[1st Edition] 9781483217048

Table of contents :
Content:
Contributors to Volume 5Page ii
Front MatterPage iii
Copyright pagePage iv
List of ContributorsPage ix
PrefacePage xiBERNARD L. HORECKER, EARL R. STADTMAN
Contents of Previous VolumesPages xiii-xvi
Phosphofructokinase*Pages 1-46TAG E. MANSOUR
A Theoretical Background to the Use of Measured Concentrations of Intermediates in Study of the Control of Intermediary MetabolismPages 47-75F.S. ROLLESTON
Memory MoleculesPages 77-97GÖTZ F. DOMAGK
Protein Kinases*Pages 99-133EDWIN G. KREBS
Glutamine Phosphoribosylpyrophosphate AmidotransferasePages 135-176JAMES B. WYNGAARDEN
The Regulatory Influence of Allosteric Effectors on Deoxycytidylate DeaminasesPages 177-228FRANK MALEY, GLADYS F. MALEY
The Citrate Enzymes: Their Structures, Mechanisms, and Biological FunctionsPages 229-283PAUL A. SRERE
Regulation of Histidine Biosynthesis in Salmonella TyphimuriumPages 285-308ROBERT F. GOLDBERGER, JOHN S. KOVACH
Author IndexPages 309-326
Subject IndexPages 327-331

Citation preview

Contributors to Volume 5 GÖTZ F. DOMAGK ROBERT F. GOLDBERGER JOHN S. KOVACH EDWIN G. KREBS FRANK MALEY GLADYS F. MALEY TAG E. MANSOUR F. S. ROLLESTON PAUL A. SRERE JAMES B. WYNGAARDEN

CURRENT TOPICS IN

Cellular Regulation edited by Bernard L Horecker Albert Einstein College of Medicine Bronx, New York

·

Earl R. Stadtman National Institutes of Health Bethesda, Maryland

Volume 5 7972

ACADEMIC PRESS

(^20) K

y

New York and London

COPYRIGHT © 1972, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED NO PART OF THIS BOOK MAY BE REPRODUCED IN ANY FORM, BY PHOTOSTAT, MICROFILM, RETRIEVAL SYSTEM, OR ANY OTHER MEANS, WITHOUT WRITTEN PERMISSION FROM THE PUBLISHERS.

ACADEMIC PRESS, INC. Ill Fifth Avenue, New York, New York 10003

United Kingdom Edition published by ACADEMIC PRESS, INC. (LONDON) LTD. 24/28 Oval Road, London NW1

LIBRARY OF CONGRESS CATALOG CARD NUMBER:

PRINTED IN THE UNITED STATES OF AMERICA

72-84153

List of Contributors Numbers in parentheses indicate the pages on which the authors' contributions begin.

F. DOMAGK* (77), Unité de Biochimie, Université de Louvain, Louvain, Belgium ROBERT F. GOLDBERGER (285), Laboratory of Chemical Biology, National Institute of Arthritis and Metabolic Diseases, National Institutes of Health, Bethesda, Maryland JOHN S. KOVACH (285), Laboratory of Chemical Biology, National Institute of Arthritis and Metabolic Diseases, National Institutes of Health, Bethesda, Maryland

GÖTZ

EDWIN G. KREBS (99), Department

of Biological Chemistry,

School of

Medicine, University of California, Davis, California FRANK MALEY (177), Division of Laboratories and Research, New York State Department of Health, Albany, New York GLADYS F. MALEY (177), Division of Laboratories and Research,

New

York State Department of Health, Albany, New York TAG E. MANSOUR (1), Department of Pharmacology, Stanford University School of Medicine, Stanford, California F. S. ROLLESTON (47), Banting and Best Department of Medical Research, University of Toronto, Toronto, Canada PAUL A. SRERE (229), Veteran's Administration

Hospital and

Department

of Biochemistry, University of Texas Southwestern Medical School, Dallas, Texas JAMES B. WYNGAARDEN (135), Department of Medicine, Duke University Medical Center, Durham, North Carolina

* Present address : Physiologisch-Chemisches Institut, 34 Göttingen, Humboldtallee 7, Germany. ix

Preface Recent years have witnessed rapid advances in our knowledge of the basic mechanisms involved in the regulation of diverse cellular activities such as intermediary metabolism, the transfer of genetic information, membrane permeability, and cellular differentiation and other organ functions. Information gained from -the detailed analyses of a large num­ ber of isolated enzyme systems, together with results derived from physio­ logical investigations of metabolic processes in vivo, constitutes an everincreasing body of knowledge from which important generalized concepts and basic principles of cellular regulation are beginning to emerge. How­ ever, so rapid are the present advances in the general area of cellular regulation and so diverse are the disciplines involved, that it has become a formidable task for even the expert in a specialized area to keep abreast of the progress in his field. This series of volumes is concerned with such recent developments in various areas of cellular regulation. We do not in­ tend that it will consist of comprehensive annual reviews of the literature. We hope rather that it will constitute a medium which will, on the one hand, provide contributing authors with an opportunity to summarize progress in specialized areas of study that have undergone substantial de­ velopments and, on the other hand, serve as a forum for the enunciation of general principles and for the formulation of provocative theories and novel concepts. To this end editorial review of individual contributions will be concerned primarily with the clarity of presentation and con­ formity to publication policies. It is hoped in this manner to bring together current knowledge of various aspects of cellular regulation so as both to enlighten the uninformed and to provide a base of knowledge for those engaged in research in this subject. BERNARD L. HORECKER EARL R. STADTMAN

Contents of Previous Volumes

Volume 1

Conformational Aspects of Enzyme Regulation D.E.

Koshland, Jr.

Limitation of Metabolite Concentrations and the Conservation of Solvent Capacity in the Living Cell Daniel E.

Atkinson

The Role of Equilibria in the Regulation of Metabolism H. A. Krebs Regulation of the Biosynthesis of the Branched-Chain Amino Acids H. E. Umbarger On the Roles of Synthesis and Degradation in Regulation of Enzyme Levels in Mammalian Tissues Robert T. Schimke The Regulation of the Biosynthesis of a-l,4-Glucans in Bacteria and Plants Jack Preiss Allosteric L-Threonine Dehydrases of Microorganisms W. A. Wood The Aspartokinases and Homoserine Dehydrogenases of Eschenchia coli Georges N. Cohen Pyruvate Dehydrogenase Complex Lester J. Reed xiii

XIV

CONTENTS OF PREVIOUS VOLUMES

Pyruvate Carboxylase Merton F. Utter and Michael C. Scrutton Author Index—Subject Index

Volume 2

DPN-Linked Isocitrate Dehydrogenase of Animal Tissues Gerhard W. E. Plant The Regulation of Biosynthesis of Aromatic Amino Acids and Vitamins J. Pittard and F. Gibson Regulation of Cholesterol Biosynthesis in Normal and Malignant Tissues Marvin D. Siperstein The Biogenesis of Yeast Mitochondria Anthony W. Linnane and J. M. Haslam Fructose 1,6-Diphosphatase from Rabbit Liver S. Pontremoli and B. L. Horecker The Role of Phosphoribosyltransferases in Purine Metabolism Kan 0. Raivio and J. Edwin Seegmiller Concentrations of Metabolites and Binding Sites. Implications in Meta­ bolic Regulation A. Sols and R. Marco A Discussion of the Regulatory Properties of Aspartate Transcarbamylase from Escherichia coli J. C. G erhart Author Index—Subject Index

CONTENTS OF PREVIOUS VOLUMES

XV

Volume 3

The Regulation of Branched and Converging Pathways B. D. Sanwal, M. Kapoor, and H. Duckworth The Role of Cyclic AMP in Bacteria Robert L. Perlman and Ira Pastan Cell Surfaces in Neoplastic Transformation Max M. Burger Glycogen Synthase and Its Control Joseph Lamer and Carlos Villar-Palasi The Regulation of Pyruvate Kinase Werner Seubert and Wilhelm Schoner Author Index—Subject Index

Volume 4

The Regulation of Arginine Metabolism in Saccharomyces Exclusion Mechanisms J. M. Wiame The Lac Repressor Suzanne Bourgeois L-Glutamate Dehydrogenases Barry R. Goldin and Carl Frieden Regulation of Fatty Acid Biosynthesis P. Roy Vagelos

cerevisiae:

CONTENTS OF PREVIOUS VOLUMES

XVI

Kinetic Analysis of Allosteric Enzymes Kasper

Kirschner

Phosphorylase and the Control of Glycogen Degradation Edmond H. Fischer, Ludwig M. G. Heilmeyer, Jr., and Richard H. Haschke Author Index—Subject Index

Phosphofructokinase'1 TAG E.

MANSOUR

Department of Pharmacology Stanford University School of Medicine Stanford, California I. Introduction—The Discovery of the Regulatory Role of Phosphofructokinase II. Molecular Structure and Association-Dissociation Systems . . A. Heart and Skeletal Muscle Enzymes B. Enzymes from Sources Other Than Heart and Skeletal Muscle III. Allosteric Kinetics A. Early Studies on the Kinetic Control of Phosphofructokinase B. The Effect of Different Modifiers on Phosphofructokinase Kinetics C. Studies on the Kinetics of Phosphofructokinase from Mammalian Sources Other Than Muscle D. Kinetics of Phosphofructokinase from Nonmammalian Sources IV. Nature of Allosteric Control A. Desensitization of Phosphofructokinase to Allosteric Inhibition B. Identification of Amino Acids That Influence Allosteric Kinetics C. ATP-Insensitive Form of Phosphofructokinase . . . . D. Ligand Binding by Native Enzyme and Enzyme Desensitized to Allosteric Control E. Relationship between Structure and Activity of Different Enzyme Modifiers and Substrates V. Identification of Functional Groups in the Enzyme . . . A. Role of Thiol Groups in Regulation and Catalysis . . . B. Role of Lysine and Tyrosine in Enzyme Activity . . . VI. Mechanism of the Reaction VII. Physiological Interpretation of Phosphofructokinase Regulation A. Synchrony between Cellular Demand for High Energy and Enzyme Activity B. The Role of Citrate in Phosphofructokinase Regulation .

2 4 4 11 11 11 12 14 15 17 17 17 19 20 24 25 25 28 29 30 30 33

* Some of the studies carried out by the author and reviewed here were supported by the U. S. Public Health Service Research Grant (AI04214) from the National Institute of Allergy and Infectious Diseases and a grant-in-aid from the American Heart Association. 1

2

TAG E. MANSOUR

C. Influence of Hormones on Phosphofructokinase Activity D. Activation of Phosphofructokinase by Serotonin in the Liver Fluke Fasciola hepatica VIII. Complementarity in Enzyme Regulation IX. Conclusion References

.

34 37 40 42 42

I. Introduction—The Discovery of the Regulatory Role of Phosphofructokinase A role for phosphofructokinase (ATP:D-fructose-6-phosphate 1-phosphotransferase, EC 2.7.1.11) in the regulation of glycolysis was first sug­ gested by Cori in the course of his classical investigations on muscle metabolism during work {24). He reported that the reaction catalyzed by phosphofructokinase [Reaction (1)] is rate limiting. This was indi­ cated by the fact that the hexosemonophosphate can be increased under certain experimental conditions without any increase in lactic acid {26, Fructose-6-P + ATP -> fructose-1,6-di-P + ADP

(1)

27y 41 ) · For example, incubation of frog muscle with epinephrine was fol­ lowed by an increase in the endogenous concentration of the hexose monophosphate, as a result of glycogen phosphorylase activation, while the increase in lactic acid was smaller. Similarly, contraction of the gastrocnemius muscle following electric stimulation results in a marked accumu­ lation of hexose monophosphate esters and an increase in lactic acid production. Since in these experiments the sum of the hexose monophos­ phates and lactic acid formed can account for the amount of glycogen lost during contraction, the assumption was made that other intermedi­ ates of glycolysis do not accumulate to an appreciable extent. The experi­ ments thus indicated "that the reaction glycogen -» hexose monophos­ phate occurs more rapidly than the reaction hexose monophosphate -» lactic acid and points to phosphofructokinase as the rate-limiting step for lactic acid formation during contraction" {24, 25). The idea that phosphofructokinase is a key enzyme in regulating glycolysis was adopted by several workers who were interested in under­ standing the mechanism of the Pasteur effect. Engelhardt and Sakov {32) found that the enzyme is highly sensitive to oxidative agents, an effect they thought could explain the role of the enzyme in the regulation of glycolysis during the shift from anaerobic to aerobic conditions. Since it was not possible to demonstrate an actual increase in the activity of phosphofructokinase following stimulation of glycolysis during anoxia or after muscle contraction (similar to that observed with phosphorylase),

PHOSPHOFRUCTOKINASE

3

an indirect means was used to get an idea about changes in enzyme ac­ tivity in the intact cell. The procedure of determining the steady-state levels of intermediary metabolites, originally used by Cori, was used by several early investigators in this problem to assess this point. For ex­ ample, Lynen et al (66) reported that, in yeast cells, anoxia, which causes an increase in the rate of glucose phosphorylation, also causes a reduction in the intracellular concentration of free-glucose and fructose-6-P as well as an increase in the intracellular levels of fructose1,6-di-P. This meant a decrease in the levels of the enzyme substrate and an increase in its products and suggested an increase in the activity of phosphofructokinase. The use of substrate and product levels to determine enzyme activity following anoxia was subsequently studied in the heart (89) and in the brain {65). Similarly antimonial agents were shown to affect phosphofructokinase in schistosomes (18, 77) and 5-hydroxytryptamine (serotonin) affected phosphofructokinase from the liver fluke (Fasciola hepatica) (69, 79). What was originally intended by the author to be a project to learn more about flukes turned out to be a twelve-year investigation on the cellular regulation of phosphofructokinase. All these investigations were a prelude to a much greater interest in phosphofructokinase. The interest was partially prompted by the realiza­ tion that the enzyme has an important physiological role analogous to that of glycogen phosphorylase in the regulation of carbohydrate metab­ olism. A thorough understanding of the regulatory role of phosphofructo­ kinase had to await a better knowledge of the molecular properties of the enzyme. It is the intent of this review to discuss the structural prop­ erties of the enzyme particularly as it relates to its activity. Since the unusual kinetics of the enzyme appear to play an important role in its regulation and have been extensively studied in our laboratory as well as others, they will receive special emphasis. Although the work on the mechanism of phosphofructokinase reaction is not yet conclusive, a review of the present status in this aspect is included. Physiological interpretation of some of the apparent changes of phosphofructokinase during different physiological changes in the cell is discussed. The evi­ dence is now accumulating, indicating complementarity in the regulation of different enzymes. A special section in this review is devoted to several examples where the regulation of phosphofructokinase appears to com­ plement the regulation of other enzymes. Since most of the work on phosphofructokinase has been done on mammalian skeletal muscle or heart enzymes, these enzymes will receive the greatest emphasis. How­ ever, the properties of enzymes from some other tissues as well as other sources are also discussed.

4

TAG E. MANSOUR

II. Molecular Structure and AssociationDissociation Systems A. Heart and Skeletal Muscle Enzymes One reason for the delay in our knowledge of the molecular properties of phosphofructokinase was the unavailability of purified enzyme prep­ arations. Early attempts to purify the enzyme were handicapped by its marked instability (86,124), particularly at a midly acidic pH (28, 127). Low degree of purification of the enzyme results in its inactivation, sug­ gesting that certain tissue components are essential for its stability. These tissue components were later identified to be hexose phosphates and adenine nucleotides. Of these agents, fructose- 1,6-di-P was the most active in stabilizing the enzyme (131). The use of these stabilizers facilitated the purification of the enzyme from the sheep heart to homogeneity (82) with good yield. Other purified enzymes were isolated from skeletal muscle (60, 98), erythrocytes (56), yeast (9), E. coli (16), and liver

(17,W). Early experiments with partially purified phosphofructokinase have indicated that the enzyme is reversibly converted to an inactive form. To those who are working in the area of the regulation of carbohydrate metabolism it suggested an enzyme-catalyzed active-inactive system similar to that known for glycogen phosphorylase. Now that the pure enzyme is available the evidence points to a pH- and ligand-directed association-dissociation system. Experiments on partially purified phos­ phofructokinase showed that, at a pH range of 5.8-6.5, enzyme activity when measured under optimal conditions was reduced to about 10% (72). The inactivation process is rapid, and accurate study of its kinetics could not be carried out. Mild acidification of the enzyme when carried out at low enzyme concentrations results in an irreversible loss of activity. How­ ever, at moderately high protein concentrations the inactive enzyme can be reactivated by incubation at pH 8.0. The sedimentation velocity of the inactive enzyme varies from 7 S to 8 S. The reactivated enzyme has a sedimentation coefficient which is identical to that of the native enzyme (14S-15S). While enzyme dissociation is almost instantaneous, the process of reassociation is slower (72). It is time dependent, enhanced by temperature and by several ligands. Reactivation of the enzyme is increased by the following nucleotides and hexose phosphates: ATP, ADP, cyclic 3',5'-AMP, fructose-6-P, and fructose-1,6-di-P. All these agents are also allosteric effectors of the enzyme. Combinations of a nucleotide and a hexose phosphate are much more effective in reactivation of phos­ phofructokinase than either nucleotide or hexose phosphate alone (Fig. 1).

5

PHOSPHOFRUCTOKINASE

5 4 3 2. - l o g fructose-ö-P concentration iM)

=> Θ

7

6

5

A

-log FDP concentration

3

(M)

FIG. 1. Effect of hexose phosphates on reactivation of phosphofructokinase. Enzyme was inactiviated by incubation in 0.05 M Tris maleate buffer, pH 5.8, at 37°C. Assays were carried out as described before (72). Reactivation was carried out in the presence of 0.08 M glycylglycine buffer, pH 7.5. Initial enzyme activity was 155 units/ml and was reduced to 5.4 units/ml after inactivation at pH 5.8. All enzyme activities were calculated back to the initial concentrated enzyme. The ordinates at the left give the net enzyme activity recovered in 10 minutes. The ordinates at the right give enzyme recovery as percentage of the initial enzyme ac­ tivity. The abscissas give negative log molar concentration of (a) fructose 6-phosphate and (b) fructose 1,6-diphosphate (FDP). Data from Mansour (72).

The process of reactivation is dependent on protein concentration in the absence of any ligand. Thus, as mentioned above, in the absence of an activator and at low enzyme concentration, no reactivation occurs. The pH-dependent dissociation system has been verified in purified enzyme preparations both from the skeletal muscle (93) and the heart (76). Paetkau and Lardy (93) demonstrated that at pH 5.8 and in the pres­ ence of 0.8 M urea and 0.5 milf of fructose- 1,6-di-P a form with mo­ lecular weight of 1.92 X 105 and sedimentation coefficient of 7 S was obtained. The specific activity of this form of the enzyme was 2 to 7 in­ stead of 140 for the native enzyme. When the pH was raised to 8, reaggregation occurred and was accompanied by reactivation. The re­ activation was shown to be due to the change in the state of aggregation

6

TAG E. MANSOUR

■■■illilillili

18 min

22 min

14 min

22 min

B

FIG. 2. Schlieren patterns of phosphofructokinase taken from data by Mansour et al. (82). (A) Crystalline phosphofructokinase taken in a double sector cell, solvent, 0.05 M potassium phosphate buffer (pH 8.0) ; 0.01 M 2-mercaptoethanol ; IO"4 M ATP; 10"5M fructose-l,6-di-P. (B) Schlieren patterns of the enzyme before crystal­ lization but having the same specific activity as in (A) in a Yphantis-Waugh moving partition cell. Protein concentration, 0.612% in 0.05 M potassium phosphate buffer (pH 8); 0.005M 2-mercaptoethanol; 10~4Μ ATP and lO^ikf fructose-l,6-di-P.

PHOSPHOFRUCTOKINASE

7

of the enzyme, not to pH. Various oligomeric forms of phosphofructokinase have been reported by Lardy's group {92, 93) with the following molecular weights: 43,000, 191,000, 380,000, 770,000, and 1,540,000. In an attempt to find whether reactivation of the enzyme is catalyzed by an enzyme system, the effect of heart extracts on enzyme reactivation was tested {72). Heart extracts reactivated acid-inactivated enzyme in the presence of ATP/Mg 2+ . This effect does not appear to be an enzymatic phosphorylation of phosphofructokinase for the following reasons: (a) boiled heart extracts have at least 40% of the fresh extract activity; thus a large part of the effect of the extracts is probably due to a nonenzymatic component; (b) ATP can be replaced without loss of activity with AMP, ADP, or cyclic 3'5'-AMP; (c) Mg-+ was not essential for the combined effect of ATP and heart extract. In view of this, it may be that the effect of the fresh heart extract in accelerating phosphofructo­ kinase reactivation in the presence of ATP/Mg 2 + is due to the presence of an endogenous hexose phosphate or another tissue component that might enhance the effect of the adenylic nucleotides. One important property of phosphofructokinase is its ability to form aggregates. Sedimentation coefficient determination of high enzyme con­ centrations revealed that it is present as high aggregates {82). For ex­ ample, in the concentrations required for the determination of schlieren patterns in the analytical ultracentrifuge (2 mg/ml), the crystalline en­ zyme gave a schlieren pattern with an asymmetrical peak with an s20,w of 25.4 (Fig. 2A). The fact that the schlieren pattern had a sharp boundary at the bottom and a partially skewed edge trailing toward the top of the cell suggests a monomer-polymer system in rapid equilibrium. In other experiments the schlieren pattern was composed of the area with an s20,w value of 41 and a light component which had an s20,w value of 8.2 (Fig. 2B). The s20,w value for the heavy component varies from 38 to 51. Separation of the two components through the use of the Yphantis-Waugh moving partition cell {143) revealed that the light component had lower specific activity than the heavy component. The double peak schlieren pattern could therefore represent an associating-dissociating system in equilibrium, as was shown first by Gilbert with other enzymes {Jfi). The effect of different ligands on the different molecular forms is of interest since it may suggest control through changes in enzyme mo­ lecular forms by different substrates {82, 97). Of particular interest are the substrates and the products of the enzyme. When the enzyme was tested under conditions that favor its aggregation into a single sedimenting peak (53 S) (top pattern in Fig. 3), ATP/Mg 2 + resulted in the ap­ pearance of multiple peaks with s20,w values of 7.1, 38.7, 44.4 accounting for 16, 58, and 33% of the total area, respectively (bottom pattern in

8

TAG E. MANSOUR

Fig. 3). On the other hand, when the enzyme was studied under condi­ tions that favor enzyme dissociation (pH 6.5) ATP was shown to favor the equilibrium toward the 7 S inactive form while fructose-6-P or fructose-l,6-di-P favors the associated fully active form (82). Experiments were carried out in several laboratories to investigate the nature of the subunits of both heart and skeletal enzymes. Ultracentrifugal analysis of heart enzyme in 5 M guanidine HCl revealed a single peak with an s20,w value of 2.75 (76). Similarly the skeletal muscle en-

18 min

26 min

26 min

36 min

FIG. 3. Schlieren patterns of phosphofructokinase taken in a double sector cell, solvent; 0.05M glycylglycine buffer pH 6.9 and 0.05M mercaptoethanol, Top: No ATP, no MgCl2; bottom: with 2 X 10"3M ATP and 4 X 10"3M MgCl2. From the data by Mansour et al. (82).

