
1216
SUBJECT INDEX
riboflavin complexes of, 545
Putrescine,
kynureninase, 595
structure of, 361
Pyrazoloisoguanine,
structure of, 280
xanthine oxidase, 281-282
Pyridine,
glucose dehydrogenase, 501
NAD nucleosidase, 488
Pyridine-2-carboxylate, NAD nucleosi-
dase, 491
Pyridine-2,6-dicarboxylate,
diaminopimelate decarboxylase, 593
glutamate dehydrogenase, 331
Pyridine-3-suIfonamide,
bacterial growth, 504
glucose dehydrogenase, 500-502
lactate dehydrogenase, 500-502
NAD nucleosidase, 491, 504
nicotinamide deaminase, 504
Pyridine-3-sulfonate,
alcohol dehydrogenase, 504
bacterial growth, 504
glucose dehydrogenase, 500-502, 504
lactate dehydrogenase, 500-502, 504
NAD nucleosidase, 491
respiration of mycobacteria, 504
sulfite oxidase, 451
toxicity, 504
Pyridoxal,
analogs of,
active transport, 574-575
bacterial growth, 575-576
blood urea, 572-573
carcinostasis, 576-577
central nervous system, 573-574
embryogenesis, 576
enzyme inhibitions, 564-566, 569-572
fatty acid biosynthesis, 574
GABA level in brain, 573-574
pyridoxal kinase, 564-565
pyridoxine deficiency, 562, 577-578
pyridoxine levels in tissues, 566-569
pyridoxine metabolism, 564
structures of, 563
toxicity, 573-574, 577-578
tjrpes of, 562
glucose dehydrogenase, 501-502
metabolism of, pathways, 561-562
pjrridoxamine-P oxidase, 566
pyridoxol-P oxidase, 566
Pyridoxal azine, pjTidoxal kinase, 564
Pyridoxal kinase,
aminooxyacetate, 358-359
analogs, 564-565
ATP analogs, 465, 477
toxopyrimidine, 578
Pyridoxal oxime,
pyridoxamine-P oxidase, 566
pyridoxol-P oxidase, 566
PjTidoxal-phosphate, metabolic functions
of, 561
Pyridoxal semicarbazone, pyridoxal ki-
nase, 564
Pyridoxamine,
pyridoxamine-P oxidase, 566
pyridoxamine-P oxidative deamina-
tion, 564
pyridoxol-P oxidase, 566
Pyridoxamineroxalacetate transaminase,
see Transaminases
Pyridoxamine-phosphate, oxidative de-
amination of,
pyridoxamine, 564
Pyridoxamine-phosphate oxidase, ana-
logs, 566
4-Pyridoxate,
glucose dehydrogenase, 501-502
pyridoxamine-P oxidase, 566
pyridoxol-P oxidase, 566
4-Pyridoxate-phosphate,
pyridoxamine-P oxidase, 566
pyridoxol-P oxidase, 566
Pyridoxol,
oxidation of, analogs, 564
pyridoxamine-P oxidase, 566
pyridoxol-P oxidase, 566
Pyridoxol dehydrogenase, mercurials, 817
Pyridoxol-phosphate oxidase, analogs,
564, 566
Pyridoxyl - L - alanine, alanine : pyruvate
transaminase, 569
2-Pyridylalanine, L-phenylalaninersRNA
ligase (AMP), 355
4(5)-3'-Pyridylglyoxaline,
SUBJECT INDEX
1217
NAD nucleosidase, 489, 491
structure of, 488
PjTimethamine (Daraprim),
folate deficiency, 584
folinate formation, 584
uptake by bacteria, 584
Pyrimidines, metabolism of,
fluoropyrimidines, 478-481
Pyrithiaminase, 528-529
Pyrithiamine (neopyrithiamine),
bacterial growth, 528-530
bacterial resistance to, 528-529
blood pyruvate, 520, 527
central nervous system, 527, 530
fungal growth, 516, 528-529
glutamate metabolism in seeds, 522
heart rate in vivo, 527
miosis, 532
nerve membrane potentials, 531
neuromuscular block, 531-532
phosphorylation of, 519, 527
pyruvate decarboxylase in vivo, 531
pyruvate dismutation in vivo, 521
pyruvate oxidation in vivo, 520-521
rat growth, 527
respiratory quotient in rats, 521-522
structure of, 517
thiaminase, 523-524
thiamine deficiency, 516, 530-532
thiamine kinase, 522-523
thiamine levels in tissues, 525-527
thiamine-PP levels in tissues, 521
tissue levels of, 527-528
toxicity, 530-531
Vibrio growth, 530
Pyrithiamine-diphosphate,
pyruvate decarboxylase, 518
pjTuvate oxidase, 519
Pyrocatechase, mercurials, 853
PjTOgallol,
adrenergic potentiation, 611
D-amino acid oxidase, 344
blood pressure, 611
catecholamine metabolism, 611-612
catechol-0-methyl transferase, 595, 611-
612
dehydroshikimate reductase, 593
epinephrine responses, 611
histidine decarboxylase, 352
norepinephrine level in brain, 611
urinary catecholamines, 611-612
Pyrophosphatase,
o-iodosobenzoate, 712
malonate, 64
mercurials, relation to SH groups, 802
nucleotides, 475
phosphate, 439
P>Tophosphate,
glutamate semialdehyde reductase, 507
ionization constants, 242
NAD nucleosidase, 492
oxidative phosphorylation, 448
phosphatases, 439
pyridoxal kinase, 477
succinate dehydrogenase, 243
tyrosinase, 301
Pyrophosphite, oxidative phosphoryla-
tion, 448
Pyrrole-2-carboxylate, D-amino acid oxi-
dase, 342, 346
A 1 - Pyrroline - 5 - carboxylate dehydroge-
nase, analogs, 336, 355
A 1 - Pyrroline - 5 - carboxylate reductase,
analogs, 355
a-Pyrrone-5-carboxylate, see Coumalate
Pyruvate,
accumulation of, ferricyanide, 677
D-amino acid oxidase, 340
blood levels of,
malonate, 219
thiamine analogs, 520, 527
decarboxylation of,
2-deoxyglucose, 396
lipoate analogs, 590
dismutation of, pyrithiamine, 521
glycerate dehydrogenase, 430
glyoxylate reductase, 438
lactate dehydrogenase, 437
malate dehydrogenase, 596
oxidation of,
acetylene-dicarboxylate, 240-241
analogs, 429-432
benzoate, 349
trans - cyclopentane -1,2- dicarboxy-
late, 241
2-deoxyglucose, 391-392, 397, 399
1218
SUBJECT INDEX
D-glucosamine, 383
lipoate analogs, 590
malonate, 69, 74-77, 128, 135
mercurials, 878-879
oxamate, 434
oxygen, 658-659
propane-tricarboxylate, 240-241
pyrithiamine in vivo, 520-521
quinacrine, 560
thiamine analogs, 519
phosphatase (acid), 442
tartronate semialdehyde reductase, 602
Pyruvate carboxylase (pyruvateiCOa li-
ferricyanide, 676
mercurials, 853
PyruvaterCOj ligase, see Pyruvate carbo-
xylase
Pyruvate decarboxylase,
acetaldehyde, 432, 600
acetamide, 430
iodine, 682, 687
o-iodosobenzoate, 712
mercurials, 775, 783, 810, 853
mutual depletion behavior, 775
protection by pyruvate, 783
rate of inhibition, 810
oxythiamine-PP, 518
porphyrexide, 668
porphyrindin, 668
pyrithiamine in vivo, 521
pyrithiamine-PP, 518
pyruvate analogs, 600
thiamine analogs, 516, 518, 521
Pyruvate dehydrogenase, mercurials, 854
Pyruvate kinase,
6-aminonicotinamide-NAD, 505
malonate, 64, 129
mercurials, 825, 854
Pyruvate oxidase,
ferricyanide, 676
malonate, 64
mercurials, 751, 774-775, 783, 854
mutual depletion behavior, 775
protection by thiamine-PP, 783
type of inhibition, 774
mercuric ion, antagonism by hpoate,
751
oxygen inactivation of, 659
phenylpyruvate, 430
thiamine analogs, 519
Pyruvic ethyl ester, pyruvate decarboxy-
lase, 430
Quinacrine (Atabrine, Atebrin, Mepa-
crine)
bacterial growth, 546
binding to cell components, 546
competition with FAD or FMN, 546-
547, 556
distribution in tissues, 546
enzyme inhibitions, 546-559
flavin complexes with, 556
germination of spores, 546
glucose utilization, 560
nucleotide complexes with, 556
oxidative phosphorylation, 556-557
pH effects, 557-558
photophosphorylation, 557
respiration, 559-560
structure of, 546
toxicity, 546
use to detect flavin components, 559
Quinine, D-amino acid oxidase, 557
Quinine oxidase, pterin-6-aldehyde, 288
Quinolinate, NAD nucleosidase, 488
Quinolines, chymotrypsin, 373
Quinone reductase, see NADH:quinone