PHOSPHOFRUCTOKINASE

9

zyme gives a single peak of 2.3 S in the presence of 5.5 M guanidine HC1 or 8 M urea {94). The molecular weight of these subunits when de­ termined on Sephadex columns was found to be 23,000 to 24,000 for the heart enzyme {115). The subunit of the skeletal muscle enzyme was determined by Paetkau et al. {94) using the equilibrium ultracentrifugation procedure and was found to be 24,000 to 25,000. Experiments in our laboratory using acrylamide electrophoresis and electrofocusing procedure indicated that phosphofructokinase subunits are identical. However, re­ sults based on the procedure of tryptic peptide mapping first reported by Paetkau et al. {94) with the skeletal muscle enzyme and confirmed on the heart enzyme in our laboratory {115) indicated that there are more than one kind of subunit for phosphofructokinase. Further experi­ ments are necessary to explain the discrepancy between the results of the tryptic peptide mapping and other physicochemical data. The data discussed above can be presented in a tentative model for the molecular structure of phosphofructokinase from either heart muscle or skeletal muscle. The scheme represents fully active phosphofructokinase as a tetramer with a molecular weight of 3.6 X 105 which can be reversibly converted to several aggregated forms with as yet an undefined molecular weight. The equilibrium in such a conversion is a function of enzyme concentration. ATP favors dissociation of these enzyme aggre­ gates. The specific activity of the aggregated forms has as yet not been defined because enzyme at assay levels is highly diluted and presumably is in the nonaggregated form. This information will have to await studies on the activity of high enzyme concentrations. The fact, however, that the binding of both substrates is reduced at high protein concentrations {62) suggests a lower specific activity for these aggregates. According to the model in Fig. 4 the tetrameric form of the enzyme can be reversibly dissociated at a mildly acidic pH to a relatively inactive dimeric form with a molecular weight of 1.8 X 105. The degree of dissociation is a function of pH, enzyme concentration and ATP concentration. The dis­ sociated enzyme can be reassociated at an alkaline pH. High enzyme concentration and the presence of fructose-6-P or fructose-1,6-di-P favors association of the enzyme. The scheme also shows that phosphofructo­ kinase beside being able to be dissociated to 4 protomers can be further converted in 5-6 M guanidine HC1 to 16 small subunits which are approxi­ mately 24,000 in molecular weight. Reversal of this dissociation has not as yet, been achieved. The enzyme in fresh extracts from skeletal muscle or the heart was shown in our laboratory to exist in different aggregated forms. Presuma­ bly in the cell at high ATP levels the equilibrium is shifted to low mo­ lecular forms which is more sensitive to dissociation to smaller inactive

10

TAG E. MANSOUR

AGGREGATES SA = not defined MW =not defined | s 32 6 - 5 4 20.W è Q.

S

"6

σ »φ Φ φ

M

£ 9 X

υ 3 Ü.

*σ *-

1 • E

0.

5

Ν C Φ

3*

r

FULLY ACTIVE PFK MW » 3.6 x 10 8 2 Q , W * 13-14 i e

ο Β TO

»

it Ν or

C Φ £σ» f

k

2 Ο o 3 ω X ul Q.

s

4 m/ifSDS.■*

^

± 4 PROTOMERS No known octivity 4 MW«9x IO

5 M guonidine HCL

► 16 SUBÜNITS MW = 2 4 x I 0 s 9 f t l«2.75 if

4

| |

c Φ

o c o Ü

Φ

E Ü.

5

h-


+ and the par­ ticulate fraction is just as effective provided cyclic 3',5'-AMP is present. The process of activation is time and temperature dependent, can be reversed by dialysis, and appears to involve a monomer-polymer con­ version. Sucrose gradient ultracentrifugai analysis of the enzyme before and after activation gave an s20,w of 5.5 S for the inactive enzyme and 12.8 S for the activated enzyme. More information concerning the nature of the fluke enzyme will have to await its purification. In.addition to a control mechanism that involves activation of an inactive form of phos­ phofructokinase, the fluke enzyme, once activated, is subject to all the allosteric kinetics that have been described above (121). The experiments described above on flukes not only focused attention on phosphofructokinase as a regulatory enzyme but also on serotonin, rather than epinephrine, as a possible hormone affecting carbohydrate metabolism in invertebrates. Subsequent experiments in other laboratories showed that the serotonin has metabolic effects on other invertebrate tissues similar to those described above (84, HO, 111). In all these experi­ ments epinephrine either has no effect on the carbohydrate metabolism, as in the case with the flukes, or could show some effect at concentrations which are much higher than that of serotonin. All these findings strengthen the idea that serotonin or a related indolealkylalamine has a hormonal function in invertebrates similar to that of epinephrine in higher organisms (33, 74, 81, 136). At a much higher phylogenetic level (mam­ mals), serotonin has little or no effect on the carbohydrate metabolism

40

TAG E. MANSOUR

and does not act as a humoral transmitter. At that level epinephrine is the amine responsible for the control of both physiological functions. VIII. Complementarity in Enzyme Regulation The complex nature of the control of phosphofructokinase makes one wonder whether the control of other related enzymes could complement phosphofructokinase regulation for the benefit of cellular metabolic con­ trol. With the recent accumulation of information on the control of these enzymes, we are now beginning to see that such complementarity does exist. AMP appears to play an important role in the regulation of en­ zymes catalyzing glycogen and glucose· utilization. An increase in the level of AMP in the cell, besides activating phosphofructokinase, will also simultaneously inhibit the activity of fructose-l,6-diphosphatase. Activation of the first enzyme would favor glycogen breakdown and also increase the substrate flux through the glycolytic enzyme system. Inhibi­ tion of fructose-1,6-diphosphatase by AMP would favor an increase in glycogen breakdown and could maintain high levels of fructose-1,6diphosphate. The latter effect tends to shift the direction of the metabo­ lism toward glycolysis rather than toward glycogen synthesis. The high levels of fructose-l,6-di-P in the cell would amplify these effects by being the allosteric activator of both phosphofructokinase and pyruvic kinase. Recent studies on the effect of sulfhydryl reagents on phosphofructo­ kinase and fructose-1,6-diphosphatase shed further light on complemen­ tarity in the design of the two enzymes. Pontremoli and Horecker have recently reported that disulfides, such as cystamine, activate fructose-1,6diphosphatase (104). The same reagents have been reported to inhibit phosphofructokinase (37). It is therefore possible that, in addition to having the common effector, AMP, these two enzymes could be controlled by the level of disulfides present in the cell, as has been suggested for fructose diphosphatase (103). The presence of fructose-1,6-diphosphatase in the muscle has been puzzling to investigators since in the muscle glycolysis is not reversed toward glycogen synthesis as it is in the liver. Newsholme and Crabtree (88) have recently discussed the role of this enzyme in the regulation of glycolysis. They found that in muscles with an extremely high demand for energy during contraction, such as the pheasant pectoral muscle, the activity of phosphofructokinase has to be stimulated to 90-fold that of the resting level activity. Presumably in these muscles, activation of phosphofructokinase through changes in AMP concentration is not suf­ ficient. Accordingly the sensitivity of the control mechanism can be in­ creased by the operation of a cycle between fructose-6-P and fructose1,6-di-P catalyzed by the simultaneous activities of phosphofructokinase

PHOSPHOFRUCTOKINASE

41

and fructose-1,6-diphosphatase in response to changes in AMP concen­ tration. In case of a great demand of energy for the muscle, the increase in the level of AMP will ensure simultaneous inhibition of the fructose diphosphatase and activation of phosphofructokinase. Such an effect would maintain a high level of fructose-1,6-di-P and could ensure that restriction of the rate of glycolysis at the stage of fructose-6-P is reduced to a minimum. AMP appears also to synchronize the activity of glycogen phosphorylase b and phosphofructokinase. Both enzymes are activated by the adenine nucleotide. There is also complementarity between phospho­ fructokinase and glycogen phosphorylase kinase. The procedure which was used to isolate the latter enzyme was reported to copurify phospho­ fructokinase (97). Furthermore cyclic 3',5'-AMP and epinephrine increase the activity of both enzymes at pH 6.9 while the activity of both enzymes at pH 8.2 is not significantly changed (53). The possible role of citrate in the regulation of phosphofructokinase was discussed above. This ligand appears to occupy a central role in the regulation of energy metabolism. It inhibits phosphofructokinase activity at levels which might well exist in the cell (87, 108). Oxidation of fatty acids and ketone bodies in diabetes and starvation results in an increase in the levels of citrate, a process which could inhibit phosphofructokinase and activate acetyl-CoA carboxylase in fat cells and liver (129a). These two complementary regulatory mechanisms on two different enzymes are designed to provide energy to the muscle from fatty acids rather than from carbohydrate and to stimulate fatty acid synthesis in the liver and in fat deposits. Mention also should be made here to the complementarity between hexokinase and phosphofructokinase. The former enzyme is known to be inhibited by glucose-6-P. An increase in phosphofructokinase activity will result in a decrease in the cellular levels of fructose-6-P. Since there is equilibrium between glucose-6-P and fructose-6-P through the phosphoglucose isomerase reaction, the level of glucose-6-P will also fall. This will result in a parallel increase in the activity of hexokinase. More recent studies on hexokinase by Berthillier et al. (13, 14) show striking simi­ larities between the kinetics of a form of glucokinase (GM1) isolated from the liver and phosphofructokinase. This form of the enzyme, like phosphofructokinase, is inhibited by ATP and activated by AMP. The inhibition by ATP appears to be allosteric in nature since the enzyme can be desensitized to such inhibition by méthylène blue without much loss to catalytic activity. Another form of hexokinase (GM2) has been isolated by these authors which is half the molecular weight of the ATPsensitive form (50,000) and was shown to be insensitive to allosteric

42

TAG E. MANSOUR

control. This illustrates another way of having complementarity in the regulation of glucokinase and phosphofructokinase. IX. Conclusion This reviewer has attempted to outline the structural features of phos­ phofructokinase, its principal kinetic properties, and the different mecha­ nisms that have been proposed for its regulation. One conclusion which seems to be overwhelmingly accepted as a result of studies on species at different phylogenetic levels is that the activity of this enzyme is geared to energy demand by the cell. The reader of this review will note that the striking feature of this enzyme is the multiplicity of its molecular forms and the large number of modifying ligands available. After con­ sidering the innumerable factors that could modify enzyme activity the reader may well come to the conclusion that phosphofructokinase is be­ yond control. This is a pessimistic view with which the author cannot concur. The fact that phosphofructokinase occupies a crucial position within the cell's metabolic machinery may explain the variety of ways that the activity of the enzyme could be changed. Future advances in our knowledge of the regulation of phosphofructokinase will have to await studies on the nature of the enzyme in the resting cell and its response to different physiological conditions. Studies on the relationship between enzyme aggregates and their catalytic activity may shed some light on the physiological significance of these forms of the enzyme. Last, a better understanding of the mechanism of the phosphofructokinase reaction will certainly contribute to a better understanding of the regulation of this important enzyme. REFERENCES 1. Abrahams, L. S., and Younathan, E. S., J. Biol. Chem. 246, 2464 (1971). 2. Afting, E . G., Ruppert, D., Hagmaier, V., and Hölzer, H., Arch. Biochem. Biophys. 143,587 (1971). 3. Ahlfors, C. E., and Mansour, T. E., / . Biol. Chem. 244, 1247 (1969). 4. Atkinson, D . E., Annu. Rev. Biochem. 35, 85 (1966). 5. Atkinson, D. E., Biochemistry 7, 4030 (1968). 6. Atkinson, D. E., Annu. Rev. Microbiol. 23, 47 (1969). 7. Atkinson, D . E., in "The Enzymes" (P. Boyer, ed.), 3rd ed., Vol. 1, p. 461. Academic Press, New York, 1970. 8. Atkinson, D . E., and Walton, G. M., J. Biol. Chem. 240, 757 (1965). 9. Atzpodien, W., and Bode, H., Euro. J. Biochem. 12, 126 (1970). 10. Atzpodien, W., Gancedo, J. M., Hagmaier, V., and Hölzer, H., Eur. J. Biochem. 12, 6 (1970). 11. Baumann, P., and Wright, B. E., Biochemistry 7, 3653 (1968). 12. Beitner, R., and Kalant, N., / . Biol. Chem. 246, 500 (1971).

PHOSPHOFRUCTOKINASE

43

13. Berthillier, G., Colobert, L., Richard, M., and Got, R., Biochim. Biophys. Ada 206, 1 (1970). 14. Berthillier, G., and Got, R., Biochim. Biophys. Ada 258, 88 (1972). 15. Betz, A., and Moore, C., Arch. Biochem. Biophys. 120, 268 (1967). 16. Blangy, D., Bue, H., and Monod, J., / . Mol. Biol. 31, 13 (1968). 17. Brock, D . J. H., Biochem. J. 113, 235 (1969). 18. Bueding, E., and Mansour, J. M., Brit. J. Pharmacol. 12, 159 (1957). 19. Chapman, A., Sanner, T., and Pihl, A., Eur. J. Biochem. 7, 588 (1969). 20. Chapman, A., Sanner, T., and Pihl, A., Biochim. Biophys. Ada 178, 74 (1969). 21. Cleland, W. W., Biochim. Biophys. Ada 67, 104 (1963). 22. Cleland, W. W., Anna. Rev. Biochem. 36, 77 (1967). 23. Colowick, S. P., Abstr., 112th Meet., Amer. Chem. Soc. p. 56C (1947). 24- Cori, C. F., in "A Symposium on Respiratory Enzymes," p. 175. Univ. of Wis­ consin Press, Madison, 1942. 25. Cori, C. F., in "Enzymes: Units of Biological Structure and Function" (O. H. Gaebler, ed.), p. 573. Academic Press, New York, 1956. 26. Cori, G. T., and Cori, C. F., J. Biol. Chem. 116, 119 (1936). 27. Cori, G. T., and Cori, C. F., J. Biol. Chem. 116, 129 (1936). 28. Cori, G. T., and Illingworth, B., Biochim. Biophys. Acta 21, 105 (1956). 29. Danforth, W. H., and Helmreich, E., / . Biol. Chem. 239, 3133 (1964). 30. Dennis, D. T., and Coultate, T. P., Biochim. Biophys. Ada 146, 129 (1967). 31. El-Badry, A., Otani, A., and Mansour, T. E., manuscript in preparation (1972). 32. Engelhardt, V. A., and Sakov, N . E., Biokhimiya 8, 9 (1943). 33. Erspamer, V., in "Handbuch der experimentellen Pharmakologie" (O. Eichler and A. Farah, eds.), Vol. 19, p. 132. Springer-Verlag, Berlin and New York, 1966. 34. Forest, P . B., and Kemp, R. G., Biochem. Biophys. Res. Commun. 33, 763 (1968). 35. Freyer, R., Liebe, S., Kopperschlager, G., and Hofmann, E., Eur. J. Biochem. 17, 386 (1970). 36. Frieden, C , in "The Regulation of Enzyme Activity and Allosteric Interactions" (E. Kvamme and A. Pihl, eds.), p. 59. Academic Press, New York, 1968. 37. Froede, H . C , Geraci, G., and Mansour, T. E., J. Biol. Chem. 243, 6021 (1968). 38. Garland, P . B., Rändle, P . J., and Newsholme, E . A., Nature {London) 200, 169 (1963). 39. Gerhart, J. C , and Schachman, H . K., Biochemistry 4, 1054 (1965). 40. Gilbert, G. A., Proc. Roy. Soc, Ser. A 250, 377 (1959). 41. Hegnauer, A. H., and Cori, G. T., / . Biol. Chem. 105, 691 (1934). 42. Helmreich, E., and Cori, C. F., Pharmacol. Rev. 18, 189 (1966). 43. Helmreich, E., Danforth, W. H., Karpatkin, S., and Cori, C. F., in "Control of Energy Metabolism" (B. Chance, R. W. Eastabrook, and J. R. Williamson, eds.), p. 299. Academic Press, New York, 1965. 44- Hill, A. V., / . Physiol (London) 40, 4P (1910). 45. Hochrein, H., and Döring, H . J., Pfluegers Arch. Gesamte Physiol. Menschen Tiere 267,313 (1958). 46. Hoskins, D . D., and Stephens, D. T., Biochim. Biophys. Ada 191, 292, 1969. 47. Hulme, E . C , and Tipton, K. F., Biochem. J. 122, 181 (1971). 48. Kemp, R. G., Biochemistry 8, 4490 (1969). 49. Kemp, R. G., J. Biol. Chem. 246, 245 (1971). 50. Kemp, R. G., and Forest, P . B., Biochemistry 7, 2596 (1968).

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Kemp, R. G., and Krebs, E. G., Biochemistry 6, 423 (1967). Koshland, D. E., Jr., Némethy, G., and Filmer, D., Biochemistry 5, 365 (1966). Krebs, E. G., Graves, D. J., and Fischer, E. H., / . Biol. Chem. 234, 2867 (1959). Krebs, H . A., Broc. Roy. Soc, Ser. B 159, 545 (1964). Krzanowski, J., and Matschinsky, F . M., Biochem. Biophys. Res. Commun. 34, 816 (1969). 56. Kuhn, B., Jacobasch, G., and Rapoport, S., Acta Biol. Med. Ger. 23, 1 (1969). 57. Lardy, H. A., and Parks, R. E., Jr., in "Enzymes: Units of Biological Structure and Function" ( 0 . H. Gaebler, ed.), p. 584. Academic Press, New York, 1956. 58. Liebe, S., Kopperschlager, G., Diezel, W., Nissler, K., Wolff, J., and Hofmann, E., FEBS Lett. 8, 20 (1970). 59. Lindell, T. J., and Stellwagen, E., / . Biol. Chem. 243, 907 (1968). 60. Ling, K. H., Marcus, F., and Lardy, H . A., J. Biol. Chem. 240, 1893 (1965). 61. Lorenson, M. Y., and Mansour, T. E., J. Biol. Chem. 243, 4677 (1968). 62. Lorenson, M. Y., and Mansour, T. E., / . Biol. Chem. 244, 6420 (1969). 63. Lowry, O. H., and Passonneau, J. V., Naunyn-Schmiedebergs Arch. Exp. Pathol. Pharmakol. 248, 185 (1964). 64. Lowry, O. H., and Passonneau, J. V., / . Biol. Chem. 241, 2268 (1966). 65. Lowry, O. H., Passonneau, J. V., Hasselberger, F . X., and Schulz, D . W., J. Biol. Chem. 239, 18 (1964). 66. Lynen, F., Hartmann, G., Netter, K. F., and Schuegrat, A., Regni. Cell Metab., Ciba Found. Symp., 1958 p. 256 (1959). 67. Mansour, T. E., Biochim. Biophys. Acta 34, 464 (1959). 68. Mansour, T. E., / . Pharmacol. Exp. Ther. 126, 212 (1959). 69. Mansour, T. E., / . Pharmacol. Exp. Ther. 135, 94 (1962). 70. Mansour, T. E., J. Biol. Chem. 238, 2285 (1963). 71. Mansour, T. E., Advan. Pharmacol. 3, 129 (1964). 72. Mansour, T. E., J. Biol. Chem. 240, 2165 (1965). 73. Mansour, T. E., Pharmacol. Rev. 18, 173 (1966). 74- Mansour, T. E., in "Biogenic Amines as Physiological Regulators" (J. J. Blum, ed.), p. 119. Prentice-Hall, Inc., Englewood Cliffs, New Jersey, 1970. 75. Mansour, T. E., manuscript in preparation (1972). 76. Mansour, T. E., and Ahlfors, C. E., J. Biol. Chem. 243, 2523 (1968). 77. Mansour, T. E., and Bueding, E., Brit. J. Pharmacol. 9, 459 (1954). 78. Mansour, T. E., Lago, A. D., and Hawkins, J. L., Fed. Proc, Fed. Amer. Soc. Exp. Biol. 16,319 (1957). 79. Mansour, T. E., and Mansour, J. M., J. Biol. Chem. 237, 629 (1962). 80. Mansour, T. E., and Stone, D . B., Biochem. Pharmacol. 19, 1137 (1970). 81. Mansour, T. E., Sutherland, E. W., Rail, T. W., and Bueding, E., J. Biol. Chem. 235,466 (1960). 82. Mansour, T. E., Wakid, N., and Sprouse, H. M., J. Biol. Chem. 241, 1512 (1966). 83. Monod, J., Wyman, J., and Changeux, J.-P., J. Mol. Biol. 12, 88 (1965). 84. Moore, K. E., and Gosselin, R. E., J. Pharmacol. Exp. Ther. 138, 145 (1962). 85. Muntz, J. A., / . Biol. Chem. 171, 653 (1947). 86. Muntz, J. A., Arch. Biochem. Biophys. 42, 435 (1953). 87. Newsholme, E . A., in "Essays in Cell Metabolism" (W. Bartley, H. L. Kornberg, and J. R. Quayle, eds.), p. 189. Wiley (Interscience), New York, 1970. 88. Newsholme, E. A., and Crabtree, B., FEBS Lett. 7, 195 (1970). 89. Newsholme, E . A., and Rändle, P . J., Biochem. J. 80, 655 (1961). 90. Newsholme, E. A., and Rändle, P . J., Biochem. J. 93, 641 (1964).