oxidoreductase
R
Rafiinose,
galactosidases, 418
a-glucosidase, 416
Rainbow trout, fatty acid oxidation,
malonate, 137
Reactivation, see Reversal
Relative /IF values, calculation of, 254-
255
Renilla reniformis (sea pansy), lumines-
cence,
mercurials, 891
Resistance,
to mercurials, 983-985
to thiamine analogs, 528-529
SUBJECT INDEX
1219
Resorcinol,
catechol oxidase, 296-298
respiration (endogenous) of apple skin,
296
tyrosinase, 304
Resorcinol monobenzoate, tyrosinase, 304
Respiration,
trans -aconitate, 273-274
kojic acid, 350
mercurials, 879-886, 984
quinacrine, 559-560
Respiration (acetate), see also Acetate,
oxidation of,
dehydroacetate, 626
Respiration (endogenous), see also speci-
fic inhibitors,
6-aminonicotinamide, 504
benzoate, 348
dehydroacetate, 623-624
2-deoxyglucose, 391-392, 396-397
y-hydroxy-a-ketoglutarate, 616
o-iodosobenzoate, 721
malonate, 166-186
nicotinamide, 503
p-nitrophenol, 297
resorcinol, 296
Respiration (galactose), see also Galac-
tose, oxidation of,
deoxygalactose, 391-392
Respiration (glucose), see also Glucose,
oxidation of,
l,5-anhydro-D-glucitol-6-P, 379
2-deoxyglucose, 391-394
ferricyanide, 677
hydrogen peroxide, 695
o-iodosobenzoate, 722
malonate, 123-125, 127, 133-135
mercurials, 885, 889-891, 893-894, 898,
927-928, 948
nicotinamide, 500
Respiratory quotient (R. Q.),
dependence on substrate, 184-185
malonate, 184-185
pyrithiamine, 521-522
Reticulocytes,
glucose utilization, quinacrine, 560
iron incorporation into heme, malonate,
163
leucine incorporation into protein, mer-
curials, 887
protein biosynthesis, mercurials, 887
respiration (endogenous), malonate, 179
succinate oxidation, malonate, 55
valine incorporation into protein, a-
amino-^-chlorobutyrate, 351
Retina,
glutamate uptake, malonate, 153
a-ketoglutarate oxidation, malonate, 81
K+ uptake, malonate, 153
respiration (endogenous), malonate, 176
183
Retinene oxidase, quinacrine, 554
Reversal,
of enzyme inhibitions, analysis of, 821-
823
of o-iodosobenzoate inhibitions, 718
of mercurial inhibitions, different meth-
ods for, 825
of SH reagent inhibitions, 650-651
Rhamnose, a-mannosidase, 422
Rhizobium japonicwm,
succinate dehydrogenase, malonate K^,
33
succinate oxidation, malonate, 53
Rhizoctonia solani,
growth of, mercurials, 973
sucrose uptake, mercurials, 911
Rhizopus nigricans, growth of,
dehydroacetate, 632
Rhodanese, see Thiosulfate transulfurase
Rhodopseudomonas spheroides, porphyrin
biosynthesis,
malonate, 162-163
mercurials, 888
Rhodospir ilium rubrum,
acetate oxidation, malonate, 77
fumarate oxidation, malonate, 81
hydrogen evolution, nitrogen, 293
lactate oxidation, malonate, 78
photophosphorylation,
malonate, 163
mercurials, 892
propionate oxidation, malonate, 146
respiration (endogenous), malonate, 168
succinate oxidation, malonate, 53
1220
SUBJECT INDEX
Rhodotorula gracilis, cycle substrate oxi-
dations,
malonate, 53, 74, 77, 79
Rhubarb leaves,
respiration (endogenous), malonate, 171
respiratory quotient, malonate, 185
Ribityllumazine, riboflavin transglucosi-
dase, 453
Riboflavin,
D-amino acid oxidase, 540-541
L-amino acid oxidase, 540-542
analogs of,
carcinostasis, 538
enzyme inhibitions, 540-545
FAD levels in tissues, 539-540
flavoenzymes, 540-545
growth of microorganisms, 537-538
metabolism of, 539-540
molecular complexes between, 544
quinacrine, 545-561
riboflavin metabolism, 539-540
structures of, 535-537
types of, 535
biosynthesis of, analogs, 539
FAD pyrophosphorylase, 542
galactono-y-lactone dehydrogenase,
540, 542
D-gluconate oxidase, 542
glutamate recemase, 542, 544
D-lactate oxidase, 542
L-lactate oxidase, 542-543
NADPH : methemoglobin oxidoreduc-
tase, 543
protein disulfide reductase, 543
purine complexes of, 545
succinate oxidase, 543
Riboflavin kinase, see Flavokinase
Riboflavin-5 '-sulfate,
D-amino acid oxidase, 540-541, 544
NADPH diaphorase, 544
Riboflavin synthetase, analogs, 543
Riboflavin transglucosidase,
analogs, 543
quinacrine, 554
Ribonuclease,
configurational changes, disulfide re-
duction, 650
DNA, 462
hydrogen peroxide, 693
o-iodosobenzoate, 712, 715-716
kinetics, 715-716
pH effects, 716
macroions, 461-463
mercurials, 750, 791, 793, 815, 817-
818, 820, 860
pH effects, 791, 793
reaction with substrate, 815
stimulation, 815, 817-818, 820
p-mercuribenzoate, splitting of disulfide
bonds, 750
nucleotides, 475
Ribonuclease T2, mercurials, 817
Ribonucleates, see also Nucleic acids,
biosynthesis of, mercurials, 969
deoxyribonuclease, 462
glycolysis (anaerobic), 414, 465
polycytidylate phosphorylase, 463
5'-Ribonucleotide phosphohydrolase, see
5'-Nucleotidase
Ribose,
a-mannosidase, 422
metabolism of, malonate, 132
NAD nucleosidase, 492
phosphoarabinose isomerase, 411
Ribose isomerase, mercurials, 854
Ribose-3-phosphate, phosphopentose iso-
merase, 411
Ribose-5-phosphate,
D-amino acid oxidase, 545
glucose dehydrogenase, 410
hexokinase, 379
phosphoarabinose isomerase, 411
Ribose-6-phosphate, phosphodeoxyribo-
mutase, 413
Ribose-phosphate isomerase,
cystine, 662
mercurials, 854-855
Ribulose-phosphate carboxylase, phos-
phate, 412
Ribulose-5-phosphate kinase, mercurials,
855
Rigor, see Contracture
RNA, see Ribonucleates
RNAase, see Ribonuclease
RNA nucleotidyltransferase, quinacrine,
555
SUBJECT INDEX
1221
Rose petals, respiration (endogenous),
malonate, 173, 182
Rubidium, uptake by roots,
mercurials, 909
Runner bean, see Phaseolus coccineus
Saccharo-l,4-lactone, see Glucaro-l,4-lac-
tone
Saccharomyces cerevisiae, see Yeast
Sake, malonate occurrence in, 224
Salicylamide,
glucose dehydrogenase, 501
lactate dehydrogenase, 501
sulfanilamide acetylase, 601
Salicylate,
D-amino acid oxidase, 348
dehydroshikimate reductase, 606
glucose dehydrogenase, 501
lactate dehydrogenase, 501
oxidative phosphorylation, 348
tricarboxylate cycle, 348
tyrosine:a-ketoglutarate transaminase,
306
Salicyloyl-^-alanide,
choline acetylase, 587
structure of, 587
Salmonella enteritidis, infection by,
malonate on antibacterial activity of
blood, 223-224
Salmonella paratyphi, growth of,
mercurials, 972
Salmonella pullorum,
growth of,
dehydroacetate, 632-633
mercurials, 972
infection by, malonate, 224
resistance to mercurials, 983, 983
Salmonelle schotmillleri, growth of,
mercurials, 972
Sahnonella typhosa,
growth of,
dehydroacetate, 632-633
mercurials, 972
resistance to mercurials, 983, 985
Salmonella typhimurium,
infection by, malonate, 221-223
malonate metabolism in, 228
Salyrgan, see Mersalyl
Samia cecropia, succinate dehydrogenase,
malonate, 29
Sarcina lutea, respiration (glucose)
malonate, 124
Sarcoma, ATP levels in,
aminopterin, 585
Sarcoma (mouse), growth of,
2-deoxyglucose, 400
Sarcoma (rat), growth of,
thiophene-2,5-dicarboxylate, 415
Sarcoma 37,
ADP-ATP levels in, 2-deoxyglucose,
395
blebbing of, malonate, 201
porphyrindin, 670
Sarcosine oxidase, methoxyacetate, 601
Saxostrea commercialis (oyster) eggs,
respiration (endogenous), malonate, 174
Saxostrea commercialis muscle,
respiration (endogenous), malonate, 174
succinate dehydrogenase, malonate, 28
Scallop muscle, glycolysis,
o-iodosobenzoate, 721
mercurials, 876
Scarlet fever toxin, porphyrindin inacti-
vation of, 667
Scenedesmus obliquus,
glucose uptake,
D-glucosamine, 382
a-L-sorbose-1-P, 379
photosynthesis, mercurials, 892
respiration (endogenous), mercurials,
881
Schistocera gregaria fat body,
acetate oxidation, malonate, 77
succinate oxidation, malonate, 54
Schizophyllum commune,
citrate accumulation, malonate, 104
cycle intermediate oxidations, 78-81
pyruvate oxidation, malonate, 74
respiration (glucose), mercurials, 881
succinate oxidation, malonate, 53
Sclerotinia fructicola,
growth of, mercurials, 973
resistance to mercurials, 983
Sea hare, see Aplysia
1222
SUBJECT INDEX
Sea pansy, see Renilla reniformis
Sea urchin, see also Arbacia, Echinus,
Lytechinus, Strongylocentrotus, and Trip-
neustes
Sea urchin eggs,
cleavage and development,
2-deoxyglucose, 400
o-iodosobenzoate, 726-727
malonate, 117
glycolysis, mercurials, 875
respiration (endogenous), malonate,
174-175
respiration (glucose), 2-deoxyglucose,
391
Sea urchin spermatozoa, respiration (en-
dogenous),
o-iodosobenzoate, 721-722
Sebacate, kynurenine : a - ketoglutarate
transaminase, 608-609
Sedoheptulose- 1 ,7-diphosphate, 2-keto-3-
deoxy - d - arabo - heptonate-7-P synthe-
tase, 413
Sedoheptulose-7-phosphate, 2-keto-3-de-
oxy-D-ara6o-heptonate-7-P synthetase,
413
Semicarbazide, diamine oxidase, 362
Seminal vesicle,
respiration (endogenous), malonate, 176
succinate dehydrogenase, malonate, 31
Sequential inhibition,
5-azaorotate and 6-azauridine, 480
malonate and fluoroacetate, 112
Serine,
biosynthesis of,
aminopterin, 585
2-deaminofolate, 585
deoxypyridoxol, 570-571
dipeptidase, 367
0-phosphoserine phosphatase, 270
D-Serine, L-alanine dehydrogenase, 354
L- Serine,
L-amino acid oxidase, 340
arginase, 337
phosphatase (acid), 441
L-threonine dehydrase, 357
Serine deaminase,
analogs, 357
o-iodosobenzoate, 712
L-Serine hydro-lyase, see Tryptophan syn-
thetase
Serotonin (5-hydroxytryptamine),
brain levels of, a-methyl-m-tyrosine,
316
metabolism of,
deoxypyridoxol, 574
galactoflavin, 544
tissue levels of,
a-methyldopa, 315-320
phenylalanine, 325
tryptophan pyrrolase, 324-325
Serratia marcescens,
fatty acid oxidation, malonate, 137
succinate oxidation, malonate, 52
SH groups, see Sulfhydryl groups
Shigella, succinate accumulation,
malonate, 90
Shikimate dehydrogenase, benzoate, 349
SH reagents, see Sulfhydryl reagents
Silicate, phosphatases, 439-440
Silkworm, see also Cecropia and Samia,
larvae of, aminomalonate decarboxyla-
tion in, 239
Silver
/S-glucuronidase, 795
glutamate dehydrogenase, 863
Sinapate, peroxidase, 599
Skatole,
chymotrypsin, 374
histamine release, 374
Skin,
electrical potential of,
iodine, 690
mercurials, 950
porphyrindin, 669-670
mercurial levels in, 930
mercurial vesication, 950
mitosis in, mercurials, 968
permeability to Cl~, mercurials, 912
pyruvate oxidation, 2-deoxyglucose,
391-392
respiration (endogenous),
mercurials, 882, 912
respiration (fructose), 2-deoxyglucose,
391-392
respiration (galactose), 2-deoxyglucose,
391-392
SUBJECT INDEX
1223
respiration (glucose), 2-deoxyglucose,
391-392
respiration (mannose), 2-deocyglucose,
391-392
Smooth muscle, see specific muscles
Snail, see, Helix
Sodium,
erythrocyte transport of, malonate,
209
intestinal transport of, mercurials, 916
phosphotransacetylase, 452
plasma levels of, malonate, 206
renal transport of,
dehydroacetate, 626
malonate, 206
mercurials, 918-920
urinary excretion of, malonate, 206
Solanain, iodine, 687
Sorbitol, mutarotase, 413-414
Sorbitol dehydrogenase,
o-iodosobenzoate, 712
mercurials in vivo, 926
Sorbitylflavin, flavokinase, 539
a - L - Sorbopyranose - 1 - phosphate, struc-
ture of, 377
a-L-Sorbose,
formation from glyceraldehyde, 377
hexokinase, 377
L-Sorbose-l,6-diphosphate, aldolase, 407
L-Sorbose- 1 -phosphate,
aldolase, 407
glucose uptake by Scenedesmus, 379
hexokinase, 377-379
structure of, 378
L-Sorbose-6-phosphate, hexokinase, 377
Soybean nodules,
hydrogen evolution, nitrogen, 293
nitrogen fixation, analogs, 292, 294
Spermatozoa,
acetate oxidation, malonate, 77
glycolysis,
malonate, 128
mercurials, 884
tetrathionate, 699
motility of,
malonate, 203
mercurials, 964
tetrathionate, 699
pyruvate oxidation, malonate, 77, 87
respiration (endogenous),
o-iodosobenzoate, 721-722
malonate, 176
mercurials, 882, 884
respiration (glucose), malonate, 124
Spermine, structure of, 361
Spinach chloroplasts,
COj photochemical fixation, mercurials,
892
fatty acid biosynthesis from malonate,
149
Hill reaction, mercurials, 891
malonate incorporation into, 232
NADP photoreduction, mercurials, 891
photophosphorylation, mercurials, 892
protein biosynthesis, mercurials, 887
Spinach levaes,
respiration (endogenous),
malonate, 170-171, 181
p-nitrophenol, 297
succinate accumulation, malonate, 94
Spindle formation, see also Cell division
mercurials, 969
Spiroporphyrexide, 666
Spirophorphyrindin, 666
Spleen,
amino acid accumulation, malonate, 103
ATP levels in, aminopterin, 585
citrate levels in, sequential inhibition
by malonate and fluoroacetate, 112
glycolysis, mercurials, 875
malonate decarboxylation in, 232
malonate levels in vivo, 102
mercurial levels in vivo, 930, 959-960
succinate accumulation, malonate, 102
succinate levels in vivo, malonate, 102
Spore germination, see Germination
Squalene, formation from mevalonate,
mercurials, 886
Staphylococcal phage, inactivation by
mercurials, 976, 979-980
Staphylococcus aureus,
acetate oxidation, malonate, 77
growth of,
dehydroacetate, 632-633
dichlororiboflavin analog, 537
mercurials, 972-973, 975
1224
SUBJECT INDEX
infection by, malonate, 221-222
o-iodosobenzoate killing of, 727
phosphate transport,
analog ions, 267
iodine, 690
mercurials, 910, 912-913
pyruvate oxidation, malonate, 74
resistance,
to D- cycloserine, 359
to mercurials, 983-984
to pyrithiamine, 529
respiration (glucose), mercurials, 880
Stearyl-CoA, acetyl-CoA carboxylase, 614
Stemphyllium sarcinaeforme, growth of,
mercurials, 973
Zl^-Steroid dehydrogenase, o-iodosoben-
zoate, 712
zl*-5a-Steroid dehydrogenase, o-iodoso-
benzoate, 713
Steroid hydroxylase,
o-iodosobenzoate, 713
quinacrine, 555
Sterol ester hydrolase, see Cholesterol
esterase
Sterols, see also Cholesterol,
biosynthesis of,
malonate, 149-150
mercurials, 886-887
Stomach, see Gastric acid secretion and
Gastric mucose
Strain L cells, galactose uptake,
2-deoxyglucose, 394
Streptococcus faecalis,
acetoin formation from pyruvate, phe-
nylpyruvate, 430
folate metabolism, analogs, 582
growth of, pyridoxal analogs, 575
pyruvate oxidation, mercuric ion, 751
Streptococcus hemolyticus, growth of,
mercurials, 973
Streptococcus plantarum, growth of,