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91. 92. 93. 94. 95.

45

Ozand, P., and Narahara, H . T., J. Biol. Chem. 239, 3146 (1964). Paetkau, V., Biochemistry 6, 2767 (1967). Paetkau, V., and Lardy, H . A., J. Biol. Chem. 242, 2035 (1967). Paetkau, V., Younathan, E . S., and Lardy, H. A., / . Mol. Biol. 33, 721 (1968). Park, C. R., Morgan, H. E., Henderson, M. J., Regen, D. M., Cadenas, E., and Post, R. L., Recent Progr. Hor. Res. 17, 493 (1961). 96. Parmeggiani, A., and Bowman, R. H., Biochem. Biophys. Res. Commun. 12, 268 (1963). 97. Parmeggiani, A., and Krebs, E. G., Biochem. Biophys. Res. Commun. 19, 89 (1965). 98. Parmeggiani, A., Luft, J. H., Love, D . S., and Krebs, E. G., J. Biol. Chem. 241, 4625 (1966). 99. Passonneau, J. V., and Lowry, O. H., Biochem. Biophys. Res. Commun. 7, 10 (1962). 100. Passonneau, J. V., and Lowry, 0 . H., Biochem. Biophys. Res. Commun. 13, 372 (1963). 101. Passonneau, J. V., and Lowry, O. H., Advan. Enzyme Regul. 2, 265 (1964). 102. Pogson, C. J., and Rändle, P . J., Biochem. J. 100, 683 (1966). 108. Pontremoli, S., and Horecker, B . L., Curr. Top. Cell. Regul 3, 174 (1970). 104. Pontremoli, S., Traniello, S., Enser, M., Shapiro, S., and Horecker, B. L., Proc. Nat. Acad. Sci. U. S. 58, 286 (1967). 105. Posner, J. B., Stern, R., and Krebs, E . G., / . Biol. Chem. 240, 982 (1965). 106. Rail, T. W., and Sutherland, E . W., / . Biol. Chem. 232, 1065 (1958). 107. Ramaiah, A., Hathaway, J. A., and Atkinson, D. E., J. Biol. Chem. 239, 3619 (1964). 108. Rändle, P . J., Denton, R. M., and England, P . J., in "Metabolic Roles of Citrate" (T. W. Goodwin, ed.), p. 87. Academic Press, New York, 1968. 109. Regen, D . M., Davis, W. W., Morgan, H . E., and Park, C. R., / . Biol. Chem. 239, 43 (1964). 110. Rozsa, K. S., Life Sci. 8, 229 (1969). 111. Rozsa, K. S., and Nagy, I. Zs., Comp. Biochem. Physiol. 23, 351 (1967). 112. Salas, M. L., Salas, J., and Sols, J., Biochem. Biophys. Res. Commun. 3 1 , 461 (1968). 118. Setlow, B., and Mansour, T. E., / . Biol. Chem. 245, 5524 (1970). 114. Setlow, B., and Mansour, T. E., Biochim. Biophys. Acta 258, 106 (1972). 115. Setlow, B., and Mansour, T. E., unpublished observation (1970). 116. Setlow, B., and Mansour, T. E., Biochemistry (in press) (1972). 117. Shen, L. C , Fall, L., Walton, G. M., and Atkinson, D. E., Biochemistry 7, 4041 (1968). 118. Singhal, R. L., Valadares, J. R. E., and Ling, G. M., J. Biol. Chem. 242, 2593 (1967). 119. Stadtman, E. R., in "The Enzymes" (P. Boyer, ed.), 3rd ed., Vol. 1, p. 425. Academic Press, New York, 1970. 120. Stone, D. B., and Mansour, T. E., Mol. Pharmacol. 3, 161 (1967). 121. Stone, D. B., and Mansour, T. E., Mol. Pharmacol. 3, 177 (1967). 122. Sutherland, E . W., Phosphorus Metab. 1, 53 (1951). 123. Sutherland, E . W., and Rail, T. W., J. Biol. Chem. 232, 1077 (1958). 124. Taylor, J. F., Phosphorus Metab. 1, 104 (1951). 125. Taylor, J. F., Antonini, E., Brunari, M., and Wyman, J., J. Biol. Chem. 241, 241 (1966).

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A Theoretical Background to the Use of Measured Concentrations of Intermediates in Study of the Control of Intermediary Metabolism I

F . S. ROLLESTON

I I I I

Banting and Best Department of Medical Research University of Toronto Toronto, Canada

I. Introduction II. Theoretical Background to the Study of Intermediary Metabolism and a General Approach A. The General Case Considered B. Disequilibrium and Metabolic Control C. Division between Nonequilibrium and Equilibrium Reactions . D. Regulatory Enzymes E. The Significance of Equilibrium Reactions to Metabolic Control F. Rate-Limiting Systems G. Relationships between Intermediate Concentrations and Kinetic Properties of Equilibrium and Nonequilibrium Reactions . . H. Summary III. The Crossover Theorem A. Development and Proof B. The Meaning of Interaction C. Application to Nonconserved Reactions D. The Fault Theorem IV. Application of the Principles Developed A. The Interconversion of Glucose and Fructose Diphosphate . B. Glyceraldehyde-3-Phosphate Dehydrogenase C. Citrate Synthase and the Control of the Tricarboxylic Acid Cycle References

47 49 49 50 51 53 55 55 56 57 58 58 60 62 62 64 64 68 69 73

I. Introduction Most of the pathways of intermediary metabolism are known, and their control mechanisms are now under active investigation. A four-stage ap­ proach to analysis of control mechanisms has been suggested by Newsholme (44, 46) · First, the enzymes involved in metabolic regulation are 47

48

F . S. ROLLESTON

identified ; second, their kinetic properties are investigated in detail ; third, the kinetic properties are used to postulate a hypothesis of metabolic control; and fourth, the hypothesis is tested and is modified as necessary. This paper is concerned with the first stage, the identification of those enzymes at which metabolic regulation is exerted. Three general methods of approaching this question have been suggested (18, 44, 46)· The teleological approach depends on the belief that control should occur early in the pathway and shortly after branch points in order to avoid long stretches of uncontrolled metabolism (31). Use of this approach requires no experiment, simply inspection of the pathway. A second ap­ proach is based on analysis of the properties of enzymes, on the premise that enzymes possessing such phenomena as changing maximal activities on dietary or hormonal manipulation, low maximal activities in com­ parison with other enzymes of the pathway, or activation or inhibition by other intermediates, are likely control sites. Use of this approach requires extensive experimental analysis, but derives its conclusions from experiments with enzymes extracted from tissues. There is no obligatory link between the properties observed in vitro and the conclusions pre­ sumed to apply in vivo, for this approach shows only that the enzyme has properties consistent with control, not that it actually does control metabolism. In the third method, intact tissue preparations are subjected to various treatments to give different rates of flow through the metabolic pathway and the steady-state concentrations of intermediates measured and used to deduce sites of metabolic control. This approach permits analysis of intact tissue preparations, and is potentially able to provide unequivocal identification of control sites. Since the development of the crossover theorem (7-9) measured con­ centrations of intermediates of metabolism have often been used to identify control sites. However, few workers have adhered rigidly to the theorem as originally stated, and the variations or shortened versions that have been substituted have not always been logically valid. Consequently, there is much confusion in the literature as to how measured concentra­ tions of intermediates can be used to identify control sites. This review is intended to discuss a theoretical background to this general topic. The presentation is divided into three sections. The first is an expansion of a brief survey carried out in 1966 (55) in which the general theory of application of measurements of concentrations of metabolic intermediates is discussed and a general approach is presented. The next section dis­ cusses the crossover theorem, and the last discusses specific problems in the control of intermediary metabolism in the light of the general theory developed.

49

INTERMEDIATE CONCENTRATIONS AND CONTROL SITES

II. Theoretical Background to the Study of Intermediary Metabolism and a General Approach A. The General Case Considered Figure la describes a generalized metabolic pathway in which a large, or constantly maintained, pool of substrate (A) is converted by a series of five reactions to an equally well maintained pool of product F. The rate of flow through this pathway can be controlled in three general ways: (1) by changing the concentrations of A or F, the overall substrates or products of the pathway; (2) by changing the concentrations of the cofactors of the pathway; or (3) by changing the activities of the en­ zymes involved in the pathway. Mechanisms of control involving changes in concentrations of substrates, products, or cofactors are likely to be secondary to controls operating on other pathways since there is exten­ sive sharing of cofactors among pathways and dependence of substrate supply or product removal on diet or on alterations of other pathways. However, control mechanisms involving changes in the activities of en­ zymes are more likely to involve primary responses specific to the path­ way. These could, for example, enable it to take a larger share of the available substrates and cofactors, rather than merely consuming those that are left by other pathways. The following discussion will be con­ cerned mainly with the identification of those reactions at which control of the activity of the enzyme has significant effect on the overall pathway. (a)

(b)

A / V B■ m n

100% -I

2

■C y ^ op

'D-^-^E—^—F

,

-AG

0%

J

I

FIG. 1. (a) A and F are large, or constantly maintained pools of substrate and product for the pathway. B, C, D, and E are all intermediates in the carbon sequence of the pathway and are called pathway substrates and products for their respective enzymes, m, n, o, and p are cofactors that participate in the overall reaction of the pathway and are referred to as cofactor intermediates, (b) Repre­ sentation of the distribution of — AG through the reactions of the pathway as a percentage of the total — AG.

50

F. S. ROLLESTON

For this discussion I have assumed that the concentrations of inter­ mediates as measured are relevant to the intracellular environment. Prob­ lems arising from comparution* of intermediates have recently been discussed by Sols and Marco (60) in a useful and thoughtful review, and will be briefly considered in Section IV. B. Disequilibrium and Metabolic Control A theoretical background to the investigation of control mechanisms can be formulated by considering the driving force for a metabolic path­ way. The general pathway in Fig. la catalyzes the net reaction A + m + o ^ F + n + p. Thermodynamically, this reaction can be de­ scribed by a mass action ratio (6, 21, 22) : r =

[F][n][p] [A][m][o]

At thermodynamic equilibrium, Γ is equal to Keq, the equilibrium con­ stant, and there can be no net flow. Keq is related to the standard free energy (AG°) across the reaction by AG° = - RT \n Keci. To obtain net flow through the pathway from A to F, the reaction must be displaced from equilibrium such that Γ is less than Keq. The extent of the displace­ ment from equilibrium determines the free energy (AG) across the reac­ tion, which can be expressed by the equation AG = AG° 4- RT In Γ = RT In (Γ/Χβ α ). Since (T/Keq) < 1, AG is always negative. Metabolic pathways usually operate under conditions in which —AG is large (21, 3Jf). This large free energy drop, or displacement from equilibrium, is generally distributed unevenly over the individual steps of the pathway such that some steps are closer to thermodynamic equi­ librium than others (5, 6, 21, 30, 42). This is illustrated in Fig. lb. In a dynamic steady state, there are only two obligatory restrictions on the free energy changes across each reaction: (1) that they add up together to the total free energy across the pathway, and (2) that they all be negative. Limited only by these two restrictions, the free energies across the individual reactions can fall anywhere on a spectrum of values. They depend only on the relationship of Γ to Keq and cannot be predicted with any certainty from kinetic constants of enzymes nor from the values oiKeq U ) . * The noun comparution is chosen as it is the least redundant of the three pos­ sibilities available; it is the abstract noun from the verb to compart (Shorter Oxford Dictionary). Of the other two possibilities, compartmentation is a noun ab­ stracted from another noun, whereas compartmentalization starts from a verb, proceeds through a noun to an adjective, thènce back to a verb, which finally is abstracted.

INTERMEDIATE CONCENTRATIONS AND CONTROL SITES

51

Control of metabolism through alteration of the activity of an enzyme must be exerted at reactions that are far from thermodynamic equilib­ rium (6, 21, 22, 30, Iß, 55). The argument advanced in support of this statement is that if the enzyme has enough activity to allow its reactants to approach equilibrium and therefore to catalyze the reverse reaction at almost the same rate as the forward reaction in spite of flow through the pathway, then it cannot be limiting flow through the pathway (6, 21, 30, 33). A similar conclusion is reached if the extent to which alteration of the activity of one particular enzyme can influence the remainder of the pathway is analyzed in terms of the free energy across the reaction. The effects of such an alteration of activity must be transmitted to the re­ maining enzymes through changes in the concentrations of pathway intermediates. In a dynamic steady state, activation of an enzyme by an outside agent will tend to bring the reaction closer to equilibrium. This will result from tendency for the steady state concentration of sub­ strates to decrease and for those of the products to increase (8, 9). Since AG must remain negative, the size of these changes is limited by the initial value of AG. Thus the efficiency with which activation of an enzyme can influence the overall rate of flow through the pathway is directly related to the extent to which the information that this activa­ tion has taken place can be conveyed to the other reactions. This, in turn, is directly related to the degree of displacement of that reaction from equilibrium before the activation occurred.* From the above, it follows that alteration of the activity of a reaction in a metabolic pathway will have more influence on the pathway if that reaction is initially far from equilibrium (6, 21, 22, 30, 4$, 55). This suggests that the first step in analysis of mechanism of control is to classify the reactions on the basis of their displacement from equilibrium. C. Division between Nonequilibrium and Equilibrium Reactions Subsequent discussion has been simplified by considering only two classes of reaction, those that are close to thermodynamic equilibrium (equilibrium reactions), and those that are far from thermodynamic equilibrium (nonequilibrium reactions) (6, 21, 55). While classification into these two groups is somewhat arbitrary, the point of division can be assessed by evaluating the extent to which the rate of flow through an enzyme-catalyzed reaction is dependent on the value of — AG (10). The rate of an enzyme-catalyzed reaction can be regarded as the sum of its * This argument is harder to apply if it is assumed that the enzyme is inhibited, for provision must then be made against complete inhibition of the reaction, which would cause 100% of the AG across the pathway to occur at that reaction.

52

F. S. ROLLESTON

forward and reverse components. Near equilibrium, the rate of the reverse reaction (υ_ι) is almost as great as the rate of the forward reac­ tion (υ+ι) and the net rate of flow is low compared with the capacity of the enzyme to catalyze either reaction. However, far from equilibrium, the forward reaction dominates and the reverse reaction is of negligible significance to the overall rate of flow. The rate of the reverse reaction can be regarded as negligible when less than 5% of the rate of forward reaction, and to play a large part in determination of the overall rate of flow through the reaction if it is greater than 20% of that of the forward reaction (Table I ) . Application of the relationship υ_ι/υ+ι = T/Keq (22) leads to a range of values of T/Keq (0.05-0.2), that separates nonequilibrium from equilibrium reactions. The suggestion that a range of values of T/Ke(l from 0.05 to 0.2 serves to separate nonequilibrium from equilibrium reactions leaves open the question of how those falling within this range are to be regarded with respect to their potential as control sites. Activation of such reac­ tions could exert significant control over the pathway as up to 20-fold increases in Γ could be tolerated by the system without reversing the flow, and the effects of such activation could therefore be transmitted to the remainder of the pathway. Conversely, the flow through the en­ zyme is sensitive to changes in Γ caused by changes in product con­ centration. It is therefore clear that the range of values of T/Keq that separates equilibrium and nonequilibrium enzymes is determined in part by what one regards as a significant effect on the pathway, and that reactions falling on the borderline between these two categories will share the control characteristics of both. The definition of T/Keq in terms of υ_ι/υ+1 (22) also provides the basis for the conclusion that, for nonequilibrium reactions, changes in the rate of the reverse reaction, υ_1? have no necessary influence on the rate of flow. Since υ_ι is primarily determined by the concentration of product for the reaction, changes in product concentration have no necessary relationship to flow rates, and therefore cannot be used to infer changes in the activity of nonequilibrium enzymes. Nonequilibrium reactions, therefore, provide the points at which con­ trol mechanisms involving alteration of the activities of one enzyme of the pathway would efficiently influence the whole pathway. Such reactions can be identified by measurement of steady-state concentrations of all participants of the reaction, calculation of the mass action ratio, and comparison with the equilibrium constant (6, 21, 42, 55, 56, 69, 70). Identification of the nonequilibrium reactions can also be based on cor­ relations between changes in Γ and changes in the rate of flow, v. If, in comparing the effects of two different treatments of a tissue, it is observed that υι < v2 and I \ < Γ2 then, at least under the condition giving the

INTERMEDIATE CONCENTRATIONS AND CONTROL SITES

53

TABLE I RELATION BETWEEN T/Ke/Vmax — v) = n log S 4- log K, where v is the initial velocity, F m a x is the maximal velocity, K is the apparent overall disso­ ciation constant, and n is a coefficient which is a function of the number of binding sites per molecule of enzyme and of the strength of the coop­ erative interactions between sites. When interactions are strong, n will be numerically equal to the number of sites. Regardless of the number of sites, n must decrease to a value of 1, if intersite interactions are weakened (3). A value of n = 1 was repeatedly obtained by Caskey et al. (10) with pigeon liver enzyme studied in the standard assay. With enzyme preincu­ bated with PP-ribose-P and Mg2+, and studied at concentrations of glutamine of 4 times [S] 0 . 5 , values for n of 1.7 to 3.2 were found by Rowe et al. (71, 84). Although in one experiment the Hill coefficient decreased in a stepwise manner in the presence of increasing concentrations of AMP, in two other studies AMP produced no significant change (Table I I ) . A value for n of 1.9 was reported by D. L. Hill and Bennett (34)

154

J A M E S B. WYNGAARDEN

£

«5-H

1/5

FIG. 6. Competitive inhibition of PP-ribose-P amidotransferase by purine ribonucleo tides. (A) Enzyme prepared according to Wyngaarden and Ashton, frac­ tion V (86), not preincubated with substrate. Nucleotide concentration 0.2 milf. (B) Enzyme prepared according to Rowe and Wyngaarden, fraction 5 (72), pre­ incubated 15 minutes with PP-ribose-P and Mg2+. v = Δ ÒDxsesnm/lO minutes. S = mM. Figure 6A from Wyngaarden and Ashton (86) by permission of American Society of Biological Chemists.

for the enzyme for adenocarcinoma cells studied at two concentrations of glutamine (approximately two and three times greater than the Km value of glutamine). A value of 1.5 was found by Shiio and Ishii {74a) with enzyme from B. subtilis studied in the presence of AMP. Values of n increasing from 1.7 to 2.5 with increases of glutamine (from concentra­ tions nearly equal to Km values to concentrations six times greater) were

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

155

TABLE II PP-RlBOSE-P AMIDOTRANSFERASE : HlLL COEFFICIENTS"

Coefficient AMP (ml)

n

n

n

0 0.5 1.0 2.0

3.2 2.5 1.7 1.4

1.7 1.7

2.3

—

1.8

—

2.2 2.2

β Enzyme was preincubated 15 minutes with PP-ribose-P and Mg2+, then assayed at 5 levels of PP-ribose-P, in the presence of several concentrations of AMP (71, 84).

reported by Nagy for the enzyme from S. pombe {59). Values of n as high as 3 were found with a mutant form of the enzyme from S. pombe (aza-I) {59) (see Section IV, B, 5, a), and values of "effectively" 4 with the enzyme in extracts of lymphoblasts grown in culture {82a) (Table I I I ) . d. Ligand-Induced Conformational Changes. In a preceding section (II, C, 3), it was pointed out that the lag phase observed after the addi­ tion of reactants was quite variable with different preparations of pigeon liver enzyme, and could be appreciably shortened by prior incubation with PP-ribose-P and Mg2+. Preincubation also appears to result in changes in the enzyme molecule which lead to strengthening of coopera­ tive interactions between PP-ribose-P binding sites, as reflected in values of n > 1. The activation process has been interpreted as representing a sequential progression of ligand-induced conformational changes {71); this conclusion is based upon changing responses to inhibitors such as 1,10-o-phenanthroline and differing results in studies with the fluorescent probe, 2-p-toluidinylnaphthaline-6-sulfonate (TNS). This aspect of the behavior of glutamine PP-ribose-P amidotransferase will be discussed more extensively below, under Functional and Conformational Hetero­ geneity (Section IV, C). 2. GLUTAMINE

a. Michaelis-Menten Plots. Velocity-substrate plots form typical hyperbolic curves for glutamine with all enzymes studied, irrespective of the concentration of PP-ribose-P {10, 59, 62, 86). b. Lineweaver-Burk Plots. Double reciprocal plots are linear, except under two conditions: at very low concentrations of glutamine there is slight upward curvilinearity with the pigeon liver enzyme {10), and in the presence of I M P the otherwise linear plot becomes markedly curvi-

10, 53, 84

3.4 4.8

— — — 71,84

— — — — — — — — — — —

1

rì

rì 2.0 3.2 1.9 1.6 1.3 2.3 2.5

3.2 1.0

S4

— —

1.0 1.0 1.6 1.6

— — — — — —

rì

—

1.9

n

n 1.0 1.0

n

Pigeon liver a

Pigeon liver

° Preincubated enzyme; see Table II.