dichlororiboflavin analog, 537
Streptococcus pyogenes, infection by,
malonate, 221
Streptomyces coelicolor,
a-ketoglutarate oxidation, malonate,
79, 84
succinate levels in, malonate, 97
Streptomyces olivaceus,
malonate metabolism in, 228
succinate oxidation, malonate, 53
Strigonionas oncopelti, respiration (endo-
genous),
malonate, 173
Strong ylocentrotus purpuratus eggs,
development of, malonate, 198-199
respiration (endogenous), malonate,
175-176, 181
Stylonychia pusulata, succinate dehydro-
genase,
malonate, 28
Suberate,
aspartate : a - ketoglutarate transami-
nase, 334
glutamate decarboxylase, 328
kynurenine: a-ketoglutarate transami-
nase, 608
Substitution analog, definition of, 257
Succinate,
accumulation of, malonate, 70-71, 90-
104
aspartate : a - ketoglutarate transami-
nase, 334
citrate sjnithetase, 597
fumarase, 275, 277
glutamate decarboxylase, 328
glutamate dehydrogenase, 330, 332
intercharge distance, 5-6
ionic species, pH effect, 10
ionization of, 8
kynurenine:a-ketoglutarate transami-
nase, 608
D -lactate dehydrogenase, 437
oxidation of,
benzoate, 348
malonate, 50-58
mercurials, 879
phosphofructokinase, 385
urinary excretion of, dehydroacetate,
628
Succinate dehydrogenase,
acetylene-dicarboxylate, 240-241
active center of, 18, 40-42, 44-45
arsonoacetate, 243
1,4-butane-disphophonate, 243
caffeate, 314
SUBJECT INDEX
1225
<raw5-cyclopentane-l,2-dicarboxylate,
241
cysteine, 659
dehydroacetate, 620-622
dicarboxylate ions, 34-40
difluoromalonate, 239
ethane- 1,2-disulfonate, 242-243
ferricyanide, 676
fluoromalonate, 239
GSSG, 661-662
hydrogen peroxide, 693
3-hydroxycinnamate, 314
hypophosphate, 243
intercharge distance in active site, 42-
44
iodine, 682, 687
o-iodosobenzoate, 713, 715-718
itaconate, 601
malonate,
absorption spectrum changes, 18-19
activation of enzyme, 45-46
binding energy, 42
competitive nature, 21-25
dependence on electron acceptor, 18-
20
discovery of inhibition, 2
effect of ATP, 48
effect of Ca++, 46-48
effect of liquid nitrogen, 187
effect of Mg++, 48
effect of osmolarity, 46-47
effect of succinate concentration, 25
effect of temperature, 46
interaction distance, 42-43
KJKi ratio, 33
reversibility, 24-25
species variation, 49-50
malondialdehyde, 40-41
malondiamide, 41
malonic diethyl ester, 236
mercurials, 773, 779, 783-784, 787-788,
802, 810, 825, 855-856, 870-872, 925-926
acceleration of nonheme Fe chela-
tion, 788
in vivo inhibition, 925-926
loss of nonheme Fe, 787
protection by XAD, 783
protection by oxalacetate, 779
protection by succinate, 783-784
rate of inhibition, 810
relation to SH groups, 802
reversal by different methods, 825
type of inhibition, 773
methane-diphosphonate, 243
methionate, 243
methods for measuring activity of, 18-
20
3-nitropropionate, 244
oxygen inactivation of, 659
phosphonoacetate, 243
^-phosphonopropionate, 242-243
porphyrindin, 668
propane-tricarboxylate, 240-241
pyrophosphate, 243
quinacrine, 555
succinyl peroxide, 694
o-sulfobenzoate, 243
/S-sulfopropionate, 242-243
tetrathionate, 696, 700
Succinate: methylmalonate isomerase, ma-
lonate, 64
Succinate oxidase,
composition of, 16-17
dehydroacetate, 620-622
FAD, 543
fiavinmononucleotide, 543
y-hydroxy-a-ketoglutarate, 616
o-iodosobenzoate, 715-718
kinetics, 715
protection by succinate, 717
reversal by thiols, 718
temperature effects, 715
kojic acid, 350
malonate, localization of site of action,
18, 20-21
mercurials, 817, 826, 870-872
A"-piperidinomethylnicotinamide, 503
quinacrine, 547, 555, 560
riboflavin, 543
riboflavin analogs, 543
Succinate semialdehyde dehydrogenase,
j3-hydroxybenzaldehyde, 601
mercurials, 856
Succinyl-CoA deacylase, ATP, 475
Succinyl-/S-ketoacyl-CoA transferase,
transfer of Co A to malonate, 234
1226
SUBJECT INDEX
Succinyl peroxide, enzyme inhibitions by,
694
Sucrose,
a-amylase, 420
/S-amylase, 421
a-dextran-l,6-glucosidase, 417
a-glucosidase, 416-417, 423
a-mannosidase, 422
uptake by Fhizoctonia, mercurials, 911
Sucrose transfructosylase, analogs, 421
Sugar cane, glucose uptake,
mercurials, 910-911
Sugar pine seedlings, see also Pinus lam-
bertiana,
tricarboxylate cycle oxidations, ma-
lonate, 74, 80, 82
Sulfamate, phosphatase (acid), 441
Sulfanilamide, acetylation of,
mercurials, 927
Sulfanilamide acetylase,
analogs, 601
mercurials, 750
Sulfanilate, D-amino acid oxidase, 343
Sulfatases,
analogs, 443-444
mercurials, 860
Sulfate,
aldolase, 414
arylsulfatases, 443-444
choline sulfatase, 444
creatine kinase, 446
glycolysis (anaerobic), 414
thiosulfate utilization by Neurospora,
451-452
tyrosinase, 301
uptake by yeast, thiosulfate, 267
Sulfated hyaluronate, hyaluronidase, 459
Sulfated pectate, hyaluronidase, 459
Sulfated polysaccharides,
lipoprotein lipase, 463
ribonuclease, 462-463
Sulfenyl iodine groups, formation by io-
dine, 680-681
Sulfhydryl enzymes,
definition of, 635, 647
oxygen inactivation of, 659
reactivity of SH groups, factors deter-
mining, 643-647
Sulfhydryl groups,
addition to double bonds, 643
alkylation of, 642
bond characteristics, 639
chemical properties, 637-642
determination of, 640-642, 667, 669,
671-672, 680-681, 697-698, 702-704,
752-757, 762-768, 798-809
hydrogen bonding, 640
ionization of, 638
oxidation of, 642, 655-700
oxidation-reduction potentials, 656
reactivity of, factors determining, 643-
647
role in cellular function, 637
role in metabolism, 636-637
Sulfhydryl reagents, 635-653
configurational changes in enzymes,
649-650
enzyme inhibitions,
interpretation of, 647-650
mechanisms of, 647-648
protection against, 650-651
reversal of, 650-651
reaction with disulfides, 640
reaction with SH groups, pH effects,
638
specificity of, 652-653
types of reactions with enzymes, 642-
643
uses of, general considerations, 651-
653
Sulfite,
arylsulfatase, 443-444
choline sulfatase, 444
oxidation of, thiosulfate, 451
Sulfite oxidase,
analogs, 451
quinacrine, 555
Sulfoacetate, aspartase, 355
o-Sulfobenzoate,
intercharge distance, 7
succinate dehydrogenase, 243
Sulfocholine, thetin:homocysteine trans-
methylase, 356
Sulfoglycolate,
glycolate oxidase, 438
structure of, 438
SUBJECT INDEX
1227
Sulfonamides, see also Sulfanilamide,
acetylation of, folate analogs, 586-587
/9-Sulfopropionate,
intercharge distance, 7
succinate dehydrogenase, 242-243
Sunflower stems,
auxin transport, mercurials, 967
growth of, malonate, 197
respiration (endogenous), mercurials,
881
Sweet potato, see also Ipomea,
citrate oxidation, mercurials, 878
oxidative phosphorylation, mercurials,
873
Sympathetic ganglia, transmission,
malonate, 211
Taka-/?-glucosidase, phenol-a-glucoside,
271
Talose, fructokinase, 376
Tapeworms, see Echinococcus and Hyme-
nolepis
Tartrate,
fumarase, 275, 277-279
ionization constants, 8
lactate dehydrogenase, 437
permeability of erythrocytes to, 188
phosphatases, 440-442
D-Tartrate,
L-tartrate dehydrase, 601
meso-tartrate dehydrase, 601
L-tartrate, D-tartrate dehydrase, 601
meso-Tartrate,
a-ketoglutarate oxidation, 432
malate dehydrogenase, 596
phosphatases, 440-441
pyruvate oxidation, 432
D-Tartrate dehydrase, analogs, 601
L-Tartrate dehydrase, analogs, 601
meso-Tartrate dehydrase, analogs, 601
Tartronate (hydroxy malonate),
glycolysis, 238
D-a-hydroxy acid dehydrogenase, 437
intercharge distance, 6
ionization constants, 8
ketogenic activity, 237
lactate dehydrogenase, 237-238, 436-437
malate dehydrogenase, 237, 596
malate dehydrogenase (decarboxylat-
ing), 238, 597
occurrence in tissues, 225, 238
phosphatases, 238
respiration, 238
succinate dehydrogenase, 36, 40, 237
D-tartrate dehydrase, 601
weso-tartrate dehydrase, 601
tartronate semialdehyde reductase, 602
Tartronate semialdehyde reductase, ana-
logs, 602
Terephthalate,
D-amino acid oxidase, 341, 344
aspartate : a - ketoglutarate transami-
nase, 334
glutamate dehydrogenase, 331
intercharge distance, 6
kynurenine: a-ketoglutarate transami-
nase, 607-608
succinate dehydrogenase, 37
tyrosinase, 300
Testis,
citrate formation in, malonate, 105, 108
malonate decarboxylation in, 232
respiration (endogenous), malonate, 175
transaminases in vivo, deoxypyridoxol,
570
Tetradecane-l,14-dicarboxylare, kynure-
nine:a-ketoglutarate transaminase, 608
Tetraethylammonium ion, renal transoprt
of,
dehydroacetate, 626
malonate, 204
mercurials, 921
Tetrahydrofolate, biosynthesis of,
analogs, 581-584
Tetrahymena geleii,
acetate utilization, <raw5-cyclopentane-
1,2-dicarboxylate, 241
succinate dehydrogenase, irans-cyclo-
pentane-l,2-dicarboxylate, 37
Tetrahymena pyriformis, glycolysis (an-
aerobic),
oxamate, 434
Tetraiodothyroacetate, thyroxine deiodi-
nase, 602
1228
SUBJECT INDEX
3,5,3',5'-Tetraiodothyropropionate, thy-
roxine deiodinase, 602
Tetrathionate, 696-700,
blood GSH, 696
chemical properties, 697
conversion to thiosulfate, 699-700
cyanide poisoning, 696
cytochrome reduction by, 696
determination of protein methionine,
696
enzyme inhibitions, 698-699
GSH levels in tissues, 700
nephrotoxicity, 700
oxidation of protein SH groups, 697-
698
oxidation of thiols, 697-698
oxidation-reduction potential, 697
reaction with SH groups, 697
thromboangiitis obliterans, 696
toxicity, 700
Tetrolate, fatty acid biosynthesis, 614
Tetrolyl-CoA, fatty acid biosynthesis,
613-614
Tetrose-diphosphate, see D-Threose-2,4-
diphosphate
Thea pollen, malonate metabolism in, 228
Theobromine, NAD nucleosidase, 492
Theophylline, NAD nucleosidase, 492
Thetin:homocysteine transmethylase, a-
nalogs, 356-357
Thiaminase, analogs, 523-525
Thiamine,
analogs of, see also Oxythiamine, Pyri-
thiamine, and others, 514-534,
central nervous system, 527, 530-531
enzyme inhibitions, 518-519
growth of microorganisms, 528-530
mechanisms of action, 532-534
metabolic disturbances by, 533-534
neuromuscular function, 531
phosphorylation of, 519
pyruvate accumulation, 520-521
resistance to, 528-529
sites of action, 516
structures of, 517
summary of actions, 532
thiaminase, 523-525
thiamine deficiency, 530-531
thiamine kinase, 522-523
thiamine levels in tissues, 525-526
thiamine-PP levels in tissues, 525-527
tissue levels of, 527-528
toxicity, 530-531
types of, 516-518
metabolism of, pathways, 514-515
reaction with mercurials, 774
urinary excretion of, oxythiamine, 525
Thiamine disphosphatase, mercurials, 860
Thiamine disulfide,
structure of, 517
thiamine kinase, 523
Thiamine-diphosphate,
metabolic functions of, 514-516
tissue levels of, analogs, 525-527
Thiamine kinase,
analogs, 522-523
role of inhibition in toxicity of, 532-
533
nucleotides, 475
Thiazole-pyrophosphate, see also 4-Me-
thyl-5-hydroxyethylthiazole-PP, struc-
ture of, 517
Thiazolidine rings, reactivity of enzyme
SH groups, 644, 646
Thiazoline rings, reactivity of enzyme SH
groups, 644, 646
/3-2-Thienylalanine,
phenylalanine hydroxylase, 354
L-phenylalanine:sRNA ligase (AMP),
354
Thienylglycine, glycine uptake by ascites
cells, 265
Thimerosal (Merthiolate), structure of,
970
Thiobacillus thiooxidans,
CO2 fixation, mercurials, 892
respiration (endogenous), malonate, 168
Thioesters, splitting by mercurials, 751
Thioglycolate, glutathione oxidase, 593
Thioglycosidase, glucono-l,4-lactone, 429
6-Thioinosinemonophosphate, IMP dehy-
drogenase, 471, 481
Thiol-disulfide equilibria, 639, 656
Thiols, see also Sulfhydryl groups,
equilibria with disulfides, 639, 656
role in metabolism, 636-637
SUBJECT INDEX
1229
Thiomerin, see Mercurin
Thionicotinamide-NAD,
alcohol dehydrogenase, 497
lactate dehydrogenase, 497
Thiophene-2,5-dicarboxylate,
carcinostasis, 415
glucose metabolism in tumors, 415
Thiophosphate, oxidative phosphoryla-
tion, 447-448
Thiosulfate,
formation from tetrathionate, 699-700
sulfate uptake by yeast, 267
sulfite oxidase, 451
utilization by Neurospora, sulfate, 451-
452
Thiosulfate reductase, quinacrine, 555
Thiosulfate transulfurase,
o-iodosobenzoate, 713
mercurials, relation to SH groups, 803
Thiothiamine, thiamine kinase, 523
Thiourea, urease, 603, 610
L-Threonine,
L-amino acid oxidase, 340
aspartokinase, 356
homoserine kinase, 357
Threonine aldolase, o-iodosobenzoate, 713
718
Threonine dehydrase,
mercurials, coenzyme displacement, 787
serine, 357
Threonine dehydrogenase,
o-iodosobenzoate, 713
mercurials, 856
Threonine synthetase, analogs, 357
D-Threose-2,4-diphosphate,
glycolysis, 409
3 - phosphoglyceraldehyde dehydroge-
nase, 408-409
photosynthesis, 409
Thrombin, tosylagmatine, 375
Thromboangiitis obliterans, use of tetra-
thionate in, 696
Thymidine,
aspartate carbamyltransferase, 468
5'-nucleotidase, 472
Thymidinemonophosphate (TMP),
aspartate carbamyltransferase, 468
phosphodiesterase, 473
Thymidylate kinase, deoxy-GMP, 475
Thymidylate synthetase, analogs, 476,
479
Thymine, NAD nucleosidase, 492-493
Thymocytes, glycolysis,
malonate, 126
Thymus,
amino acid accumulation, malonate,
103
citrate levels in, sequential inhibition
by malonate and fluoroacetate, 112
malonate levels in vivo, 102
succinate dehydrogenase, 32
succinate levels in vivo, malonate, 102
Thymus nuclei,
ATP levels in, malonate, 189
glucose metabolism,
dehydroacetate, 624
2-deoxyglucose, 393-394
respiration, malonate, 189
respiration (glucose), dehydroacetate,
624
Thyone muscle, glycolysis,
mercurials, 876
Thyroid,
glucose metabolism, malonate, 131
glycolysis (aerobic), malonate, 128
iodide uptake,
malonate, 209
mercurials, 910
pyruvate oxidation, malonate, 76
respiration (endogenous), malonate 176
respiration (glucose), mercurials, 883
Thyronine, thyroxine deiodinase, 603
D-Thyroxine, L-thyroxine deiodinase, 602
Thyroxine deiodinase,
analogs, 602-603
mercurials, 778
quinacrine, 555
Tibial condyles, chondroitin sulfate bio-
synthesis,
malonate, 166
Titration of enzyme SH groups,
o-iodosobenzoate, 714-715
mercurials, 766, 798-809
porphyrindin, 667
tetrathionate, 699
Titration of protein SH groups.