Reference

Inhibitor AMP ADP ATP GMP GDP IMP 6-MP-RP 6-MeThioG-RP 6-ThioG-RP dGDP AMP + IMP AMP + 6-MP-RP

Substrate PP-ribose-P Glutamine

Source :

Mouse adenocarcinoma 755

82a

— — — — — — — — — — — —

rì

~4 ~2

n

Human lymphoblast

74a

— — — — — — — — — —

3.1 2.5

rì

—

1.6

n

B. subtilis

P P - R I B O S E - P AMIDOTRANSFERASE : H I L L COEFFICIENTS

TABLE III

— — — — — —

— — — .— — — 69

59

1

—

— 2.5

— — — 1

rì

3 1

n

S. pombe (aza-1)

— — — 2.5

rì

2.5 2.2

n

Sçhizosaccharomyces pombe (wild)

Oi Ci

J A M E S B. WYNGAARDEN

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

157

linear in studies with enzyme from S. pombe {59). With enzyme from chicken liver {22), pigeon liver {10, 86), adenocarcinoma 755 cells {34), or B. subtilis {74a), the curve is displaced in the presence of inhibitors such as nucleotides {10, 74a, 86) or DON {22, 34), but it remains linear over the range of concentrations of glutamine studied. c. Hill Plots. Values of n = 1 would be anticipated for substrate ranges in which Lineweaver-Burk plots are linear, and indeed are ob­ tained. With pigeon liver enzyme the value of n is 1 with preincubated as well as with standard enzyme, and is unchanged in the presence of concentrations of AMP up to 2 m l {84)- A Hill coefficient of about 2 is found with enzyme in extracts of human lymphoblasts grown in cul­ ture {82a). With S. pombe enzyme, the value of 1 increases to 1.8 and 2.2 in the presence of 0.2 or 0.4 m ¥ IMP, indicating cooperativity of glutamine binding under these circumstances (Table III) {59). d. Ligand-Induced Conformational Changes. The enzyme previously activated by PP-ribose-P and Mg2+ is still not capable of achieving an immediately maximal velocity upon addition of glutamine (Fig. 1). The final activation induced by glutamine modifies the fluorescence response of the enzyme-PP-ribose-P—Mg 2+ complex upon addition of TNS (see Section IV, C). B. Effects of Purine Ribonucleotide Inhibitors 1. INHIBITION BY INDIVIDUAL NUCLEOTIDES

The pigeon liver enzyme is inhibited by purine 5'-ribonucleotides, but not by (2',3') ribonucleotides, 5'-deoxyribonucleotides, ribonucleosides or free bases, nor by pyrimidine compounds. Effective inhibitors include AMP, ADP, GMP, GDP, IMP, 4-amino-5-imidazolecarboxamide ribo­ nucleotide, and ATP, but not GTP, IDP, or I T P {86). In addition, the enzyme is inhibited by 5'-phosphoribosyl derivatives of 6-thiopurine, 6-thioguanine, 8-azaguanine, and allopurinol {53), and by the corre­ sponding derivative of cordycepin {70). Studies with PP-ribose-P amidotransferase from other sources have disclosed many similarities and some interesting differences. The en­ zyme from A. aerogenes is inhibited by AMP, ADP, GMP, and IMP. It is also inhibited by GTP but not by ATP {62). The enzyme from B. subtilis is weakly inhibited by GMP and GTP {56, 74a), more strongly by AMP and ADP {70, 74a) and 5'-phosphoribosyl cordycepin {70). The amidotransferase from S. pombe is sensitive to GMP and I M P but less well inhibited by AMP {59). PP-ribose-P amidotransferase from rat liver is inhibited by GMP, AMP, and ATP, and the kinetics of competition are like those of the pigeon liver enzyme {10). The P P -

158

J A M E S B. WYNGAARDEN

ribose-P amidotransferase of mouse spleen, induced by Friend leukemia virus, appears to be more sensitive to guanyl ribonucleotides than to adenyl compounds, but has only been studied in crude extracts (67, 68). The enzyme from adenocarcinoma 755 cells is sensitive to some extent to every nucleotide tested. Inhibition by ribonucleoside and deoxyribonucleoside triphosphates can be overcome by additional magnesium (84) · This is not the case with inhibition of the pigeon liver enzyme by ATP (86). Some nucleotides were only moderately inhibitory even at very high concentrations. The most strongly inhibitory of the natural nucleo­ tides tested was dGDP. 6-Methylthiopurine ribonucleotide was the most potent inhibitor of all the nucleotides tested (84)- Enzyme from human lymphoblasts is inhibited by AMP and GMP (82a). With all enzymes studied, suitable concentrations of effective ribonucleotides produce 100% inhibition (10, 84, 58, 59, 62, 70, 74a, 86). a. Michaelis-Menten Plots. With both S. pombe (59) and preincubated pigeon liver enzymes (71), the sigmoidicity of the plot of velocity as a function of PP-ribose-P concentration is more readily apparent in the presence of nucleotide inhibitor, but the maximal velocity appears to be unchanged. b. Lineweaver-Burk Plots. With pigeon liver enzyme not preincubated with PP-ribose-P and Mg2+, double reciprocal plots show changes of slope but no changes in F max in the presence of inhibitors, when PPribose-P is the substrate whose concentration is varied (10, 58, 86). These results describe an apparent competitive or "i£-type" inhibition with respect to PP-ribose-P. When glutamine is the substrate whose concen­ tration is varied, there is both an apparent increase in Km and a decrease in Fmax. These results describe a mixed competitive-noncompetitive or "mixed K-V type" inhibition (58) with respect to glutamine (10, 58). With enzyme from S. pombe the effects of inhibitor (IMP) appear to be more strictly competitive with respect to glutamine (59). The competi­ tive kinetics of inhibition suggested initially that purine ribonucleotides might compete for the substrate site at which PP-ribose-P is bound. The effects of ribonucleotides upon enzymes that were desensitized to their inhibitory .effects (see below) indicated that this interpretation was not correct and that the enzyme was "allosteric," i.e., that special regu­ latory sites existed on the enzyme (58). c. Hill Plots. Cooperativity between inhibitor binding sites may be evaluated by use of the equation, log v/V0 — v) = log K' — n' log I, where v is the initial velocity in the absence of inhibitor, nf is a coeffi­ cient which is a function of the number of inhibitor binding sites per en­ zyme molecule and of the strength of cooperative interactions between sites, I is the concentration of inhibitor, and K! is the apparent overall

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

159

dissociation constant (36, 57, 77). With pigeon liver enzyme (10) in the tetramer form (72), Hill plots are sigmoidal and yield values of rì ranging from 1.3 to 3.2 for AMP, ADP, ATP, GMP, GDP, IMP, and 5-phosphoribosyl-6-mercaptopurine at [I] 0 .s concentrations (Table I I I ) . A curious finding is that pigeon liver enzyme which has been preincubated for 15 minutes with PP-ribose-P and Mg2+, and which now shows a shortened lag period, sigmoidal kinetics and n values for PP-ribose-P of 1.7 to 3.2, no longer shows cooperativity between AMP sites. Values of rì = 1 were found at substrate concentrations of PP-ribose-P of 0.1, 0.25, or 0.5 mikf (84). The amidotransferase from B. subtilis (74a) has n values of 3.1 for AMP and 2.5 for ADP. With enzyme from S. pombe (59), rì values for I M P and GMP were 2.5 at saturating values of PP-ribose-P and 1.3 at [S] 0 . 5 concentrations of PP-ribose-P. With en­ zyme from adenocarcinoma 755 cells (34), rì values were independent of PP-ribose-P concentrations. For 6-thioguanine ribonucleotide and for dGDP, rì was 1.6 ± 0.2, but for 5-phosphoribosyl derivations of 6-mercaptopurine and 6-methylthiopurine the values were 1.0 ± 0.1 or 0.2. Kiy and [I]o.s values for various inhibitors obtained from Lineweaver-Burk or Hill plots, respectively, are listed in Table IV. d. Effect of Inhibitors on Substrate Binding. With enzyme from S. pombe (59) or with preincubated enzyme from pigeon liver (71), nucleotide inhibitors lower the affinity of enzyme for substrates, but do not modify the cooperativity of PP-ribose-P binding. The values of n remain unchanged (84)- With enzyme from S. pombe, the glutamine concentra­ tion-velocity curve, which is strictly hyperbolic in the absence of IMP, becomes strongly sigmoidal, and n values for cooperativity of glutamine binding increase from 1.0 to 2.2 (59). 2. INHIBITION BY COMBINATIONS OF NUCLEOTIDES

a. Studies with Partially Purified Preparations of PP-ribose-P Amidotransferase. In initial studies of end-product inhibition of PP-ribose-P amidotransferase from pigeon liver, Caskey et al. (10) encountered rather bizarre variations in nucleotide sensitivity of certain preparations. In some instances the amidotransferase lost sensitivity to AMP and ATP while standing at 4°C and regained ATP sensitivity after re freez­ ing. Eventually several preparations lost all sensitivity to ATP while undergoing as much as an 8-fold gain in specific activity. In other ex­ periments enzyme preparations lost and regained sensitivities to AMP and GMP, but the variations with respect to the two nucleotides oc­ curred independently. The lack of parallelism of responses to AMP and GMP suggested the possibility of more than one type of inhibitor binding site. A series of

10,86

— — — — — — — —

0.09-2.5 0.039-0.64 0.037-1.1 0.18-3.5 0.086-1.0 0.38-5.4

Ki

53

— —

0.05 0.19 0.54 0.6

— — — — — — —

1.0

Ki

70

—

0.7

— — — — — — — — — — —

0.7

Ki

VALUES0

u

0.09

0.65

1.8 0.4

> 6 ..7* > 3 .. 3 * > 3 .. 3 * > 2 ..3* 1..2 1.1 0.27 >3.3*

[I]0.5

0.55 1.3

[I]0.5

Aerobacter aerogenes

>2.3* >3.3*

[I]0.5

Mouse adeno carcinoma Human 755 lymphoblast

A N D [ I ] 0. 5

70

0.1

[I]0.5

Bacillus subtilis

69

0.03

[I]0.5

Schizosaccharomyces pombe

a Ki values (mM) determined from Lineweaver-Burk plots. [I]o.5 values (mM) determined from Hill plots, except that those marked with an asterisk (*) were estimated from tabulated data. b R P = ribonucleoside 5'-monophosphate.

Reference

Inhibitor AMP ADP ATP IMP GMP GDP GTP dGDP 6-Mercaptopurine-RP 6 6-Thioguanine-RP 8-Azaguanine-RP Allopurinol-RP Cordycepin-RP 6-Methylthiopurine-RP

Source Ki or [I]o.5

Pigeon liver

P P - R i B O S E - P A M I D O T R A N S F E R A S E : KÌ

TABLE IV

o

J A M E S B. WYNGAARDEN

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

161

studies was therefore conducted with different pairs of ribonucleotide in­ hibitors in which the effects of each ribonucleotide alone were compared with inhibitions caused by two ribonucleotides acting together. It was reasoned that if a single species of regulatory site was involved, mixtures of two inhibitors should have additive effects, whereas if there were two or more types of sites capable of interacting, the effects of two inhibitors acting simultaneously should be greater than the sum of the individual contributions. In the cases of AMP plus ADP, GMP plus IMP, GMP plus GDP, and AMP plus ATP, the residual fractional activity in the presence of the two nucleotides acting together was not less than the product of the residual fractional activities observed with an equal total nucleotide concentration of each inhibitor alone. However, in the cases of AMP plus GMP, AMP plus IMP, AMP plus GDP, and AMP plus 6-thiopurine ribonucleotide, the effects of the two inhibitors acting to­ gether were significantly greater than the predicted additive effects (Fig. 7) (10). In Hill plots, values of ri as high as 4.8 were obtained at total concentrations of two nonhomologous inhibitors (AMP plus 6-mercaptopurine ribonucleotide) which reduced the velocity by 50% (Table HI) (84). These results cannot be explained by the hypothesis of binding of these nucleotide pairs to identical subgroups of a single type of inhibitor site, and they indicate that the pigeon liver enzyme contains either separate binding sites for 6-amino and 6-hydroxypurine ribonucleotide compounds or exclusive binding subgroups for each type of inhibitor within a single type of site, and that these sites are interacting when occupied by nonhomologous inhibitors. Ribonucleotides that are nonhomologous at the 6-position can be bound simultaneously, and binding of one inhibitor either increases the affinity for the second, or it increases the inhibitory effect of binding of the second (10). This type of allosteric regulation was initially termed "cooperative feedback inhibition" (10, 75). This term is ambiguous in this context, however, in that many regulatory en­ zymes show cooperative effects of homotropic or heterotropic type but do show potentiative effects in the presence of mixtures of inhibitors. The term "synergistic inhibition" (76) more appropriately describes the phenomenon observed with the amidotransferase in the presence of mix­ tures of 6-amino and 6-hydroxypurine ribonucleotides and is now the preferred designation. Very similar control features have been observed by Nierlich and Magasanik (62) with the PP-ribose-P amidotransferase of A. aerogenes which also shows synergistic inhibitions by 6-hydroxy- and 6-aminopurine compounds. The synergistic inhibition is competitive and can be completely overcome by high concentrations of PP-ribose-P. The P P -

162

JAMES B. WYNGAARDEN

FIG. 7. Synergistic inhibition of PP-ribose-P amidotransferase by pairs of nonhomologous purine ribonucleotides. I n all four frames, the observed activity values in the presence of two nucleotides are plotted in terms of total nucleotide concen­ trations, and the two inhibitors are present in equimolar concentrations. The predicted curve is determined by the product of the fractional inhibitions caused by each inhibitor alone. For example, in the upper left-hand panel, there was 57% residual activity in the presence of 1 m M ADP, and 50% residual activity in the presence of 1 m M A M P . The predicted activity in the presence of 1 m M A D P and 1 m l A M P is (0.57 X 0.50) 100 or 28.5%. Thus the predicted residual activity curve represents the minimal activity (maximal inhibition) anticipated if each inhibitor acts independently of the other. Observed inhibitions greater than those found with equimolar total concentration of either inhibitor alone, and greater than the individual inhibitions (predicted residual activity curves) represent synergistic effects of two inhibitors. As shown here, A M P + G M P (lower left-hand panel) and A M P + I M P (lower right-hand panel) act synergistically. The velocities in the absence of inhibitor were 0.023-0.030 optical density unit/minute. From Caskey et al. (10) by permission of the American Society of Biological Chemists.

ribose-P amidotransferase of S. pombe {59) also shows inhibitory effects of equimolar mixtures of AMP plus IMP, and of AMP plus GMP, that are greater than the calculated additive effects, but the synergism is minimal, perhaps owing to the relative insensitivity of the S. pombe en­ zyme to AMP. It is reported {34) that PP-ribose-P amidotransferase

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

163

preparations from adenocarcinoma 755 cells do not show synergistic ef­ fects of combinations of inhibitory ribonucleotides. The pigeon liver enzyme appears to be the only one studied thus far that is inhibited by ATP even in the presence of excess magnesium (86). Independent spontaneous variations in sensitivity to ATP and AMP have been mentioned above. In addition, many preparations which showed high grade sensitivity to AMP and GMP were totally insensitive to ATP. However, in enzymes sensitive to both AMP and ATP, the ef­ fects of both inhibitors acting together were additive (10). These effects can be accommodated within the hypothesis of a single site for 6-aminopurine compounds, if it is assumed that different subgroups are involved in binding of AMP and ATP, which possess different binding energies in different conformational states. The synergistic nature of the inhibitions by 6-amino- and 6-hydroxypurine ribonucleotides on the first step in their biosynthesis should per­ mit the more effective curtailment of purine biosynthesis when both types of inhibitors are in surplus simultaneously, but allow for a more moder­ ate control when only one kind of purine is present in excess. b. Studies in Intact Cells. In studies of feedback inhibition of purine biosynthesis in azaserine-treated ascites tumor cells, Henderson (25) found that purine bases inhibited synthesis of α-iV-formyl glycinamide ribonucleotide (FGAR). By analogy with the effects of purine ribo­ nucleotides on purified PP-ribose-P amidotransferase of pigeon liver, these effects were assumed to be dependent upon prior conversion of base to ribonucleotide. Mutant strains of tumor cells which lack hypoxanthine-guanine phosphoribosyltransferase are resistant to inhibition of purine biosynthesis by hypoxanthine analogs (8, 44)· Combinations of weakly inhibitory concentrations of adenine and guanine, adenine and AIC (25), adenine and 2,6-diaminopurine, adenine and purine, or of 2,6diaminopurine and purine (26) exerted stronger inhibitory effects than even high concentrations of each compound alone. These latter observa­ tions were without explanation at the time. In subsequent studies, Henderson and Khoo (30) found that both glutamine-dependent and NH 4 Cl-dependent utilization of PP-ribose-P were inhibited by purine analogs which by themselves had little or no effect upon PP-ribose-P concentrations. Methylthioinosine (a substi­ tuted 6-hydroxypurine) inhibited PP-ribose-P disappearance only when glutamine was the second substrate, whereas psicofuranine (a 6-aminopurine) was exclusively active when NH4C1 was used. Apparently these inhibitors bind to the enzyme at different sites. Psicofuranine potentiated the inhibitory effects of methylthioinosine, suggesting that it was bound even when its effects were not independently apparent. These studies

164

JAMES B. WYNGAARDEN

were considered confirmatory in the intact cell of concepts developed with partially purified enzymes from other sources. 3. SENSITIVITY OF SUBUNITS TO RIBONUCLEOTIDE INHIBITORS

Limited studies of the three enzyme fractions obtained from Bio-Gel P-300, or from sucrose density gradient separations, disclosed inhibition of all fractions by AMP and GMP, tested individually (71). Approxi­ mate [I]o.5 values of 10~3M were observed for each nucleotide for all three fractions. Other studies reviewed above provided no evidence for reaggregation of monomer and dimer under conditions of the assay em­ ployed. These inhibition studies therefore suggest that PP-ribose-P amidotransferase is both active as enzyme and sensitive to nucleotides in tetramer, dimer and monomer states. No studies of cooperativity of homologous or nonhomologous inhibitor binding have been conducted with dimers or monomers as yet. 4. DESENSITIZATION OF P P - R I B O S E - P AMIDOTRANSFERASE TO INHIBITORS

Sensitivity to nucleotide inhibitors may change in the course of en­ zyme purification. Some preparations display no sensitivity whatever. With pigeon liver enzyme (10) desensitization occurred most often dur­ ing dialysis of an ammonium sulfate fraction against distilled water. In this procedure the enzyme precipitates, and on solution in buffer, the en­ zyme, although catalytically active, is frequently nucleotide insensitive. A revision of the purification procedure was developed (72) in which the ammonium sulfate fraction is applied directly to a Sephadex column, as. described above. With this procedure desensitization occurs only rarely, and an enzyme with reproducible properties is obtained, but K\ values for inhibitors are higher than those observed with the most sensi­ tive preparations obtained with the earlier procedure (86). The sum total of experience gives a wide range of Ki or [I]0.5 values for nucleotides even with sensitive pigeon liver enzyme (Table IV). Desensitization of more highly purified enzyme to inhibitors sometimes occurred during study in the absence of additional purification procedures. Examples of unusual variation in sensitivity to AMP and ATP were cited above. The 8-fold gain in specific activity that accompanied total loss of sensitivity to ATP and diminished response to AMP of an enzyme preparation dur­ ing freezing and thawing, strongly suggests that spontaneous conformational changes were responsible for these variations in properties. Never­ theless, deliberate desensitization of nucleotide-sensitive enzyme could not be achieved by measures which have been successfully employed with other allosteric enzymes, such as treatment with heat, metabolic

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

165

inhibitors, or urea. In fact, urea increases the sensitivity of enzyme to A M P , A D P o r A T P (10). Results with PP-ribose-P amidotransferases from other sources re­ capitulate the full range of experiences with the pigeon liver enzyme. The highly purified enzyme from chicken liver prepared by Hartman (21) is insensitive to nucleotides. The procedure employed includes a dialysis step during which the enzyme precipitates. The amidotransferase from S. pombe retains its activity and its sensitivity toward inhibitors for about a week when stored at 0°, but loses sensitivity toward inhibi­ tors in 2-3 days at — 20°C (59). The enzymes from A. aerogenes (##), B. subtilis (70), and adenocarcinoma 755 cells (34) apparently retain nucleotide sensitivity during partial purification. An interesting observation with the pigeon liver enzyme is that even the nucleotide-insensitive PP-ribose-P amidotransferase is protected against inhibition by 1,10-o-phenanthroline by preincubation with AMP plus Mg2+, or AMP plus Mg2+, but not by nucleotide or Mg2+ alone (71). These results suggest that the nucleotide-insensitive enzyme (in State II form, see below) is still capable of binding AMP or GMP. In this respect the enzyme appears to be similar to the first enzyme of histidine biosyn­ thesis, phosphoribosyl-ATP-pyrophosphorylase, in which Martin (50) has shown by equilibrium dialysis studies that insensitive enzyme binds the "inhibitor" histidine. 5. EFFECTS OF MUTATIONS

a. Schizosaccharomyces pombe (aza-I) (59). This mutant was se­ lected for its resistance to 8-azaguanine. Aza-I grows normally on mini­ mal medium and excretes hypoxanthine and inosine. The aza-I mutation is closely linked to the structural gene ad-4 of glutamine phosphoribosylpyrophosphate amidotransferase. The purified amidotransferase from mutant strain aza-I shows changes in both substrate binding and effects of inhibitors. The [S] 0 . 5 value for PP-ribose-P is 1 X 10~4M, three times lower than that of enzyme from the wild strain. The value of n increased from 2.0 to 3.0 with increases Qf glutamine concentration from 4 to 20 mM. The [S] 0 . 5 value for glutamine was 5.5 X 10~3M, and was now in­ dependent of PP-ribose-P concentration. The inhibition curve for AMP did not differ significantly from that of the wild-type enzyme, but the [I]0.5 value for I M P and GMP was about 10 times higher. Furthermore, there was no cooperativity between I M P (or GMP) binding sites of the mutant enzyme, the ri value of the Hill plots being 1. Two other differ­ ences were found: total inhibition by I M P could not be achieved even at I M P concentrations as high as 8 mikf, and no stimulation of activity was found at low concentrations of I M P such as was observed with en-

166

JAMES B. WYNGAARDEN

zyme from wild-type cells. In addition, IMP lowered the cooperativity of PP-ribose-P binding (from n values of 2.8-1.5) and failed to induce cooperativity of glutamine binding with enzyme from the mutant strain. Thus, the mutational event conferring resistance toward 8-azaguanine seems to result in an alteration of at least some of the IMP and GMP binding sites of PP-ribose-P amidotransferase, which also prevents the cooperativity of IMP binding and modifies the patterns of substrate binding. fa. Ehrlich Ascites Tumor Cells. Henderson et al. (29) have described a mutant strain of ascites cells which is resistant to 6-methylmercaptopurine, a compound that is converted to its ribonucleoside by purine ribonucleoside phosphorylase, and is then converted to its ribonucleotide by adenosine kinase. The mutant cells make the ribonucleotide deriva­ tive at a normal rate, but FGAR synthesis in azaserine-treated cells is less well suppressed by 6-methylmercaptopuririe and by guanine than in normal cell, whereas suppression of FGAR synthesis by adenine is normal. Glutamine-dependent diappearance of PP-ribose-P is also less well suppressed by the analog (presumably its ribonucleotide) in the mutant than the wild-type cell. The authors propose that the mutant cells contain an altered amidotransferase which exhibits reduced sensi­ tivity to 6-hydroxypurine ribonucleotides, and normal sensitivity to 6-aminopurine ribonucleotides. c. Human "Fibroblasts." Cells with fibroblast morphology obtained by tissue culture of punch biopsies of human skin are capable of purine synthesis de novo and are useful for study of biochemical abnormalities of this process. In the presence of azaserine they accumulate FGAR, and the addition of purine bases results in inhibition of purine biosyn­ thesis de novo and reduction in the amount of FGAR formed. Cultured fibroblasts from two gouty subjects, both flamboyant overproducers of uric acid, have been found by Henderson and colleagues (31) to show reduced feedback inhibition in response to adenine, hypoxanthine, and 6-methylmercaptopurine ribonucleoside. The extent of conversion of purine bases to nucleotides, and of the interconversion of ribonucleotides, was apparently normal in these cells. The possibility of an altered P P ribose-P amidotransferase is discussed by the authors, who also point out, however, that both patients responded normally to feedback inhibi­ tion of purine biosynthesis by azathiopurine in vivo. C. Functional and Conformational Heterogeneity of PP-Ribose-P Amidotransferase from Pigeon Liver The lag phase that precedes development of maximal velocity upon addition of substrates, and the variable sensitivities of enzyme to chelat-

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

167

ing agents and to nucleotides, have indicated that the enzyme may exist in a range of conformational states in which it exhibits varied responses to substrates and modifiers. With respect to catalytic activity, Rowe et al. (71) have demonstrated four major forms of the enzyme, which may of course be present in mixtures in some preparations: (a) enzyme that is initially inactive in the presence of substrate, but is capable of being activated, referred to as State I enzyme; (b) enzyme, initially in­ active, which after maximal activation by PP-ribose-P and Mg2+ cata­ lyzes a reaction upon addition of glutamine which exhibits only a short lag phase before attaining maximal velocity, referred to as State II en­ zyme; (c) enzyme which demonstrates an initial high rate of activity, albeit not maximal, in the assay system without preincubation, but which differs in other ways from State II enzyme, referred to as State IV enzyme; (d) enzyme under conditions where all substrates are pres­ ent and the reaction is proceeding maximally, referred to as State III enzyme. The transition of the State I enzyme into State II form is strongly inhibited by the iron chelators, implying that the iron moieties essential to catalytic activity are in an exposed or accessible position in the State I molecule. Incubation of the State I enzyme with PP-ribose-P and Mg-+ results in protection of these iron atoms, implying that the conforma­ tional changes leading to activation to State II also render the iron in­ accessible to chelators. AMP and MgL>+ will also block chelator effects in the State I enzyme provided the enzyme is sensitive to inhibition by the nucleotide. The nucleotide-insensitive State I enzyme either does not bind the nucleotide or the nucleotide binding site is in such a position that even though filled it cannot afford protection to the active site against the effects of the chelator. The State II enzyme becomes fully active only after the addition of glutamine. The progressive acceleration during the course of the reaction implies a further conformational change following the binding of this second substrate. The fully activated or State I I I molecule is totally resistant to chelators; apparently the iron is now inaccessible to the relatively bulky o-phenanthroline molecule. Kinetic studies employing substrate exhaustion and substrate removal techniques indicate that the postulated conformational changes of the activation process are reversible (Fig. 8). When levels of PP-ribose-P become limiting during the reaction, the enzyme, initially added in State I form, reverts from State III to a mixture of State I and State II forms, indicating the necessity of PP-ribose-P for the binding of glutamine. If PP-ribose-P is exhausted or totally removed from the reaction, a com­ plete reversal to State I occurs. Removal or depletion of glutamine from

168

JAMES B. WYNGAARDEN

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FIG. 8. Kinetic curves illustrating the effect of substrate limitation and sub­ strate depletion on various forms of the enzyme. Curve A represents the effect of the addition of further PP-ribose-P to enzyme initially added in State I I ' form when this substrate is becoming limiting. Curve B is the result of a similar experi­ ment with enzyme initially in State I or State I I form. The results of a similar experiment when PP-ribose-P has been exhausted are shown with curve C (State I F enzyme) and curve D (State I or State II enzyme). Note the time scale indi­ cating that the velocities to the left are not initial velocities. Arrows indicate the point of substrate addition. From Rowe et al. (71) by permission of the American Chemical Society.

fully activated (State III) enzyme causes a reversion to State II pro­ vided PP-ribose-P and Mg2+ are present. The reversal of the changes that characterize the activation process is accompanied by appropriate change in sensitivity to o-phenanthroline. The State I F form appears to be different from the State II form in that it does not require PP-ribose-P and Mg2+ for the maintenance of its activated state, and removal of these substrates does not cause an im­ mediate transition to State I form. The enzyme is more resistant to o-phenanthroline and is fully protected by AMP plus Mg2+. This implies that, irrespective of the nucleotide inhibition phenomenon, the State I I ' form of the enzyme binds nucleotide and the resulting conformational change affords protection of the active site against the effects of the chelator. Occurrence of conformational changes is given further support by studies with the fluorescent hydrophobic probe TNS. State I enzyme, under the influence of Mg2+, exposes a more favorable hydrophobic en-

GLTJTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

169

vironment for the binding of the fluorophore. For glutamine to induce an additional conformational change, the protein molecule must first be further modified by PP-ribose-P. State Ι Γ enzyme also undergoes an instantaneous conformational change with the addition of Mg2+. However, in the presence of Mg2+, glutamine will induce a conformational change without prior conditioning of the enzyme by added PP-ribose-P. State ΙΓ enzyme may represent some type of stable enzyme-PP-ribose-P complex. The conformational changes induced by glutamine occur almost in­ stantaneously, whereas the acceleration of the enzymatic reaction occurs over a period of several minutes. Perhaps the instantaneous changes are followed by as yet undetectable conformational changes more directly related to the active site and largely unrelated to those changes which are creating a more favorable hydrophobic environment for the fluorophore. The purified enzyme loses all measurable activity in 72 hr, at — 20°C, and during this process State ΙΓ enzyme is converted into State I en­ zyme. Preparations of PP-ribose-P amidotransferase eluted from DEAE or DEAE-Sephadex columns, which have s values of about 9 S in sucrose density gradients, and appear to represent homogeneous fractions of 200,000 MW, behave variously as State I, State I F , or mixed forms of the enzyme. It was pointed out above that the 200,000 MW enzyme dis­ sociates into 100,000 MW dimers on dilution, and further dissociates into 50,000 MW monomer subunits in the presence of 60 milf mercaptoethanol. The monomer behaves as a State I form of the enzyme in the assay system. However, we have not been able to demonstrate any reaggregation of subunits in the presence of various combinations of sub­ strates. Thus, although changes in molecular weight during transitions involving State I, State II or I F , and State III forms have not been rigorously excluded, such changes appear improbable on the basis of available information. V. Enzyme Induction, Derepression, and Repression A. Bacterial Systems Nierlich and Magasanik (61) have demonstrated repression and derepression of six enzymes of purine biosynthesis in A. aerogenes, includ­ ing three of the pathway leading to IMP. Changes of activity of P P ribose-P amidotransferase are coordinate with those of FGAR-amidotransferase but noncoordinate with changes of activity of the other enzymes, including that of GAR synthetase, the second enzyme of the pathway.

170

JAMES B. WYNGAARDEN

Pur D mutants of Salmonella typhimurium (80) which are completely or almost completely deficient in activity of the second enzyme of purine biosynthesis, GAR synthetase, contain wild-type levels of PP-ribose-P amidotransferase when grown on xanthine (5 mg/ml). By contrast, Pur F mutants, which lack PP-ribose-P amidotransferase, produce higher, derepressed levels of GAR synthetase (88-180 nmoles total GAR per milligram of protein) compared with wild-type levels (35 units/mg) under similar growth conditions. Reem (66) has partially purified an enzymatic fraction from pigeon liver which appears to catalyze the formation of phosphoribosylamine from ribose 5-phosphate, ammonia, and ATP, and has summarized the literature suggesting the presence of this potential alternative first step of purine biosynthesis in crude extracts of bacterial and mammalian tissues. The pigeon liver fraction does not exhibit activity with P P ribose-P, Mg2+, and glutamine. However, ribose 5-phosphate aminotransferase activity has not been separated from GAR-synthetase ac­ tivity, which is much the more active of the two enzymatic functions. The Km values for ribose 5-phosphate and ammonia, and pH optimum, are high, rather similar to those of the nonenzymatic reaction between ribose 5-phosphate and ammonia (60, 63). The observation that Pur F mutants of S. typhimurium are stringent purine auxotrophs, suggests that the alternative reaction for synthesis of phosphoribosylamine does not function in vivo, at least in this organism. Doubt also exists whether this reaction functions in ascites cells (27). B. Mammalian Systems The work of McFall and Magasanik (54) contains very indirect in­ dications that enzymes of purine biosynthesis de novo, perhaps including PP-ribose-P amidotransferase, may be repressed in strain L of mouse fibroblasts cultured for several generations in adenine or guanosine. These agents produce a complete suppression of synthesis of purines de novo when offered to cells with such growth histories, and all purines of the soluble pools are then derived from the exogenous supplement. By contrast, addition of adenine or guanosine to cells not previously exposed to purines produces a condition in which one-half of the purines of the soluble pools still arise by synthesis de novo. Reem and Friend (68) find that mouse spleen normally contains no detectable activity of PP-ribose-P amidotransferase by the assay em­ ployed. Following infection with Friend leukemia virus, activity of PPribose-P amidotransferase appears by day 4, increases rapidly to a maxi­ mal peak of activity by days, 6-9, and thereafter declines gradually over 2-4 weeks. The enzyme activity which appears is subject to ribo-

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTRANSFERASE

171

nucleotide feedback inhibition both in vivo and in vitro. The appearance of enzyme correlates with the extent of infiltration of spleen with tumor cells. Whether PP-ribose-P amidotransferase is produced by derepression of host genome or by induction under the influence of viral genome is not known.

VI. Summary Kinetic studies do not unequivocally provide a determination of the minimal number of binding sites, since rate kinetic effects alone can give ligand concentrations of powers higher than the number of interact­ ing sites (73). In most actual cases, however, Hill coefficients have given an indication of the minimum number of sites (2> 3> 13, 75). In a few examples, the coefficients have equaled the theoretical number of sites, indicating a very strong degree of cooperativity (4, 47', 48). In many instances the Hill coefficient is less than the theoretical value. Hemo­ globin has a Hill coefficient of 2.8 for a total of 4 oxygen binding sites (82, 83). Fructose-1,6-diphosphatase from rabbit liver has a Hill coeffi­ cient of 1.7 for the binding of 4 equivalents of F D P per mole of tetrameric enzyme at saturation (65). In considering a schematic model of PP-ribose-P amidotransferase the main observations to be accommodated are (a) tetrameric structure (pigeon liver enzyme) ; (b) kinetic data for binding of at least 3 and possibly 4 molecules of PP-ribose-P ; of at least 2 molecules of glutamine ; and of 2 or 3 molecules of a single type of nucleotide and of as many as 5 molecules of mixtures of 6-amino and 6-hydroxypurine ribonucleotides (pigeon liver enzyme) ; (c) catalytic activity and nucleotide sensitivity of monomer and dimer forms under conditions in which deliberate at­ tempts to achieve and demonstrate reaggregation do not indicate a change in molecular weight. The inhibitor sites would have to be distinct from substrate sites, i.e., with no shared binding subgroups, in order to explain protection of the enzyme from o-phenanthroline inhibition by AMP or GMP, even in the enzyme which is insensitive to inhibition by nucleotide. Reasonable limit­ ing cases of models extend from 1 to 4 active sites, and from 2 to 8 nucleotide regulatory sites per tetramer. A few examples will be discussed. A first model envisions 4 identical subunits forming a tetramer with four catalytic sites and eight regulatory sites. Each monomer has one site for PP-ribose-P, glutamine, 6-amino- and 6-hydroxypurine ribo­ nucleotides, respectively. Theoretical values of n or n' of 4 for each ligand, or of 8 for nonhomologous ribonucleotide inhibitors acting to­ gether, are not observed because the strengths of the cooperative inter-

172

JAMES B. WYNGAARDEN

actions between sites are not sufficiently great, at least under the con­ ditions of study. A second model envisions two catalytic sites per tetrameric enzyme of 200,000 MW (22). Kinetic data do not indicate binding of more than two glutamine molecules by any PP-ribose-P amidotransferase studied thus far. Furthermore, affinity labeling studies show maximal binding of only 1 mole of 14C-labeled DON per 198,000 gm of enzyme (22). In this model two PP-ribose-P sites function as catalytic sites and two as modifiers. In addition, the enzyme has 8 nucleotide regulatory sites, as proposed above. Either catalytic site model may be altered to allow for 4 overlapping rather than 8 discrete nucleotide sites. Such a model would allow for cooperative effects between homologous inhibitor sites, and also for synergistic effects of binding of nonhomologous inhibitors, if it is assumed that binding energies for the two types of inhibitors vary with changes of conformational state (10), and that the tetramer may exist in several different hybrid forms such that one subunit binds a 6-aminopurine preferentially while another binds a 6-hydroxypurine. This model predicts that the maximal value of nf is 4; the one observed value of 4.8 would then presumably be 4 rather than 5. These several schematic possibilities are shown in Fig. 9. On the basis of these models, the experimental observations with respect to glutamine sites could be explained in two ways, which are not mutually exclusive, (a) The potential glutamine sites are distorted in the State I form of the enzyme; ligand-induced conformational changes brought about by PP-ribose-P and Mg2+ are required to generate four glutamine-binding sites in the tetramer ; cooperatively is only moderately strong as indicated by n values of 2. (b) The tetramer possesses four glutamine-binding sites but only two are available for the catalytic process and only one can be filled by DON. CTP-synthetase of E. coli is a tetrameric enzyme (47) with reactive sites for glutamine, UTP, and ATP. The glutamine sites show negative cooperativity (45 > 48), but values of n > 2 are required to explain the biphasic substrate-velocity curve of glutamine (45\ 78). The maximal n values of 3.4 for UTP, and of 3.8 for ATP (48) strongly suggest that four binding sites exist for each nucleoside triphosphate, and by inference for glutamine as well (47). Nevertheless, only 2 of the 4 probable gluta­ mine sites can be filled by 14C-labeled DON in affinity labeling experi­ ments (47)· The additional event involved in the covalent binding of DON to the enzyme may induce a drastic conformational change, and result in such strong negative cooperativity between the glutamine sites that (irreversible) binding of a third or fourth DON molecule does not occur under conditions of study.

GLUTAMINE PHOSPHORIBOSYLPYROPHOSPHATE AMIDOTBANSFERASE

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173

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Activated Fully Active 1 State Π ) I State m s) PRPP+MgV^ ^Glutamine,,

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FIG. 9. Schematic models of possible structures of PP-ribose-P amidotransferase. At the top are models of two types of monomers. In the one on the left two discrete nucleotide binding sites are shown; in the one on the right overlapping sites are shown for 6-amino and 6-hydroxypurine ribonucleotides. Open and solid rectangles indicate substrate binding sites (P = PP-ribose-P, Gin = glutamine). I n the center are shown various hypothetical forms of the tetramer, illustrating the changes of conformational state brought about by interaction with substrates, and also indicating that additional conformational variations may determine whether the enzyme is nucleotide sensitive (s) or insensitive (i). I n the lower portion of the figure, variations of tetramer arrangements are shown, which place different limits on maximal binding values of substrates and inhibitors. Each of these models can be substituted for the one at the left on the line above, and carried through the same sequence of hypothetical conformational changes.

Similarly, the binding of one molecule of DON by PP-ribose-P amidotransferase may induce such strong negative cooperativity that a second molecule is not bound. Alternatively, the additional méthylène group of DON may provide added steric hindrance if the glutamine sites happen to be located near each other in adjacent subunits (47). It seems un­ likely that one DON molecule could have this effect on three neighbor­ ing glutamine sites, as would be required in the first model, but the second model would require such an effect on only one additional site. In any case, DON is not a valid probe of the number of glutaminebinding sites. Substrate and inhibitor binding data will be required to discriminate among these models of enzyme structure; the first model (four catalytic and eight nucleotide regulatory sites) appears to be the most plausible at the present time. The regulatory properties of glutamine PP-ribose-P amidotransferase have been viewed from the standpoint of the symmetry model of Monod, Wyman, and Changeux (58) by Nagy (59) in the case of the enzyme from S. pombe; and from the standpoint of the ligand-induced or sequential model of Koshland, Nemethy, and Filmer (J$) by Rowe,

174

JAMES B. WYNGAARDEN

Coleman, and Wyngaarden (71) in the case of the enzyme from pigeon liver. The slowly progressive positive homotropic effects of substrates in the activation of pigeon liver enzyme appear strongly in favor of the sequential model. Similarly, the capricious behavior of the enzyme to­ ward nucleotide inhibitors, and the variability of the indices of cooperativity of substrate or inhibitor sites favor this model. The limited observations made thus far which indicate that the monomer form of the enzyme behaves as a State I enzyme, and undergoes progressive sequential activation by the PP-ribose-P-Mg 2+ complex and by glutamine, also favor the concept of hybrid conformational states as ligand is bound, occurring within the polypeptide strand and not wholly de­ pendent upon the tetrameric structure. Koshland's concepts of enzyme flexibility, of enzyme-substrate interaction in establishment of the catalytically active site, and of enzyme-inhibitor interaction in modifying the activity of the catalytic site (39-41) would appear to form the best point of departure for considering the regulatory behavior of PP-ribose-P amidotransferase. REFERENCES

1. 2. 3. 4.

Abrams, R., Arch. Biochem. Biophys. 33, 436 (1951). Atkinson, D. E., Science 150, 851 (1965). Atkinson, D. E., Annu. Rev. Biochem. 35, 85 (1966). Atkinson, D. E., Hathaway, J. A., and Smith, E. C, / . Biol. Chem. 240, 2682 (1965). 6. Bachmayer, H., Piette, L. H., Yasunobu, K. T., and Whiteley, H. R., Proc. Nat. Acad. Sci. U. S. 57, 122 (1967). 6. Balis, M. E., Levin, D. H., Brown, G. B., Elion, G. B., Vanderwerff, H., and Hitchings, G. H., / . Biol. Chem. 196, 729 (1952). 7. Bolton, E. T., Abelson, P. H., and Aldous, E., / . Biol. Chem. 196, 179 (1952). 8. Brockman, R. W., Cancer Res. 23, 1191 (1963). 9. Buchanan, J. M., Hartman, S. C , Herrmann, R. L., and Day, R. A., J. Cell. Comp. Physiol. 54, Suppl. 1, 139 (1959). 10. Caskey, C. T., Ashton, D. M., and Wyngaarden, J. B., J. Biol. Chem. 239, 2570 (1964). 11. Flaks, J. G., Erwin, M. J., and Buchanan, J. M., J. Biol. Chem. 228, 201 (1957). 12. Frère, J.-M., Schroeder, D. D., and Buchanan, J. M., J. Biol. Chem. 246, 4727 (1971). 13. Gerhart, J. C, and Pardee, A. B., Fed. Proc, Fed. Amer. Soc. Exp. Biol. 23, 727 (1964). U. Goldthwait, D. A., J. Biol. Chem. 232, 1051 (1956). 15. Goldthwait, D. A., and Bendich, A., J. Biol. Chem. 196, 841 (1952). 16. Goldthwait, D. A., Greenberg, G. R., and Peabody, R. A., Biochim. Biophys. Ada 18, 148 (1955). 17. Goldthwait, D. A., Peabody, R. A., and Greenberg, G. R., J. Amer. Chem. Soc. 76, 5258 (1954).