1230
SUBJECT INDEX
ferricyanide, 672
iodine, 680-681
o-iodosobenzoate, 703-704
mercurials, 762-766
tetrathionate, 697
TMP, see Thymidinemonophosphate
Toadfish, see Opsanus tau
Tobacco,
citrate formation, malonate, 105
citrate oxidation, malonate, 79, 87
cycle intermediate levels in, malonate,
107, 111
glycolate levels in, a-hydroxysylforates,
439
growth of stem cultures, malonate, 197
a-ketoglutarate oxidation, malonate, 80
malate oxidation, malonate, 82
malonate,
metabolism in, 228
occurrence in, 225
pH effects, 190
NADP reduction by hexose-P's, mer-
curials, 885
pentose-P pathway, mercurials, 885
protein biosynthesis, 107, 156
respiration (endogenous), malonate, 172
succinate accumulation, malonate, 91
succinate dehydrogenase, 28
Tobacco mosaic virus,
inactivation by mercurials, 976, 979-
980
infectivity of, mercuric ion, 741
proliferation of,
malonate, 194
mercurials, 976, 979
titration of SH groups,
iodine, 681
porphyrindin, 667
Toluates, see Methylbenzoates
j)-Toluenesulfonate, D-amino acid oxidase,
343
p-Tolylalanine, L-phenylalanine:sRNA li-
gase (AMP), 355
m-Tolyl-sulfate, arylsulfatase, 443
Tomato,
pyruvate oxidation, malonate, 74
respiration (endogenous), malonate, 171
succinate dehydrogenase, malonate, 27
Tosylagmatine,
blood clotting, 375
thrombin, 375
trypsin, 375
Toxicity, see also Lethal doses,
3-acetylpyridine, 489, 494
alkylmalonates, 2
6-deoxy-6-fluoroglucose, 404-405
2-deoxyglucose, 401
fluoromalonate, 239
hydrogen peroxide, 696
o-iodosobenzoate, 725
kojic acid, 349
malonate, 1-2, 217-221
mercurials, 924-925, 950-957
oxamate, 434
pyridoxal analogs, 562, 573-574, 577-
578
quinacrine, 546
tetrathionate, 700
thiamine analogs, 530-531
Toxoflavin,
bacterial growth, 538
structure of, 537
Toxoplasma gondii,
infection by, deoxypyridoxol, 576
respiration (glucose), mercurials, 882
Toxopyrimidine,
central nervous system, 578
enzyme inhibitions, 578
glutamate decarboxylase in brain, 571
structure of, 563
toxicity, 562, 564
Toxopyrimidine-phosphate, tyrosine de-
carboxylase, 578
TPN, see NADP
TpTpTpT, phosphodiesterase, 473
Transaldolase, phosphate, 412
Transaminases,
alanine: a-ketoglutarate,
aminoxyacetate, 358
analogs, 334
L-cycloserine, 360
deoxypyridoxol in vivo, 569
malonate, 64
mercurials, 856-857
alanine:pyruvate, deoxypyridoxol, 569
y-aminobutyrate: a-ketoglutarate,
SUBJECT INDEX
1231
aminooxyacetate, 358-359
D-cycloserine, 359
malonate, 64
mercurials, 857
6- amino valerate : a -ketoglutarate, mer-
curials, 857
asparagine : a - ketoglutarate,
cycloserines, 360
asparagine: pyruvate, mercurials, 857
aspartate:a-ketoglutarate,
analogs, 334, 355
ferricyanide, 675
malonate, 64
mercurials, 808, 827, 857
glutamine:pyruvate,
GSSG, 662
mercurials, 857
glycine: a-ketoglutarate, mercurials, 857
kynurenine: a-ketoglutarate,
acetate, 608
analogs, 595, 607-610
malonate, 64
pyridoxal analogs, 569-570
pyridoxamine-oxalacetate, oxalacetate
analogs, 600
toxopyrimidine, 578
tryptophan:a-ketoglutarate, mercurials
857
tyrosine:a-ketoglutarate,
analogs, 305-306
o-iodosobenzoate, 713
mercurials, 857
Transfer RNA, biosynthesis from nucleo-
tides,
mercurials, 820
Transhydrogenase, see NAD:NADP trans-
hydrogenase
Transketolase,
mercurials, 857
oxythiamine in vivo, 522
phosphate, 412
thiamine analogs, 519, 522
Transmembrane potentials, see Membrane
potentials
Transphosporylases, analogs, 444-447
Trehalose, a-glucosidase, 416
Treponema pallidum, resistance to mer-
curials, 983-985
2, 4, 7 - Triamino - 6 - o - methylphenylpteri-
dine, folinate formation, 582
Tribromophenol, D-amino acid oxidase,
344
Tricarboxylate cycle,
ferrocyanide, 677-678
y-hydroxy-a-ketoglutarate, 615-616
intermediates of, concentrations in
cells, 88-90
limiting reactions in, 70
malonate, 69-90
mercurials, 877-879
Trichloroacetate,
pantoate:/3-alanine ligase, 598
tyrosinase, 300
a,a,/3-Trichloropropionate, pantoate:/5-a-
lanine ligase, 598
2,6,8-Trichloropurine, uricase, 284
Trichomonas foetus, respiration (endoge-
nous),
malonate, 173
Trichomonas suis, respiration (endoge-
nous),
malonate, 173
Trichomonas vaginalis, respiration (glu-
cose),
mercurials, 882
Trichophyton interdigitale, growth of,
dehydroacetate, 632
Trichophyton mentagrophytes, growth of,
dehydroacetate, 632
Trichophyton rubrum, respiration (endo-
genous),
malonate, 169
mercurials, 880
Trichophyton schoenleini, respiration (en-
dogenous),
malonate, 169
Triethylsulfonium ion, thetin: homocys-
teine transmethylase, 356
Trifluoroacetyltryptamine, chymotrypsin,
371
Trifluoroacetyltryptophanaraide, chymo-
trypsin, 371
Trifluoroacetyltyrosinamide, chymotryp-
sin, 371
Trifluorothiamine,
Bacillus growth, 531
1232
SUBJECT INDEX
thiamine deficiency, 531
tumor growth, 531
Trigonelhne,
glucose dehydrogenase, 501-502
lactate dehydrogenase, 501-502
NAD nucleosidase, 488, 491
structure of, 488
Trihydrobenzoates, dopa decarboxylase,
312
Triiodophenol, D-amino acid oxidase, 344
Triiodothyroacetate, intestinal transport
of,
mercurials, 911, 913-914
Triiodothyronines, thyronine deiodinase,
602-603
Trimesate,
glutamate dehydrogenase, 330
structure of, 330
Trimetaphosphimate, oxidative phospho-
rylation, 448
Trimethylacetate, tyrosinase, 300
Trimethylammonium ion, thetin:homo-
cysteine transmethylase, 356
Trimethylenediamine, diamine oxidase,
362
1,3,7-Trimethylurate, uricase, 283-284
1,3,9-Trimethylurate, uricase, 283-284
2,4,6-Trinitrophenol, D-amino acid oxi-
dase, 348
Triose-phosphate dehydrogenase, see 3-
Phosphoglyceraldehyde dehydrogenase
Triose-phosphate isomerase, phosphate,
412
Tri peptidase, o-iodosobenzoate, 713
Triphosphate, oxidative phosphorylation,
448
Triphosphoinositide phosphodiesterase,
mercurials, 858
Triphosphoinositide phosphomonoester-
ase, mercurials, 858
Tripneustes esclulentus, development of,
mercurials, 964
Tripolyphosphate,
creatine kinase, 446
hexokinase, 383
yeast fermentation, 383
True inhibitor constant, definition of, 252
Trypanosoma cruzi,
glucose utilization, malonate, 127
succinate accumulation, malonate, 91,
127
succinate dehydrogenase, malonate, 28
Trypanosoma hippicum, respiration (endo-
genous),
malonate, 173
Trypanosomes,