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18. Goldthwait, D. A., Peabody, R. A., and Greenberg, G. R., J. Biol. Chem. 221, 569 (1955). 19. Gots, J. S,. J. Biol. Chem. 228, 57 (1957). 20. Gots, J. S., and Goldstein, J., Science 130, 622 (1959). 21. Hartman, S. C , / . Biol. Chem. 238, 3024 (1963). 22. Hartman, S. C , / . Biol. Chem. 238, 3036 (1963). 23. Hartman, S. C , and Buchanan, J. M., J. Biol. Chem. 233, 451 (1957). 24- Hartman, S. C , Levenberg, B., and Buchanan, J. M., J. Biol. Chem. 221, 1057 (1956). 25. Henderson, J. F., J. Biol. Chem. 237, 2631 (1962). 26. Henderson, J. F., Biochem. Pharmacol. 12, 551 (1963). 27. Henderson, J. F., Biochim. Biophys. Ada 76, 173 (1963). 28. Henderson, J. F., Brox, L. W., Kelley, W. N., Rosenbloom, F. M., and Seegmiller, J. E., J. Biol. Chem. 243, 2514 (1968). 29. Henderson, J. F., Caldwell, I. C , and Paterson, A. R. P., Cancer Res. 27, 1773 (1967). 30. Henderson, J. F., and Khoo, M. K. Y., / . Biol. Chem. 240, 3104 (1965). 31. Henderson, J. F., Rosenbloom, F. M., Kelley, W. N., and Seegmiller, J. E., J. Clin. Invest. 47, 1511 (1968). 32. Hill, A. V., J. Physiol. (London) 40, iv (1910). 33. Hill, A. V., Biochem. J. 7, 471 (1913). 34. Hill, D. L., and Bennett, L. L., Jr., Biochemistry 8,122 (1969). 35. Hori, M., and Henderson, J. F., J. Biol. Chem. 241, 3404 (1966). 36. Johnson, F. H., Eyring, H., and Williams, R. W., / . Cell. Comp. Physiol. 20, 247 (1942). 37. Koch, A. L., Putnam, F. W., and Evans, N. A., Jr., / . Biol. Chem. 197, 105 (1952). 38. Kornberg, A., Lieberman, I., and Simms, E. S., / . Biol. Chem. 215, 389 (1954). 39. Koshland, D. E., Jr., Cold Spring Harbor 28, 473 (1963). 40. Koshland, D. E., Jr., Curr. Top. Cell. Regul. 1, 1 (1969). 41. Koshland, D. E., Jr., in "The Enzymes" (P. Boyer, ed.), 3rd ed., Vol. 1, p. 341. Academic Press, New York, 1970. 42. Koshland, D. E., Jr., Némethy, G., and Filmer, D., Biochemistry 5, 365 (1966). 43. Krenitsky, T. A., and Papaioannou, R., J. Biol. Chem. 244, 1271 (1969). 44. Le Page, G. A., and Jones, M., Cancer Res. 21, 642 (1961). 45. Levitzki, A., and Koshland, D. E., Jr., Proc. Nat. Acad. Sci. U. S. 62, 1121 (1969). 46. Li, H. C., and Buchanan, J. M., J. Biol Chem. 246, 4713 (1971). 47. Long, C. W., Levitzki, A., and Koshland, D. E., Jr., / . Biol. Chem. 245, 80 (1970). 48. Long, C. W., and Pardee, A. B., J. Biol. Chem. 242, 4715 (1967). 49. Love, S. H., and Gots, J. S., J. Biol. Chem. 212, 647 (1955). 50. Martin, R. G., / . Biol. Chem. 238, 257 (1963). 51. McClure, W. O., and Edelman, G. M., Biochemistry 5,1908 (1966). 52. McClure, W. O., and Edelman, G. M., Biochemistry 6, 559 (1967). 53. McCollister, R. J., Gilbert, W. R., Jr., Ashton, D. M., and Wyngaarden, J. B., J. Biol. Chem. 239, 1560 (1964). 54. McFall, E., and Magasanik, B., / . Biol. Chem. 235, 2103 (1960). 55. Mizobuchi, K., and Buchanan, J. M., J. Biol. Chem. 243, 4853 (1968).

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56. Momose, H., Nishikawa, H., and Katsuya, N., J. Gen. Appi. Microbiol. 2, 211 (1965). 57. Monod, J., Changeux, J.-P., and Jacob, F., / . Mol. Biol. 6, 306 (1963). 58. Monod, J., Wyman, J., and Changeux, J.-P., / . Mol. Biol. 12, 88 (1965). 59. Nagy, M., Biochim. Biophys. Ada 198, 471 (1970). 60. Nierlich, D. P., and Magasanik, B., / . Biol. Chem. 236, PC32 (1961). 61. Nierlich, D . P., and Magasanik, B., Fed. Proc, Fed. Amer. Soc. Exrp. Biol. 22, 476 (1963). 62. Nierlich, D. P., and Magasanik, B., J. Biol. Chem. 240, 358 (1965). 63. Nierlich, D . P., and Magasanik, B., J. Biol. Chem. 240, 366 (1965). 64- Phillips, W. D., Knight, E., Jr., and Blomstrom, D. C , in "Non-Heme Iron Proteins: Role in Energy Conversion" (A. San Pietro, ed.), p. 69. Antioch Press, Yellow Springs, Ohio, 1965. 65. Pontremoli, S., and Horecker, B. L., Curr. Top. Cell. Regul 2, 173 (1970). 66. Reem, G. H., / . Biol. Chem. 243, 5695 (1968). 67. Reem, G. H., J. Clin. Invest. 47, 83a (1968). 68. Reem, G. H., and Friend, C , Science 157, 1203 (1967). 69. Remy, C. N., Remy, W. T., and Buchanan, J. M., J. Biol. Chem. 217, 885 (1955). 70. Rottman, F., and Guarino, A. J., Biochim. Biophys. Acta 89, 465 (1964). 71. Rowe, P . B., Coleman, M. D., and Wyngaarden, J. B., Biochemistry 9, 1948 (1970). 72. Rowe, P . B., and Wyngaarden, J. B., J. Biol. Chem. 243, 6373 (1968). 73. Sanwal, B. D., and Cook, R. A., Biochemistry 5, 886 (1966). 74- Sartorelli, A. C , and LePage, G. A., Cancer Res. 18, 1329 (1958). 74a. Shiio, I., and Ishii, K., J. Biochem. 66, 175 (1969). 75. Stadtman, E. R., Advan. Enzymol. 28, 41 (1966). 76. Stadtman, E . R., in "The Enzymes" (P. Boyer, ed.), 3rd ed., Vol. 1, p. 397. Academic Press, New York, 1970. 77. Taketa, K., and Pogell, B. M., J. Biol. Chem. 240, 651 (1965). 78. Teipel, J., and Koshland, D. E., Jr., Biochemistry 8, 4656 (1969). 79. Thomson, R. Y., Paul, J., and Davidson, J. N., Biochem. J. 69, 553 (1958). 80. Westby, C. A., and Gots, J. S., J. Biol. Chem. 244, 2095 (1969). 81. Whitaker, J. R., Anal. Chem. 35, 1950 (1963). 82. Williams, R. J. P., J. Chem. Soc. London p. 137 (1955). 82a. Wood, A. W., and Seegmiller, J. E., Fed. Proc, Fed. Amer. Soc. Exp. Biol. 30, 1113 (1971) (abstr.). 83. Wyman, J., Advan. Protein Chem. 4, 407 (1948). 84- Wyngaarden, J. B., unpublished data, or recalculations from data of Caskey et al. (10), Rowe et al. (71), or Wyngaarden and Ashton (86). 85. Wyngaarden, J. B., Appel, S. H., and Rowe, P . B., in "Exploitable Molecular Mechanisms and Neoplasia," p. 415. Williams & Wilkins, Baltimore, Maryland, 1969. 86. Wyngaarden, J. B., and Ashton, D. M., J. Biol. Chem. 234, 1492 (1959). 87. Wyngaarden, J. B., Silberman, H . R., and Sadler, J. H., Ann. N. Y. Acad. Sci. 75, 45 (1958).

The Regulatory Influence of Allosterîc Effectors on Deoxycytidylate Deaminases I

FRANK MALEY

I

GLADYS F. MALEY

I Division of Laboratories and I Research I New York State Department of I Health I Albany, New York I. Introduction A. Distribution and Role of Deoxycytidylate Deaminase . . B. Methods of Detection and Assay II. Feedback Regulation of Deoxycytidylate Deaminase . . . A. Discovery B. dCTP, an Activating and Stabilizing Ligand; dTTP, a Reversible Inhibitor C. Metal Ion Requirement III. Allosteric Transitions Effected by the Regulatory Ligands . . A. Homo tropic and He tero tropic Interactions B. Evidence Implicating Conformational Transitions and Quaternary Structural Changes IV. Active and Inactive States Associated with Deoxycytidylate Deaminase A. Quaternary Structural Changes Associated with the Altered States B. Reactivation of Inactive Enzyme by the Allosteric Ligands and Thiols C. Effect of the Sulfhydryl-Disulfide Interchange (SDI) Enzyme on Subunit Structure and Function D. Utilization of the Activation-Inactivation Phenomenon to Obtain Homogeneous Enzyme V. Use of Inhibitors to Define the Physiological Significance of Deoxycytidylate Deaminase A. Incorporation of Labeled Deoxycytidine in the Presence of Deoxyuridine and Thymidine B. Tetrahydrodeoxyuridylate VI. Distribution and Role of Deoxycytidylate Deaminase in Bacterial Systems A. Induction in Bacteriophage-Infected Cells; the Titer Effect . B. Substrate Specificity of the Bacteriophage-Induced Enzyme . C. Mutants Deficient in Deoxycytidylate Deaminase . . . VII. Regulatory Responses of the Bacteriophage-Induced Deoxycytidylate Deaminase to Its Metabolic End Products . 177

178 178 179 180 180 182 185 185 186 188 191 191 192 193 194 195 195 197 199 200 201 205 205

178

FRANK MALEY AND GLADYS F. MALEY

VIII.

IX. X. XI.

A. Role of the Allosteric Ligands: dCTP, HM-dCTP, and dTTP B. Allosteric Transitions Associated with the Regulatory Process Physical and Chemical Properties of T2r+ BacteriophageInduced Deoxycytidylate Deaminase A. Purification to Homogeneity B. Molecular Weight of the Oligomer and Protomer . . C. Diffusion Coefficient and Stokes Radius D. Dependence of Oligomeric Structure on Protein Concentration E. Amino Acid Analysis F. Comparative Studies with T4 Bacteriophage-Induced Deoxycytidylate Deaminase The Effect of pH on the Allosteric Response of Deoxycytidylate Deaminase Studies on the in Vitro Synthesis of Bacteriophage-Induced Deoxycytidylate Deaminase Conclusion References

205 208

.

209 209 211 213 216 217 217 219 222 224 226

I. Introduction

A. Distribution and Role of Deoxycytidylate Deaminase The discovery of deoxycytidylate deaminase resulted from independent investigations on the metabolism of deoxycytidine derivatives in sea urchin eggs (75) and in rat embryo extracts (48). At the time, the deaminase appeared to provide the only direct means of forming dUMP, a substrate for the critical enzyme thymidylate synthetase, and thus it was viewed as playing a possible role in the regulation of DNA syn­ thesis or contributing perhaps to the efficiency of pyrimidine nucleotide utilization. Potential evidence in support of these assumptions was pro­ vided by numerous studies indicating that deaminase activity was usu­ ally elevated in mitotically active normal and neoplastic tissues. In ad­ dition to those sources already mentioned, the enzyme was found to be active in regenerating liver (40, 48, 62, 88), various solid and ascitic neoplasms (14, 16, 41, 67, 72, 86), virus-transformed cells (28, 81), and in the serum of patients with various acute pathologies (57). The sig­ nificance of these findings was somewhat clouded by the presence of surprisingly high levels of the enzyme in such mitotically inert tissues as adult rabbit, monkey, and human liver (48), a result that complicated the presentation of a rational explanation for the physiological role of the deaminase.

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

179

An additional complication was the demonstration by Reichard et al (69, 70) of a new reaction sequence U D P -► dUDP -► dUTP -> dUMP

in which dUMP was synthesized by the direct conversion of a ribose to a deoxyribose. As will be shown in Section V, studies with a specific in­ hibitor of deoxycytidylate deaminase suggest that the above metabolic sequence involving ribonucleotide reductase is probably the major source of dUMP in animal tissues. The role of the deaminase in most bacterial systems is marginal at best or nonexistent, except in those cases where the enzyme is induced on bacteriophage infection, and even here the presence of the enzyme is not essential for survival {26). For these reasons, interest in deoxy­ cytidylate deaminase as such has waned, but other factors have pre­ vented it from dissipating completely. Thus in the course of these studies, we found the enzyme to be markedly susceptible to feedback inhibition by dCTP and dTTP (50), and it has proved since to be an exceptionally convenient model for the study of the mechanisms involved in eliciting aUosteric transitions associated with deoxycytidylate de­ aminase. In addition, the discovery of a similar enzyme in T2 bacteriophage-infected Escherìchia coli provided an opportunity for a compara­ tive study of biological catalysts of different phylogenetic origin from which an interesting pattern of subtle differences has emerged. The dis­ cussion that follows will attempt to define the properties of both en­ zymes and the fascinating mechanism that nature has devised to regu­ late their activities. B. Methods of Detection and Assay The rationale for most of the assay procedures depends on the mark­ edly different absorption spectra of the substrate and product. In our initial studies (48), it was found most convenient to measure the decrease in absorption at 290 nm resulting from the conversion of dCMP to dUMP. Since the difference in the millimolar extinction co­ efficients of the two nucleotides at 290 nm is 10, a fairly sensitive assay was provided, one more convenient than the measurement of the change in absorbance at various wavelengths (30, 79). Where even more sensi­ tive methods of detection were required, the deamination of dCMP-2- 14 C, dCMP- 3 H,ordCMP- 32 Pwas followed by passage of the denatured centrifuged protein solution through a Dowex 50 H + column (40). If necessary, the products can be separated further by chromatography on DE-81 ion exchange paper (31). However, these methods are often subject to er­ roneous interpretation as the column elution does not distinguish between

180

FRANK MALEY AND GLADYS F. MALE Y

dUMP and deoxyuridine and paper chromatography does not separate dUMP from dCDP readily. The latter nucelotide was produced by a heat-stable kinase (46) that converts dCMP to dCDP, a compound not retained by Dowex 50 H + . To circumvent this problem, the reaction mix­ ture was treated with snake venom, a procedure that converts nucleotides to nucleosides and provides deoxyuridine as a measure of deoxycytidylate deaminase activity (35). These words of caution are expressed as results have been reported too often without taking the precaution of making certain that only one enzyme and one product are involved. Ammonia release has been used also as a measure of activity either following diffusion (72), or directly on the reaction mixture, by the sen­ sitive Berthelot reaction (57). Caution must be exercised here, too, be­ cause of the combined action of phosphatases and nucleoside deaminases, in addition to the nonspecific release of ammonia. The method found to be most convenient and accurate, particularly with partially purified enzyme, evolved with the advent of the Gilford recording spectrophotometer. Because this instrument, or one based on the same principle, provides a continuous kinetic assay, it has been used exclusively in our laboratory. Under the condition used to assay the de­ aminase, the millimolar extinction coefficient of dCMP at pH 8.0 and 290 nm is 2.33 and that of dUMP is 0.358. In order to convert absorbance change to micromoles of dUMP formed, the following relationship was developed (39) : y = 2.33(a - x) + 0.358z where y = absorbance, a = initial concentration of dCMP in /xmoles, and x = /xmoles dUMP. Thus when the initial concentration of dCMP is 1 mM, an absorbance change of 1.97 is equivalent to the deamination of 1 /rniole of dUMP. A unit is taken as the amount of dCMP deaminated per minute at 37°C. Although assayed at 30°C in the Gilford, the results are multiplied by 2 (obtained experimentally) to yield values comparable to those reported in the literature. In earlier studies (52), a unit of 1 /miole per 10 minutes was used. II. Feedback Regulation of Deoxycytidylate Deaminase A. Discovery One of the problems encountered in our initial studies with the de­ aminase in embryonic tissues was its apparent instability to storage. Attempts at purification were hampered similarly until, in searching for methods to stabilize the enzyme, it was found that the addition of low levels of dCTP (10_5.M) to partially or even totally inactive deaminase

181

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

TABLE I RESTORATION OF DEOXYCYTIDYLATE DEAMINASE ACTIVITY BY d C T P °

dCMP deaminated Treatment of extract

(μπιο1β/10 min)

0.286 0.002 0.000 0.007

0.360 0.333 0.280 0.307

0.198 0.071

0.401 0.369

Chick embryo None Frozen 5 days Frozen 25 days Heated at 37°C for 10 minutes Rat embryo None Dowex 1-formate a b

+dCTP>

-dCTP (μπιοΐβ/ΐθ min)

Reaction conditions and assay are described by Maley and Maley {50). d C T P was.present a t 50 nmoles in the reaction mixtures.

preparations completely restored the activity {50). These initial studies with crude embryo extracts are presented in Table I. An even more striking demonstration of this phenomenon is presented in Fig. 1, where it is shown that the extent of activation is dependent on the concentra­ tion of substrate and the age of the extract (aging conditions are given ▼

50

40

/ / /

K

O

/ /

* 30 -

»

LU —1

o \20 z

J>

5

-l^io 0

| r

f

/

y

^ ,—'

/

_Γ->—— ^ " ~ -

1

1

1

1

1

1

1

1

j(LITER/MOLE)XIO-3 FIG. 1. Effect of d C T P on deoxycytidylate deaminase in fresh and aged chick embryo (4-day) extracts. For experimental details and assay procedures, see Maley and Maley {50). Aged extract: T , - d C T P ; # , + d C T P . Fresh extract: V , - d C T P ; O , + dCTP.

182

FRANK MALEY AND GLADYS F. MALEY 1.0 (Λ Ixl _l

o

S

o_ 0.5 o ■σ

o

0.9

t\

GO

\

or o ω




< |

50

OC

o S 25 Ld O CE L±J Û-

0

MINUTES

FIG. 15. Rate of inactivation of deoxycy tidy late deaminase by the SDI enzyme. Separate reaction vessels, at 25°C, contained 3 m l MgCl2; 50 m l potassium phosphate (pH 7.5) ; 1.0 unit of chick embryo deaminase; and the indicated volume of DEAE-purified SDI'enzyme (11) in a final volume of 0.15 ml. At the indicated times, 20-μ1 aliquote w,ere removed and assayed for deaminase activity by Assay 1 (54)· Reprinted with permission from Maley and Maley (44) > copyright 1970, Pergamon Press Ltd., Oxford.

194

FRANK MALEY AND GLADYS F. MALEY

FIG. 16. Protective effect of dCTP and dTTP against the SDI enzyme. The incubation mixtures contained 2.5 /imoles of potassium phosphate (pH 7.5) ; 1 μτηοίβ of MgCl2; 20 nmoles of dCTP or dTTP as shown; 10 μΐ of chick embryo deaminase (1.0 unit, specific activity, 445); 20 μ\ of SDI enzyme; and water to a final volume of 0.1 ml at 25°C. Aliquots of 10 μΐ were removed and assayed for deaminase activity (54) at the indicated intervals. Reprinted with permission from Maley and Maley (44), copyright 1970, Pergamon Press Ltd., Oxford.