growth of, quinacrine, 559
respiration, quinacrine, 559
Trypsin,
dehydroacetate, 622
inactivation of phosphoribosyl-ATP py-
rophosphorylase, 351
macroions, 456-457
mercurials, 797-798
tosylagmatine, 375
Tryptamine,
chymotrypsin, 371, 374
dopa decarboxylase, 308
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
Tryptazan,
incorporation into enzymes, 326
maltase biosynthesis, 326
tryptophan pyrrolase, 325
L-tryptophan:sRNA hgase (AMP), 326
Tryptophan,
biosynthesis of, analogs, 321, 323
histidine decarboxylase, 352-353
metabolism of,
analogs, 321-326
deoxypyridoxol, 572
pathways of, 322
D -Tryptophan,
chymotrypsin, 374
tryptophan pyrrolase, 325
L-tryptophan:sRNA ligase (AMP), 326
L-Tryptophan,
arginase, 337
chymotrypsin, 374
dipeptidase, 367
feedback inhibition of anthranilate
synthesis, 321
intestinal transport of iodotyrosine, 265
tyrosine:a-ketoglutarate transaminase,
306
uptake by ascites cells, analogs, 265-266
SUBJECT INDEX
1233
Tryptophanamides, chymotrypsin, 271,
371
L-Tryptophanase,
analogs, 323-324
mercurials, 858
methylindoles, 321
toxopyrimidine, 578
D-tryptophan, 268
Tryptophan hydroxylase (phenylalanine
hydroxylase), analogs, 325-326
Tryptophan : a - ketoglutarate transami-
nase, see Transaminases
Tryptophan peroxidase, see Tryptophan
pyrrolase
Tryptophan pyrrolase (tryptophan pero-
xidase),
hematin analogs, 603
induction of, /?-azaguanine, 478
mercurials, 817
tryptophan analogs, 324-325
L-Tryptophan : sRNA ligase (AMP), ana-
logs, 326
Tryptophan synthetase,
analogs, 321
ferricyanide, 676
Tumor 2146 (mouse), growth of,
2-acetamido-2-deoxygluconolactone,
428
glucaro-l,4-lactone, 428
Tumors, see also specific tumors,
blebbing of, see Sarcoma 37
DNA levels in, 2-deoxyglucose, 399
fatty acid biosynthesis, malonate, 148-
149
glucose metabolism, thiophene-2,5- di-
carboxylate, 415
glycolysis,
2-deoxyglucose, 392
hydrogen peroxide, 695
glycolysis (anaerobic), D-glucosone, 385
growth of,
6-azauracil, 478
2-deoxyglucose, 400-401
malonate, 200-202
pyridoxal analogs, 576-577
riboflavin analogs, 538
trifluoro thiamine, 531
mercurial uptake in vivo, 969-970
metabolism in, ferricyanide, 677
protein biosynthesis, malonate, 156
respiration (endogenous), trans-aconi-
tate, 273
transaminases in vivo, deoxypyridoxol,
570
Tungstate,
Agrohacterium growth, 615
Aspergillus growth, 614
Azotobacter growth, 614
molybdate uptake, 614
molybdenum deficiency, 614-615
molybdenum levels in tissues, 614
nitrate reductase, 615
xanthine oxidase in vivo, 614
Turanose, a-glucosidase, 416, 423
Turnip yellow mosaic virus, splitting into
subunits by mercurials, 980
Tyramine,
dopa decarboxylase, 308
dopamine /^-hydroxylase, 320
oxidation of, kojic acid, 350
tyrosine:a-ketoglutarate transaminase,
306
L-tyrosine:sRNA ligase (AMP), 307
D-Tyrosinamide, chymotrypsin, 271
L-Tyrosinamide,
cathepsin C, 375
L-tyrosine:sRNA ligase (AMP), 307
Tyrosinase,
acetate, 300-301
analogs, 300-305
benzoate, 349
cafFeate, 314
fluoride, 300
maleate, 301
mercurials, 794
oxalate, 300
pyrophosphate, 301
succinyl peroxide, 694
D- and L-tyrosines, 268
)3-Tyrosinase, ferricyanide, 676
Tyrosine,
histidine decarboxylase, 352-353
metabolism of,
analogs, 302-320
pathways of, 303
phenylalanine deaminase, 355
1234
SUBJECT INDEX
D-Tyrosine, L-tyrosine:a-ketoglutarate
transaminase, 306
Tyrosine decarboxylase,
analogs, 306-307
cafFeate, 314
folate analogs, 586
permanganate, 660
toxopyrimidine, 578
toxopyrimidine-P, 578
Tyrosinehydroxamide, chymotrypsin, 371
Tyrosine:a-ketoglutarate transaminase,
see Transaminases
L-Tyrosine:sRNA ligase (AMP), analogs,
307
u
UDP, see Uridinediphosphate
UDPgalactose-4-epimerase, mercurials,
858
UDPglucose dehydrogenase,
GSSG, 662
o-iodosobenzoate, 713
mercurials, 858-859
UDPglucose:a-l,4-glucan-a-4-glucosyl-
transferase,
2-deoxyglucose-6-P, 391
UDP and UMP, 476
UDPglucose - glycogen glucosyltransfer-
ase, D-glucosamine, 382
UDPglucose pyrophosphorylase,
galactose- 1-P, 603
D-glucosamine, 382
UDPglucose-starch glucosyltransferase,
see UDPglucose:a-l,4-glucan-a-4-gIuco-
syltransferase
UDP glucuronyltransferase,
o-iodosobenzoate, 713
mercurials, 817, 859
Ulcers, treatment with pepsin inhibitors,
458
Ulothrix zonata, succinate dehydrogenase,
malonate, 27
UMP, see Uridinemonophosphate
Undecane -1,11- dicarboxylate, kynuren-
ine:a-ketoglutarate transaminase, 608
Uracil,
D-amino acid oxidase, 545
intestinal transport of, mercurials, 911
NAD nucleosidase, 493
5'-nucleotidase, 472
6-Uracilmethylsulfone, orotate transphos-
phoribosylase, 473
6-Uracilsulfonamide, orotate transphos-
phoribosylase, 473
6-Uracilsulfonate, orotate transphospho-
ribosylase, 473
Urate,
diamine oxidase, 365
oxidation of, kojic acid, 350
uptake by erythrocytes, hypoxanthine,
267
Urate oxidase, see Uricase
Urate riboside, inosine hydrolase, 471
Urea,
formation of,
deoxypyridoxol, 572-573
malonate, 157-158
Urea cycle, deoxypyridoxol, 572-573
Urease,
acetamide, 603
analogs, 603, 610
cystine, 662
dehydroacetate, 621-622
ferricyanide, 673, 676
iodine, 682-683, 687
malonate, 64
mercurials, 773, 778, 794, 859
pH effects, 794
protection by ascorbate, 778
type of inhibition, 773
methylurea, 610
phenylisocyanate, 649
porphyrindin, 669
SH groups of, titration of, 643
succinyl peroxide, 694
Urechis caupo eggs, development of,
ferricyanide, 678
Urechis unicinctus eggs, development of,
o-iodosobenzoate, 727
y-(3,4-Ureylenecyclohexyl)butyrate,
structure of, 588
yeast fermentation, 588-589
Uricase,
hydrogen peroxide, 691, 693
o-iodosobenzoate, 713
mercurials, 859
SUBJECT INDEX
1235
pterin-6-aldehyde, 288
purine analogs, 283-286
Uridine,
aspartate carbamyltransferase, 468
5 '-nucleotidase, 472
Uridinediphosphate (UDP),
fructose- 1,6-diphosphatase, 470
orotidylate decarboxylase, 473, 479
UDPglucose:a-l,4-glucan-a-4-glucosyl-
transferase, 476
Uridinemonophosphate (UMP),
adenylosuccinate synthetase, 467
aspartate carbamyltransferase, 468
deoxycytidylate deaminase, 469
5 -nucleotidase, 471
orotidylate decarboxylase, 473, 479
phosphatase, 439
pyrophosphatase, 475
UDPglucose:a-l,4-glucan-a-4-glucosyl-
transferase, 476
Uridinetriphosphatase (UTPase),
ADP, 446
IDP, 446
Uridinetriphosphate (UTP),
aspartate carbamyltransferase, 468
isocitrate dehydrogenase, 509
NADH oxidase, 511
orotidylate decarboxylase, 473, 477
Urine,
ethylmalonate in, 225
malonate in, 225-226
methylmalonate in, 224
tartronate in, 238
Urinary flow,
dehydroacetate, 625
malonate, 206
mercurials, 917-918
Urinary pH, malonate, 206
Urocanase,
hydrogen peroxide, 693
o-iodosobenzoate, 713
mercurials, protection by urocanate, 783
permanganate, 660
Urocanate oxidase, malonate, 64
Uroporphyrinogen decarobxylase, mer-
curials, 859
Ustilago maydis, respiration (glucose),
malonate, 133-134
Uterus,
contracture, o-iodosobenzoate, 724
motility, o-iodosobenzoate, 724
UTP, see Uridinetriphosphate
UTPase, see Uridinetriphosphatase
Vaccinia virus,
infectivity of, mercurials, 979-980
proliferation of, malonate, 193
Valerate (pentanoate),
carboxypeptidase, 366
glutamate decarboxylase, 328
kynurenine:a-ketoglutarate transami-
nase, 608-609
lactate dehydrogenase, 436
leucine decarboxylase, 352
f5-Valerolactam, A i-pyrroline-5-carboxy-
late dehydrogenase, 336
Valine,
alanine: ct-ketoglutarate transaminase,
334
dipeptidase, 367
incorporation into proteins, a-amino-^-
chlorobutyrate, 351
D-Valine, L-alanine dehydrogenase, 354
L-Valine, L-amino acid oxidase, 340
Valylleucine, penicillinase, 599
Valylvaline, penicillinase, 599
Vanillin, dehydroshikimate reductase,
593, 604-605
Vascular smooth muscle, malonate, 212
Ventricle, see Heart
Venturia inaegualis, ascosporulation-
malonate, 195
Veratroyl-^-glucoronide, glucuronidases,
426-427
Vessels, see Vascular smooth muscle
Vetch leaves, malonate occurrence in, 224
Vibrio cholera,
amino acid accumulation, deoxypyri-
doxol, 576
growth of,
dehydroacetate, 632
deoxypyridoxol, 576
thiamine analogs, 522, 530
Vibrio metchnikowii, growth of,
dehydroacetate, 532
1236
SUBJECT INDEX
Vigna sinensis, glutamate metabolism,
malonate, 153
Viruses, see also specific viruses,
proliferation of,
o-iodosobenzoate, 728
malonate, 192-194
mercurials, 976-981
Vitamin B,, see FAD and Riboflavin
Vitamin B^, see Thiamine
Vitamin Bg, see Pyridoxal and derivatives
Vitamin Bjj, see Cyanocobalamin
Vitamin Kj reductase, o-iodosoenzoate,
713
W
Walker carcinosarcoma,
2-deoxyglucose, 386
glutamate metabolism, malonate, 152
glycolysis, ferricyanide, 677
malonate levels in vivo, 102
protein biosynthesis, malonate, 156
pyruvate oxidation, mercurials, 878
succinate levels in vivo, malonate, 102
Water,
corneal transport of, mercurials, 91 1
intestinal transport of, mercurials, 916
renal transport of, mercurials, 917-918
Western equine encephalitis virus, in-
fectivity of,
mercurials, 979
Wheat,
malonate occurrence in, 224-225
respiration (endogenous), malonate, 170
184
Woodroach, see Leucophaea
Xanthine,
NAD nucleosidase, 492
oxidation in liver, kojic acid, 350
uricase, 285
Xanthine:cytochrome c oxidoreductase,
myoglobin, 603
Xanthine dehydrogenase, mercurials, 859
Xanthine oxidase,
analogs, 279-289
8-azaguanine, 477
hydrogen peroxide, 693-694
9-hydroxyethylriboflavin analog, 544
iodine, 687
o-iodosobenzoate, 669, 704, 713, 718
mercurials, 783, 796-797, 803, 807, 814
pH eifects, 796-797
potentiation of inhibition by sub-
strate, 807
protection by hypoxanthine, 783
relation to SH groups, 803
spontaneous reversal, 814
oxygen inactivation of, 659
porphyrindin, 668-669
pteridine analogs, 285-289
purine analogs, 280-283
quinacrine, 555
tungstate in vivo, 614
Xanthomonas phaseoli, succinate dehydro-
genase,
malonate, 19, 26
Xanthopterin,
guanase, 288
structure of, 287
xanthine oxidase, 289
Xanthopterin-7-carboxylate, xanthine o-
xidase, 289
Xanthosine, inosine hydrolase, 471
Xanthosine-5'-phosphate aminase (GMP
synthetase), psicofuranine, 476, 481
Xanthurenate, urinary excretion of,
deoxypyridoxol, 572
Xanthyl- cytochrome c, cytochrome c, 592
Xylanase, mercurials, 859
Xylenesulfonate, arylsulfatase, 444
Xylitol, mutarose, 413-414
Xylono- 1 ,4-lactone,
a-glucuronidase, 426
structure of, 425
Xylose,
a-amylase, 420
fructokinase, 376
galactose transport by intestine, 263
/3-galactosidase, 418
a-glucosidase, 423
a-mannosidase, 422
mutarotase, 413-414
phosphoarabinose isomerase, 411
photosynthesis, 414
SUBJECT INDEX
1237
uptake by diaphragm, mercurials, 911-
912
Xylulokinase,
o-iodosobenzoate, 713
mercurials, 859
Yeast,
acetate oxidation, malonate, 77, 116
catalase induction in, 8-azaguanine, 478
citrate formation from acetate, ma-
lonate, 105
coenzyme A levels in, mercurials, 750,
885
cycle intermediate concentrations in,
89
2-deoxyglucose uptake, 387
ethanol oxidation, 2-deoxyglucose, 395-
396
fermentation (fructose), 6-deoxy-6-fluo-
roglucose, 404
fermentation (glucose),
biotin analogs, 588-589
6-deoxy-6-fluoroglucose, 404
D-glucosone, 384-385
iodine, 689
mercurials, 875, 884
tripolyphosphate, 383
fumarate oxidation, malonate, 81
glycolysis, 2-deoxyglucose, 392
growth of,
benzoate, 349
dehydroacetate, 632
o-iodosobenzoate, 727
malonate, 195
mercurials, 971, 973-974
cy-methylpyridoxol, 575
growth of (menadione-stimulated), fer-
ricyanide, 678
K+ fluxes, mercurials, 898-900, 908
maltase biosynthesis, p-fluorophenyla-
lanine, 351
mercurial uptake by, 898-900, 974
nucleotide levels in, mercurials, 884-
885
pyruvate oxidation, benzoate, 349
resistance to mercurials, 983-985
respiration (endogenous),
malonate, 169
mercurials, 879-881
respiration (glucose),
malonate, 124
mercurials, 880-881, 884-885
succinate accumulation, malonate, 92-
93
succinate dehydrogenase,
fumarate K,, 38
malonate, 27, 33, 187
succinate oxidation, malonate, 22
sulfate uptake, thiosulfate, 267
Yoshida ascites hepatoma, glycolysis (ae-
robic),
tartronate, 238
Yoshida sarcoma, respiration (endoge-
nous),
malonate, 179
Zahdel hepatoma, ADP-ATP levels in,
2-deoxyglucose, 395
Zinc,
complexes with di- and tricarboxylates,
12
glutamate dehydrogenase, 863
Lactobacillus growth, 452
phosphatase (acid), 452-453
Zygorrhynchus moelleri,
acetate oxidation, malonic diethyl
ester, 236-237
glucose oxidation, malonic diethyl
ester, 236-237
succinate oxidation, malonate, 51, 53,
187
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