SDI preparation, but this appears unlikely in view of (1) the dissociation of the enzyme by the SDI enzyme preparation; (2) the restoration of enzyme activity and structure by 2-mercaptoethanol ; (3) the protection of the deaminase by dCTP (Fig. 16), which is strikingly similar to the results obtained in Fig. 4. Nonetheless, further studies will be necessary to validate the above assumption. In any event, the inhibitory effect of the microsomal SDI enzyme on the deaminase provides a plausible ex­ planation for the observations of Fiala and Fiala (16) concerning an apparent microsomal agent that was detrimental to deoxycy tidy late deaminase activity in tissue extracts. In view of the sulfhydryl sensitivity of the deaminase, it was not surprising to find in plotting pH versus pKm that the apparent pK's of the enzyme substrate complex were 6.8 and 9.5 (44)· Although the latter pK could be representative of a tyrosyl group, it is also consistent with that for a sulfhydryl group in a negatively charged environment. The titration of enzyme activity with iV-ethylmaleimide as a function of pH is consistent with the latter interpretation (unpublished observations). D. Utilization of the Activation-lnactivation Phenomenon to Obtain Homogeneous Enzyme The remarkable ability of the deaminase to undergo reversible changes in molecular weight suggested a means of obtaining pure chick embryo

195

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

TABLE V PURIFICATION OF CHICK EMBRYO DEOXYCYTIDYLATE DEAMINASE*1

Fraction 1. 2. 3. 4. 5. 6. 7. α 6

Extract Ammonium sulfate Phosphocellulose I Sèphadex G-2006 Phosphocellulose II Sephadex G-100 Sephadex G-200

Total protein (mg)

Specific activity (units/mg)

2600 1250 173 305 28 4.2 0.32

0.05 0.115 2.30 2.50 18.0 89.5 630.0

Seven-day-old chick embryos were used. Several preparations from fraction 3 were usually combined for this step.

deoxycytidylate deaminase. The major steps in the purification scheme presented in Table V are 6 and 7, where the enzyme was first inactivated by prolonged dialysis, a procedure that provides the low molecular weight disaggregated form of the deaminase. The inactive form of the deaminase was isolated by chromatography on Sephadex G-100, followed by reactivation with the combination of Mg-dTTP and 2-mercaptoethanol. The reaggregated active enzyme was chromatographed on Sepha­ dex G-200. The overall purification was 12,000-fold with only a single band of protein evident on polyacrylamide gel electrophoresis. The enzyme was associated with the protein band. V. Use of Inhibitors to Define the Physiological Significance of Deoxycytidylate Deaminase A. Incorporation of Labeled Deoxycytidine in the Presence of Deoxyuridine and Thymidine Two major pathways for the de novo synthesis of dUMP have been delineated in animal tissues, one utilizing reactions 1, 2, and 3 of Fig. 17 and the other reaction sequence 1, 2, UTP-»CTP, 8 and 5. It is a question of both academic and practical importance to determine the extent to which each pathway contributes to the formation of dUMP. With the appropriate inhibitor, a solution to this problem should be possible. Since a selective inhibitor of the CDP-»dCDP reaction prob­ ably will not be found, the most practical inhibitor would be that of the deaminase. The only known effective inhibitors of the deaminase to date suffer from the fact that they are all phosphorylated compounds, such as dTTP, dGMP, and 4-iV-hydroxy-dCMP, and will not pass the cell mem­ brane without first being dephosphorylated. However, some inferences

196

FRANK MALEY AND GLADYS F. MALEY

dTTP dUTP

dCTP dTDP CDP-

4

-»-dCDP

dUDP-

-UDP

2

CMP

-dUMP

f7 Cr

j UMP-

-(OA)

-Udr

FIG. 17. Pathway for pyrimidine metabolism in chick embryo ; orotic acid (OA).

can be made as indicated by the apparent sparing effect imposed by thymidine on deoxycytidine utilization in chick embryo (1$, 51). As shown in these studies, the normal ratio of label for dCMP to dTMP in DNA when chick embryo mince is incubated with deoxycytidine-214 C is 1:2, indicating that deamination reactions are favored. Thus dTMP may arise by a combination of two pathways: d C M P -> d U M P -* d T M P deoxycytidine /"

\

deoxyuridine -> d U M P -> d T M P

(1) (2)

By raising the external thymidine level, the dCMP to dTMP count ratio approached 2:1. Since DNA synthesis was not impaired, the en­ hanced count in the dCMP arose most probably from an elevation in the deoxycytidine nucleotide pool, produced as a consequence of the inhibition of deoxyuridine phosphorylation or of deoxycytidylate deami­ nation. As indicated earlier (4#), dTTP inhibited both reactions. How­ ever, it was not possible to determine from these experiments which of the two pathways, (1) or (2), contributes more to the formation of dTMP and thus when inhibited produces an elevated deoxycytidine nucleotide pool. Evidence favoring (1) as the major route to dTMP synthesis with deoxycytidine as a precursor is supported by the results in Fig. 18, where it is shown that cold deoxyuridine added as a trap for the deoxycytidine-2-14C metabolized by pathway (2) effected a 50% decrease in the specific activity of DNA dTMP, while that in the de­ oxyuridine pool was diluted by over 95%. Some count in the DNA was, no doubt, contributed by pathway (2) ; however, most would appear to come from (1). The cytotoxic effect of thymidine noted in cell culture

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

197

14

1" * 8

I 6

o O

1.0 2.0 DEOX Y U RI DI N E(/*moles)

FIG. 18. Effect of deoxyuridine on the incorporation of deoxycytidine-2-14C into chick embryo DNA. The incubation mixture consisted of 0.45 /imole deoxycytidine-2-14C (1.8 X 10e cpm//imole) ; deoxyuridine as indicated; 0.8 gm of 4-dayold chick embryo mince; and Krebs-Ringer phosphate to 2.0 ml. After 2 hours of shaking at 37°C, the reactions were stopped by the addition of 0.1 volume of cold 70% perchloric acid. The isolated DNA was digested with DNase and phosphodiesterase, and the 5'-nucleotides were separated by electrophoresis as described (49). Deoxyuridine was isolated from the KOH-neutralized extract following pas­ sage over a mixed ion exchange column by 2-dimensional chromatography (38).

experiments (59, 60), which is believed to be a consequence of the inhibi­ tion of the CDP—»dCDP reaction by dTTP, was not a factor in the chick embryo studies, since DNA synthesis was not impaired by thymidine. Since the apparent sparing effect described above was not observed with cytidine-2- 14 C, a plausible assumption would be that most of the de novo dUMP is formed from orotic acid, not by the reaction sequence CDP -> dCDP -> dCMP -> dUMP

(3)

A similar assumption was arrived at by Crone and Itzhaki (9) from incorporation studies with regenerating liver. B. Tetrahydrodeoxyuridylate A report by Hanze (27) that tetrahydrouridine was an effective in­ hibitor of liver cytidine deaminase suggested to us that a similar inhibi­ tion of deoxycytidylate deaminase might be effected by tetrahydrode­ oxyuridylate (H 4 -dUMP). The compound was prepared by the reduction of deoxycytidine 5'monophosphate with the aid of a rhodium on alumina catalyst (27) ; as shown in Fig. 19, it was found to be a potent competitive inhibitor of the deaminase, with a Ki of at least 1 to 2 X 10~8 M (45). The potential

198

FRANK MALEY AND GLADYS F. MALEY

effectiveness of the inhibitor against deoxycytidylate deaminase as de­ termined by the Km:K{ ratio was of the order of 3 to 6 X 104, which is of a greater magnitude than that for F-dUMP and thymidylate synthetase. Considering that the compound may not be completely pure and that an asymmetric hydroxyl is present on the 4-position of the ring, it may be even more potent. Unfortunately, this inhibitor possesses the same debilitating characteristic of the other phosphorylated compounds discussed above. However, sufficient amounts of the H 4 -deoxyuridine appear to be phosphorylated in situ to inhibit completely the deamination of dCMP (45). Accumulation of H 4 -deoxyuridine in situ also blocks the nucleoside deaminase. Still, with deoxycytidine-2-14C as a precursor, only a 62% inhibition of deoxyuridine and dUMP formation could be effected with the chick embryo mince system. An apparent increase in the thymidine level was noted in spite of the sharp reduction in deoxyuridine, suggesting the possibility of another pathway for the formation of dTMP. A compensatory pathway such as described recently by O'Donovan and Neuhard (66) for Escherichia coli and Salmonella typhimunum that involves a dCTP deaminase in the reaction sequence dCTP -» dUTP —» dUMP -» dTMP could explain these anomalous findings. However, a dCTP deaminase is yet to be reported in animal tissues. With deoxycytidine-2-14C as a precursor and H 4 -dUMP as an inhibitor, it was anticipated that the deoxycytidine nucleotide pool would be ele­ vated, which resulted in a sparing effect similar to that obtained with thymidine (J$, 50). This effect was exactly the result obtained as the dCMP/dTMP ratio in DNA increased from 0.63 to 1.6. When cytidine2-14C was used as a precursor together with H 4 -deoxyuridine and H 4 uridine as inhibitors (the latter to block cytidine deaminase), a 60% impairment in dTMP synthesis was obtained without an apparent sparing or inhibitory effect on dCMP synthesis (Table VI). The fact

H 4 -dUMP (μΜ)

[dCMP]"l(m/lO"1

FIG 19. Dixon (A) and Lineweaver-Burk (B) plots to determine the type of inhibition effected by tetrahydro-dUMP on deoxycytidylate deaminase. The en­ zyme was assayed as described by Maley and Maley (54).

199

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES T A B L E VI ENHANCEMENT OF FLUORODEOXYURIDINE INHIBITION OF C Y T I D I N E - 2 - 1 4 C INCORPORATION INTO D N A

BY H 4 - U R I D I N E AND

H^DEOXYURIDINE«

DNA DNA (cpnii/mg X IO" 3 )

Additions 0 H 4 -Uridine

+

+

37.0

28.9

35.1

7.17

17.3

20.9

4.86

14.4

16.6

2.48

18.6

\

/

H 4 -Deoxyuridine >

+

36.0 )

H 4 -Deoxyuridine ) F-Deoxyuridine H 4 -Uridine

dCMP dTMP (cpm//umole X IO - 3 )

\

F-Deoxyuridine / α

D a t a from Maley and Maley (45).

that DNA dTMP was still labeled, although reduced by 40%, indicates that the block imposed by H 4 -dUMP in situ was inadequate, or that a compensatory thymidine nucleotide pathway may be involved, as dis­ cussed above. It is doubtful that sufficient cytidine-2- 14 C was deaminated and utilized by pathways 2 and 3 (Fig. 17) to account for the incorpora­ tion into dTMP, since H 4 -uridine alone was found to inhibit the deamination of cytidine-2- 14 C by almost 90% (45). While reactions 8, 5, and 6 of Fig. 17 possess the capacity to supplement dTMP synthesis, the finding that DNA synthesis was unaffected in spite of a 40% reduction in count from cytidine-2- 14 C (Table VI) suggests that most of the DNA dTMP results from de novo synthesis via reactions 1, 2, and 3. Since these studies suggest that H 4 -deoxyuridine may be useful in limit­ ing dTMP production in certain instances, the effect of this nucleoside in potentiating the inhibition of dTMP synthesis was tested. From the results presented in Table VI, it would appear that the inhibition pro­ moted by fluorodeoxyuridine can be enhanced by H 4 -deoxyuridine. VI. Distribution and Role of Deoxycytidylate Deaminase in Bacterial Systems Although deoxycytidylate deaminase is widely distributed in the ani­ mal kingdom, it has been found outside of this realm in only a limited number of cases. Perhaps the enzyme is present at much lower levels or has not been investigated thoroughly enough, but in any event it has been located in only limited quantities in Lactobacillus acidophilus {84), Chlorella pyeonoidosa (83), and Staphylococcus aureus (4). Fortunately,

200

FRANK MALEY AND GLADYS F . MALEY

however, after the reports of the existence of the deaminase in animal tissues, Keck et al. (30) searched for the enzyme in T2 bacteriophageinfected Escherìchia coli and found a striking increase in activity. In view of present information, their finding was the result of a rather fortuitous selection of substrate concentration, for if the dCMP had been lower, the enzyme would probably have gone undetected. The existence of this enzyme was alluded to also by Flaks and Cohen (18). Pursuant to these studies, an analogous increase in deoxycytidylate deaminase activity was found in SP-8 bacteriophage infected Bacillus subtilis (64), but this enzyme was shown later to have properties strikingly different from the E. coli phage-induced deaminase. As a consequence of the discovery of the bacteriophage-induced deoxy­ cytidylate deaminases, an opportunity was presented to examine these enzymes in a manner similar to that described for the animal deaminases and to determine whether the former enzymes are regulated for the same apparent reason as the latter, that of maximizing the efficiency of sub­ strate utilization. Another more pragmatic reason for undertaking this study was the relatively greater source of enzyme available in T-even bacteriophage-infected E. coli. Our first studies were centered, therefore, about the deaminase induced by T2r+ bacteriophage (47, 53). Almost simultaneous investigations were initiated in Maurice Bessman's laboratory (19, 82) with the T4 and T6 bacteriophage-induced deaminases. As first shown by Flaks and Cohen (17), deoxycytidylate hydroxymethylase and thymidylate synthetase are induced within minutes of infection of E. coli by T2r+ bacteriophage. In a subsequent review of his work and others, Cohen (7), discusses the role of some twenty enzymes of this "delayed early" type that are induced on bacteriophage infection of E. coli. Included among these are thymidylate synthetase, deoxynucleotide kinase, deoxycytidylate hydroxymethylase, deoxycytidine 5'triphosphatase, DNA polymerase, dihydrofolic and ribonucleotide reductase, and UDPG-glycosyltransferase. Deoxycytidylate deaminase, similar to the other early enzymes, appeared to be coded for by the bacteriophage genome. More direct proof of the latter statement will be presented later (Section X ) . First, a detailed presentation of the kinetic and physical properties of the homogeneous deaminase will be provided. A. Induction in Bacteriophage-infected Cells; the Titer Effect In our initial studies, difficulty was encountered in inducing consis­ tently high deaminase activities until it was found that the specific activity of deoxycytidylate deaminase, unlike that of the T2 phageinduced thymidylate synthetase and deoxycytidylate hydroxymethylase,

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

201

was dependent on the cell titer. Thus, as shown in Fig. 20, the specific activity of the deaminase varied inversely with the E. coli growth level following T2 bacteriophage infection. A recent study undertaken to pro­ vide an explanation for this selective regulation of enzyme synthesis revealed (Fig. 21) that a similar enhancement could be promoted by a 4-fold dilution of the cells immediately following infection. However, a similar response was not obtained with thymidylate synthetase or with four other phage-induced enzymes (deoxycytidine 5'-triphosphatase, deoxycytidylate hydroxymethylase, deoxyribonucleotide kinase, and dihydrofolate reductase). Protein synthesis inhibitors blocked the in­ crease in deaminase activity, but not rifampicin, an inhibitor of mRNA initiation (91) indicating that the modulating effect is imposed most likely at the translational level. The increase represents true enzyme synthesis, as antibody to the deaminase was found to precipitate all the newly formed radioactive enzyme (Table VII). It will be necessary now to dissect the protein synthetic mechanism to determine where the block resides in this regulatory phenomenon. B. Substrate Specificity of the Bacteriophage-lnduced Enzyme As in the case of the animal deaminase, the phage-induced enzyme is subject to feedback regulation by dTTP and dCTP. A more detailed discussion of this subject is presented in Section VII. However, unlike

FIG. 20. T2r+-induced enzyme synthesis as a function of Escherichia coli growth. The cell titer at which phage infection was initiated was φ, 1.7 X IO8 cells/ml; X, 3.5 X IO8 cells/ml; O, 5 X 108 cells/ml. Specific activity of the synthetase and hydroxymethylase is expressed as nanomoles of product formed in 10 minutes per milligram of protein; that of the deaminase is expressed as micromoles dUMP formed in 10 minutes per milligram of protein. Additional experimental details are given by Maley et al. (47).

202

FRANK MALEY AND GLADYS F. MALEY

dCMP DEAMINASE 200

>-

dTMP .O

SYNTHETASE

8

I50l·

υ 100


HMdUMP (56). It cannot be assured that the restricted deoxycytidylate deaminase produced by the T-even phage provides these organisms with a selective evolutionary advantage, but it does make an interesting, if not provoca­ tive, hypothesis. In this vein, it would be of interest to determine whether a nonlethal deaminase mutant could be found with the capacity to deaminate HM-dCMP. Similar findings on the substrate specificity of the phage deaminase were obtained by Fleming and Bessman (19) with the T6 phage-induced enzyme, although their initial observations on the lack of reactivity of the enzyme with 5-methyl dCMP appeared to be in error (82).

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

205

C. Mutants Deficient in Deoxycytidylate Deaminase The elegant studies of Hall and Tessman (26), in which hydroxylamine mutants of T4 phage deficient in deaminase were produced, revealed this enzyme to be of questionable benefit to the propagation of the normal phage. However, the development of the phage does appear to be im­ paired to some degree, although the restriction in phage propagation could not be directly attributed to the absence of deoxycytidylate de­ aminase. In contrast with these findings, T4 amber mutants have been isolated that induce higher levels of deoxycytidylate deaminase than do the wild-type phage (89). As in the case of the animal deaminase, the question of the signifi­ cance of the enzyme is raised. Is its only role that of providing a supple­ mentary pathway for the synthesis of dUMP, or is the enzyme involved in a more essential task still to be discovered? VII. Regulatory Responses of the BacteriophageInduced Deoxycytidylate Deaminase to Its Metabolic End Products A. Role of the Allosteric Ligands: dCTP, HM-dCTP, and dTTP Although both dCTP .and HM-dCTP activate the deaminase, the most probable natural activator is HM-dCTP since dCTP probably does not accumulate to any extent (Fig. 22). However, in view of the greater commercial abundance of dCTP, this compound has been used in most of the following studies. As with the animal enzyme, a divalent cation, preferably Mg2+, was required with dCTP indicating that the most probable activator is a complex of [Mg^-dCTP* - ] 1 ', where x probably varies from 2 to 4 under the conditions of assay. But unlike the animal enzyme, the activator was found to be essential for the expression of activity at substrate con­ centrations below 2 mili (Fig. 23, curve B). The phage enzyme was also inhibited by p-chloromercuribenzoate, but, as indicated, the inhibition was reversed by a thiol (curve C). In addition, there appeared to be a correlation between the dCMP concentration and the level of dCTP required to achieve a maximal degree of activity. Thus, as shown in Fig. 24, the concentration of dCTP necessary to effect a 50% increase in activity increased about 2-fold in going from 0.25 m l dCMP to 1.0 m l dCMP. It appears that dCMP is able to compete with dCTP for the activator site, a competition that could account for the finding that the enzyme was less active at 1 mikf dCMP than at 0.25 m ¥ dCMP. How­ ever, it should be mentioned that Scocca et al. (82) found that the ac-

206

FRANK MALEY AND GLADYS F. MALEY

4

8 12 MINUTES

16

FIG. 23. Reversal of p-chloromercuribenzoate inhibition of deoxycytidylate de­ aminase by dithiothreitol (RSH) and demonstration of the dCTP requirement. The composition of the reaction mixtures and assay conditions are described by Maley et al. (47). A, dCTP present at zero time; B, dCTP added at 5 minutes; C, dCTP added at 5 minutes and RSH at 9 minutes.

o

100

V ~

1-

> 80 -

3 60 LU

> 40

/

3 20

lu

LU

er

0

0.2

0.4

0.6

0.8

1.0

2.0

4

dCTP (/tf xlO ) FIG. 24. The effect of substrate concentration on dCTP activation. The reaction mixtures contained the following components in a volume of 1 ml: O, dCMP, 0.25 mM; φ, 0.5 mM; Δ , 1.0 mM; 10 mM acetate-phosphate-borate wide-range buffer, pH 8.0 (22); 5 mM dithiothreitol; 1 m l MgCl2; and 0.35 unit deaminase. The reaction was followed spectrophotometrically (64). The levels of dCTP used in each assay are indicated by the symbols on the curves.

207

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

dCTP

0.5 ^ - ^ A

^^JL, \

1

^ U

\C

o 0.4 &

\ *

CM

\ \

\D

Q

O

0.3 -

0

1 2 MINUTES

3

FIG. 25. Reversal of dTTP inhibition by dCTP. The composition of the reac­ tion mixtures and assay conditions are described by Maley et al. (47). At the time indicated by ( j ), 20 nmoles of dCTP were added. A, 1.15 X HT4 M dTTP; B, 7 X 10-5M dTTP; C, 2.3 X 10"5M dTTP; D, 6.7 X 10"6M dCTP. tivity of the T4 phage-induced enzyme was independent of the dCTP level, regardless of the dCMP concentration. Inhibition of enzyme activity was readily effected by dTTP, as shown in Fig. 25, but almost immediate reversal occurred on addition of an O.I5i

no dTTP?

0.02 dCTP

0.03

0.04

(mM)

FIG. 26. Reversal of dTTP inhibition of T2r+ phage-induced deoxycytidylate deaminase by dCTP. The reaction conditions are described by Maley et al. (47).

208

FRANK MALEY AND GLADYS F . MALEY

excess of dCTP. The apparent lack of a lag effect in the activation and inhibition is another indication that the effectors are not reacting chem­ ically with the enzyme. The antagonistic interaction of the allosteric ligands is more clearly seen in Fig. 26 and is reflected in the decreased cooperativity of the curves as the ratio of positive to negative effector is increased. The fact that the curve in the absence of dTTP is still sigmoidal is possibly a response of the deaminase to the heterotropic interplay of dCTP and dCMP, an interplay that culminates in an active form of the enzyme. Because of the complex nature of the reaction, the dependence of activity on the ratio of dCTP to dCMP, and the effect of pH on the sigmoidicity of the reaction, the presentation of interaction coefficients obtained from Hill plots would be of dubious significance, al­ though it is readily apparent in Fig. 26 that dTTP does increase the cooperative nature of the reaction, an effect that will be reflected in the Hill coefficient. B. Allosteric Transitions Associated with the Regulatory Process The cooperative nature of the allosteric interactions described above is believed in most instances to be associated with ligand-induced conformational changes in the affected protein. As a consequence of the alteration in protein conformation, the protein subunits bind more or less tightly to one another, depending on whether the ligand is a positive or negative effector. In the case of deoxycytidylate deaminase, dCTP ap­ pears to enhance subunit binding as evidenced by the preference of the enzyme for the aggregate state in the presence of this nucleotide. It is not possible to state categorically whether the initial conformational transition or the ensuing disaggregation is responsible for the decrease in activity, but it is not unlikely that a combination of both is involved. The influence of ligands on subunit interactions in allosteric proteins has been the subject of considerable conjecture (15, 25, 34, 58, 65, 92), no one theory being completely satisfactory. Our early studies with a 10% purified phage enzyme using sucrose gradient centrifugation was in sub­ stantial agreement with the subunit hypothesis. Thus when the enzyme was centrifuged in the presence of dCTP, an s2o w value of 7.19 S was obtained, which shifted to 4.07 S in the presence of dTTP. Similar find­ ings were obtained on Sephadex G-200 chromatography of the deaminase as shown in Fig. 27, where apparent molecular weights of 134,000 and 47,100 were found for the dCTP and dTTP forms of the enzyme. Evi­ dence is presented ^Section V i l i , D) suggesting that the latter value is not the monomer molecular weight of the deaminase, but a weight aver­ age quantity. While the subunit composition of the deaminase from the chick em­ bryo and T2 phage has been disputed by studies from other laboratories

ALLOSTERIC REGULATION

OF DEOXYCYTIDYLATE

209

DEAMINASES

951 \f-dCMP Deammase+dTTP \Peroxidase

90 85

^Bovine Serum Albumin f Creatine Kinase

801-

^ 75L 70

LactateDehydrogenaseN d CPM Dea minase Alcohol Dehydrogenase \ +dCTP

65 60 10

30

50

100

3 0 0 5 0 0 1000

M O L E C U L A R WEIGHT x IO* 3

FIG. 27. Sephadex G-200 chromatography of T2r+ phage-induced deoxycytidylate deaminase in the presence of 40 μΜ dCTP and dTTP. A detailed description of the methods used are presented by Maley et al. (4?).

(78, 82), we have consistently obtained multiple forms, even with a sam­ ple of the T4 deaminase kindly sent to us from Bessman's laboratory. Similar findings were obtained with the T4 phage-induced deaminase prepared in our laboratory. Convincing evidence for the oligomeric structure of the pure Τ2Γ+ phage-induced deaminase will be presented below, evidence that supports our earlier observations on the aggregate nature of the enzyme. VIII. Physical and Chemical Properties of the T2r Bacteriophage-lnduced Deoxycytidylate Deaminase A. Purification to Homogeneity The purification of reasonable quantities of the deaminase to homo­ geneity was made possible through the excellent cooperation and faciliTABLE IX PURIFICATION

OF T2r + BACTERIOPHAGE-INDUCED

Step 1. 2. 3. 4. 5. 6. 7. β

Crude extract Streptomycin autolyzate Ammonium sulfate DEAE-cellulose Phosphocellulose Sephadex G-200 Isoelectric precipitation

DEOXYCYTIDYLATE

DEAMINASE0

Total protein (gm)

Specific activity (units/mg)

Yield

294 76 28 6.95 0.24 0.13 0.03

0.13 0.44 1.35 5.10 95.5 174 430

100 88 99 75 61 57 34

A detailed purification procedure has been published (47a).

(%)

210

FRANK MALEY AND GLADYS F. MALEY

ties of the New England Enzyme Center. The overall purification scheme is presented in Table IX and represents better than a 3000-fold purifi­ cation. A detailed description of the purification has been published (47a). One of the major problems encountered, common to the iso­ lation of most phage-induced enzymes, was the removal of large quanti­ ties of interfering nucleic acid. This problem was solved by streptomycin precipitation, autolysis, and sonication prior to chromatography on DEAE-cellulose. Unlike the chick embryo enzyme, which was purified to homogeneity by a novel reversible activation step, the phage enzyme did not lend itself to this technique. However, the last step in the proce­ dure, dialysis at pH 7.8, yielded the enzyme as a semicrystalline precipi­ tate, and, as shown in the polyacrylamide gel pattern of Fig. 28, most of the contaminating protein remained in the supernatant fraction. To solubilize the precipitated enzyme, high salt and thiol were necessary (0.2 M phosphate plus 0.1 M 2-mercaptoethanol, pH 7.1). The inference that the isoelectric point is about pH 7.8 was supported by the isoelectric focusing pattern presented in Fig. 29. In addition to polyacrylamide gel electrophoresis (Fig. 28), the question of homogeneity was answered by

FIG. 28. Polyacrylamide gel electrophoresis of deoxycytidylate deaminase pro­ tein fractions at various stages of purification. The gels and the protein concentra­ tions are from left to right: P-cellulose fraction, 18 μ&; Sephadex G-200 fraction, 20 /ig; supernatant fraction following isoelectric precipitation of the deaminase, 70 /ig; highest specific activity deaminase fractions, 10 /ig and 25 /xg.

211

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

15

20

TUBE

25

30

45

50

NUMBER +

FIG. 29. Isoelectric focusing of T2r bacteriophage-induced deoxycytidylate deaminase with a pH 6 to 8 ampholine gradient. Homogeneous protein (120 /ig or 55 units) in a volume of 2 ml was applied to the center of the column. The methods used are similar to those described by Durham and Ives (18).

sedimentation velocity centrifugation (Fig. 30) where the enzyme is seen to migrate as a single component. With the photoelectric scanner system (80), an s2V^% value of 6.05 ± 0.05 S was obtained. B. Molecular Weight of the Oligomer and Protomer Sedimentation equilibrium analysis of the deaminase provided addi­ tional proof for the homogeneity of the enzyme protein as evidenced by the lack of deviation of the line in Fig. 31A. The data from the slope yielded a molecular weight of 124,000 ± 1800 daltons. When the enzyme was centrifuged in 6 M guanidine · HC1 plus 0.1 Af 2-mercaptoethanol, a single component with a molecular weight of 20,200 ± 100 daltons was obtained (Fig. 31B), indicating that the enzyme is composed of six subunits. Further evidence in support of the hexameric subunit composition of the deaminase was provided by the sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis procedure of Weber and Osborn (90) (Fig. 32). As seen in the log molecular weight versus mobility of Fig. 33, the estimated molecular weight of the disaggregated enzyme protein is in excellent agreement with that obtained by the more sophisticated sedi­ mentation equilibrium analysis.

212

FRANK MALEY AND GLADYS F. MALEY

H#

D

D

FIG. 30. Ultracentrifugation of T2r + bacteriophage-induced deoxycytidylate deaminase. The protein solution (0.6% in 0.1 M potassium phosphate, pH 7.1), plus 0.1 M 2-mercaptoethanol was centrifuged at 59,780 rpm in a 12-mm cell of an AN-D rotor with the Spinco Model E ultracentrifuge. Exposures were taken with a bar angle of 60° after (A) 5 minutes; (B) 21 minutes; (C) 37 minutes; (D) 53 minutes. Temperature: 25°C (47a).

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

213

FIG. 31. Molecular weight determination of T2r+ bacteriophage-induced deoxycytidylate deaminase by sedimentation equilibrium using the photoelectric scanner system (80). The ordinate presents the logarithm of the recorder deflection (mm) on the left and the abscissa, the square of the distance (cm2) from the axis of rotation. The solvent in (A) was 0.2 M potassium phosphate, pH 7.1, and 0.1 M 2-mercaptoethanol; in (B), 6Af guanidine ■ HC1, 0.2 M potassium phosphate, pH 7.1, and 0.1 M 2-mercaptoethanol. The concentration of protein in each cell was 0.25 mg/ml.

Another method useful in clarifying the subunit composition of the enzyme is the dimethyl suberimidate procedure of Davies and Stark (10). As shown by these authors, after treatment of a multicomponent protein with this cross-linking agent, there is excellent correlation be­ tween the number of bands obtained on SDS-polyacrylamide gel electrophoresis and the identity of the protein subunits. Thus if their thesis is correct and the deaminase is composed of six identical subunits, only six protein bands should be obtained. The results presented in Fig. 34 tend to support this concept, as does the presence of a single band on SDSpolyacrylamide gel electrophoresis (Fig. 32). Similar studies in progress, such as fingerprint and end group analysis, will be necessary to validate these findings. Preliminary findings on the number of fragments formed after cyanogen bromide treatment revealed only three, a result consistent with the methionine content of the protein. If the subunits were not iden­ tical, more than three bands would be anticipated. C. Diffusion Coefficient and Stokes Radius Calculation of the Stokes radius of the monomer and hexamer using the gel procedure of Siegel and Monty {85) yielded values of 7.5 Â and

214

FRANK MALEY AND GLADYS F. MALEY

BSA CPase cyt c

T2d

tryp

T2d cyt c

FIG. 32. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis of various proteins in comparison to deoxycytidylate deaminase. The proteins used are BSA, bovine serum albumin; CPase, carboxypeptidase A; cyt c, cytochrome c; tryp, trypsin; T2d, deoxycytidylate deaminase. The conditions employed are similar to those described by Weber and Osborn (47b, 90).

44 Â, respectively. Although this method has certain inherent limitations (-0, the results are consistent with the combination of six protomers to form an. oligomer. The Stokes radius of the latter is also in the range described for aldolase, 46 A, a protein with a molecular weight of 150,000. Using the determined s and M values for the T2r+ bacteriophageinduced deoxycytidylate deaminase in the Svedberg equation:

a value of 4.61 X 10~7 cm2 sec"1 was obtained for D20. When D20 was substituted in the equation relating Stokes radius to diffusion coefficient: =

kT 6πηΏ

where a = Stokes radius, k = Boltzmann constant, T = absolute tem­ perature at 20°, and η = viscosity of the medium at this temperature, a

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE

DEAMINASES

215

.OVALBUMIN

UJ

^ · CARBOXYPEPTIDASE A

ce < _i

J"RYPSIN

o

UJ _l

o Έ

CYTOCHROMEc 1.0

FIG. 33. Molecular weight determination of deoxycytidylate deaminase by means of sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The mobility of thé proteins in Fig. 30 is plotted versus the log of molecular weight. BSA, bovine serum albumin (47b).

î2f* #04ψ ψψβφίί*ΐ$*ΐφ

■^AMÉÉÉ ^

■

o




I-




14 20 30 50

90 ' 1751 330,

40 50 60

110 160 190

(MM)

dTTP:dCTP HM-dCTP (MM) (average)

23

90^

65^

10

pH a

dTTP* (MM)

9V 11< 18|

dTTP: HMdCTP (average)

4 6 10 20

32 >

40 50 100

52 80 185

1.6

100 120 160

180 206 264

1.7

1.8

° The reaction mixtures contained the following components in a final volume of 1 ml: 0.5 mM dCMP; 5 m l dithiothreitol; 1 m l MgCl2; 10 m l acetate-phosphateborate wide-range buffer (22); 0.3 unit of enzyme and concentrations of dCTP, dTTP, and HM-dCTP as indicated. b Concentration of dTTP required to effect a 50% inhibition at the indicated dCTP level. Data from Maley et al. (47a).

222

FRANK MALEY AND GLADYS F. MALEY

X. Studies on the in Vitro Synthesis of BacteriophageInduced Deoxycytidylate Deaminase The apparent induction of enzymes on the introduction of a virus genome into a cell is often associated with the premise that the informa­ tion for the enzyme structural gene resides in the virus DNA. Since the use of inhibitors of transcription and translation to prevent the appear­ ance of induced enzymes does not constitute direct proof for the intro­ duction of new genes into a cell, more definitive results might be ob­ tained by the use of an in vitro protein-synthesizing system with viral DNA as template. This technique has been used successfully in several instances already for the synthesis of ß-galactosidase {12), phage-directed enzymes {21, 24, 81), and rabbit hemoglobin {63). While our studies were in progress, the purported demonstration of T4-induced deoxycytidylate deaminase synthesis appeared {81). Unfortunately, the effective activity measured in this system was only twice background. However, in support of these studies, no activity was produced with DNA sources that were known not to program for deoxycytidylate deaminase. We have improved on this system greatly now, and yields of deaminase activity 300 times greater than those reported earlier (81) have been obtained. As indicated in Fig. 39, there was a lag of about 8 minutes 200 'S»

=3

t too > IO


-

; Γ"

ò

'c

50

N Z L±J

0

2

I/,,,."

CP

\ e c LU

ί

0. X

i η

16

24

32

40

ÜJ

Z

o =) LU

MINUTES FIG. 39. Kinetics of the in vitro synthesis of deoxycytidylate deaminase di­ rected by T2r+ phage DNA. The system used was similar to that described by DeVries and Zubay (12). The incubation mixtures (0.5 ml) contained 3.75 mg S-30 protein and 37 /ig T2r+ DNA. At the times indicated, 25-μ1 aliquots were removed and assayed as described by Gold and Schweiger (24). Leucine incorporation into protein was determined in parallel with the enzyme assay.

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES

223

before deoxycytidylate deaminase appeared with the T2r + phage DNA as the template, although amino acid incorporation began almost immedi­ ately. This delay could be eliminated by using mRNA isolated from 8-minute T2 phage-infected cells (Fig. 40). The template capacity of the mRNA varied with the time of isolation from the infected cells (Fig. 41), a result anticipated from the time course of appearance of phage-induced enzymes (7). A plateau of deaminase activity was reached in the infected cells at about 11 minutes, but as shown, the mRNA that programs deaminase synthesis decreased rapidly after 5 minutes. The enzyme pro­ tein appears to be more stable than the mRNA, the latter with a T% of about 3.5 minutes. Somewhat similar results were reported recently by Sakiyama and Buchanan (78) for T4 phage-induced deoxynucleotide kinase, and earlier by Bose and Warren (5), for thymidylate synthetase. The deaminase does not appear, however, to respond to the allosteric effectors, dCTP and dTTP, in the manner anticipated, i.e., dCTP does not activate and dTTP does not inhibit. The latter effect is complicated though by the finding that dTTP can transfer its phosphorus to dCMP yielding dCTP. Because of the numerous proteins and cofactors involved in the synthesis, it would be premature to speculate on these findings. Nonetheless, the possibility that the enzyme is altered at some later stage of phage development, or by a still unknown cofactor, should not be excluded. Although somewhat different in its regulatory response from the iso-

io

20

30

MINUTES

FIG. 40. Kinetics of the m vitro synthesis of deoxycytidylate deaminase directed by T2r+ phage mRNA. The conditions were similar to those in Pig. 39, except that ribonucleoside 5'-triphosphates were deleted and 500 /ig of RNA were substituted for the DNA.

224

FRANK MALEY AND GLADYS F. MALEY

E ω

Έ

160 J .

>

i-

o

< < Ld 00

< < Id Q

2

5

8

II

14

17

o I

MINUTES

FIG. 41. Time course of synthesis of deoxycytidylate deaminase and its mRNA template following T2r+ phage infection. Enzyme activity was followed at various times after infection by sonication of the cells ( # ) . Deaminase activity programmed by mRNA at various times after infection was measured by isolation of the RNA and using it as a template in the in vitro synthesizing system (12).

lated T2 phage-induced deaminase, the in vitro enzyme is completely in­ hibited by antibody to the latter, thus establishing the identity of the two. XI. Conclusion The data presented in this article demonstrate that despite the wide gulf in phylogeny that separates chick embryo deoxycytidylate deaminase from the T2r + phage-induced enzyme, evolution has not imposed striking differences on their regulatory or catalytic properties. Although the phage enzyme is more restricted in its capacity to deaminate analogs of dCMP with substituents on the 5-position of the pyrimidine ring, an apparent logical outgrowth of the metabolic requirements of the T-even phages, the catalytic potential of both chick and phage enzymes is : determined by the interplay of the end products of pyrimidine deoxyribonucleotide biosynthesis. Thus, in the case of the chick embryo deaminase, the pool sizes of the pyrimidine deoxyribonucleotides are probably regulated as shown in Fig. 42. An elevation of the dTTP pool should lead to an impairment in dTTP synthesis due to its feedback inhibition of sites 3 and 4 and, as a consequence, more deoxycytidine nucleotides should be available for dCTP synthesis. With the resultant increase in the dCTP pool, an impair­ ment at site 2 would result (Jfê) while the inhibition at site 4 should be

ALLOSTERIC REGULATION OF DEOXYCYTIDYLATE DEAMINASES d-URIDINE

225

dTMP

dGTP

J

FIG. 42. Regulation of pyrimidine deoxyribonucleotide metabolism in animal tissues by dCTP and dTTP.

relieved (54), enabling the balance between dCTP and dTTP to be main­ tained. Not included in this figure is the major route for the de novo synthesis of pyrimidine deoxyribonucleotides, the CDP -» dCDP ribonucleotide reductase reaction, which appears also to be regulated by dTTP (69). Since all the above channels of supply for the production of pyrimidine deoxyribonucleotides are regulated to some degree by dCTP and dTTP, a high degree of efficiency in the maintenance of the dCTP and dTTP pool sizes should prevail. Similarly, as shown in Fig. 22, the flow of pyrimidine intermediates through HM-dCTP and dTTP in T-even phage-infected E. coli should be regulated efficiently by the interplay of these nucleotides on enzymes involved in their synthesis. Unfortunately, a more comprehensive analy­ sis cannot be presented as the only phage enzyme known to be affected by HM-dCTP and dTTP is the T-even phage-induced deoxycytidylate deaminase. Although a phage ribonucleotide reductase system is induced (8), its regulation has not been studied thoroughly enough to comment on, except that it appears to be regulated differently than the E. coli reductase (3). The studies presented in Section IX demonstrating the pH dependence of the allosteric regulation of deoxycytidylate deaminase suggest that a major role of this enzyme is to maintain a balanced level of HM-dCTP and dTTP by competing with deoxycytidylate hydroxymethylase for dCMP. The influence of HM-dCTP and dTTP on deoxy­ cytidylate deaminase no doubt contributes to this process. Whether enzymes from such divergent sources as described in this article employ the same mechanisms of cooperative feedback regulation must await a comparative analysis of the phage enzyriie with pure chick embryo deoxycytidylate deaminase, a study currently in progress.

226

FRANK MALEY AND GLADYS F. MALEY ACKNOWLEDGMENTS

We would like to express our appreciation to Mr. Don U. Guarino and Mrs. Judith Reidl for their excellent technical assistance and to Dr. Robert MacColl for his aid with the ultracentrifuge analyses. The studies of Dr. Robert B. Trimble on the in vitro synthesis of the deaminase are gratefully acknowledged. Support for this work was provided by grants from the United States Public Health Service (CA-06406), the National Science Foundation (GB-27598), and the American Heart Association. REFERENCES

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The Citrate Enzymes: Their Structures, Mechanisms, and Biological Functions I

P A U L A.

SRERE

I I I I I

Veteran's Administration Hospital and Department of Biochemistry University of Texas Southwestern Medical School Dallas, Texas

I. Introduction A. Three Citrate Enzymes B. Chemistry of Citrate C. Biochemistry of Citrate D. Common Enzyme Mechanisms II. Citrate Lyase (Citritase) A. Distribution B. Physical Properties C. The Reaction III. Citrate Synthase A. Distribution B. Physical Properties C. Chemical Properties of the Enzyme and Active Site . . D. Thermodynamics E. Kinetics F. Effectors of Citrate Synthase Activity G. Mechanism of Citrate Synthase H. Biological Aspects of Citrate Synthase IV. ATP Citrate Lyase (Citrate Cleavage Enzyme) . . . . A. Distribution B. The Reaction C. Physical Properties D. Chemical Properties E. Mechanism Studies F. Physiological Aspects of ATP Citrate Lyase V. Overview References

229 229 230 234 235 242 242 242 242 245 245 248 249 252 253 254 257 262 266 266 266 269 269 270 273 276 276

I. Introduction A. Three Citrate Enzymes There are three enzymes that catalyze lyase reactions on citrate to yield a C2 and a C4 unit: 1. Citrate lya^e (citrate oxaloacetate-lyase EC 4.1.3.6.) 229

230

PAUL A. SRERE M2+

Citrate 3 - = acetate" + oxaloacetate 2 -

2. Citrate synthase [citrate oxaloacetate-lyase (CoA-acetylating) EC 4.1.3.7.] C i t r a t e 3 - + CoA + H + = acetyl-CoA + oxaloacetate 2 - + H 2 0

3. ATP citrate lyase [ATP:citrate oxaloacetate-lyase (CoA-acetylat­ ing and ATP-dephosphorylating) EC 4.1.3.8.] M2+

C i t r a t e 3 - + CoA + A T P 4 - = acetyl-CoA + oxaloacetate 2 - + A D P 3 - + Pi 2 "

Citrate lyase has been reported only in certain bacteria, citrate synthase has been found in all cells examined for it, and ATP citrate lyase has been found only in eukaryotic cells. The reaction catalyzed in common by these three enzymes is an aldol type, that is the reversible formation of a C—H bond from a C—C bond. It is possible that there exist some catalytic similarities in the three enzymatic reactions so that comparison of the results obtained with each enzyme may be useful in understanding the others. The citrate enzymes comprise a unique biological system in which three enzymes catalyze the same bond-breaking-making reaction. The fact that the simplest of these reactions, that catalyzed by citrate lyase, occurs in the most primitive organisms and that the most complex, the ATP citrate lyase reaction, occurs in the most recently evolved orga­ nisms, suggests a possible evolutionary relation between these enzymes. B. Chemistry of Citrate Citric acid has pK values (0 ionic strength at 25°C) of 3.13, 4.76, and 6.40 {10) y while at 0.1 ionic strength at 25°C they are 2.88, 4.36, and 5.84 (221). Cellular pH is assumed to be 7.4 and the ionic strength is not less than 0.3, so that in most biological circumstances we are deal­ ing primarily with the trianion. Titration studies of Martin (133) indi­ cated that the central carboxylic acid group is the most acidic and that the symmetrical monoionized species is predominant. Analysis of X-ray diffraction data indicates also that the central carboxyl group is ionized in solid sodium dihydrogen citrate (76). Of great but unassessed biologi­ cal importance is the chelating ability of citrate. The log of the forma­ tion constant of magnesium citrate is 3.2, that for Fe2+ citrate is 2.15, while that of calcium citrate is 3.22 (33). Tate et al. (221) have indi­ cated that the log of the stability constant of MgHcit is 1.84 at 0.1 ionic strength and 25°C while that of Mg citrate - is 3.73 under similar condi­ tions. Johnson has shown (106) by X-ray crystallography that in mag-

THE CITRATE ENZYMES

231

nesium citrate decahydrate each citrate chelates to one magnesium atom in a tridentate manner through one terminal carboxyl, the central carboxyl and the hydroxyl group oxygen. This indicates that the citrate chelates are extremely stable and citric acid has been used as a chelator in many industrial processes. Nuclear magnetic resonance (NMR) studies of citric acid by Loewenstein and Roberts {126) showed a quadruplet (a strong doublet and two weak satellites) due to the nonequivalence of the méthylène hydrogens. X-Ray crystallographic studies show that various forms of citric acid and its salts have a remarkably similar conformation. The molecule is invariably fully extended with the central carboxyl group and the adja­ cent hydroxyl always coplanar (Fig. 1) (74). This constant conforma­ tion of the various citrate molecules has been used by Glusker to aid in depicting the mechanism of the aconitase reaction (7%). A similar ap­ proximation as to the geometry of the citrate site on other citrate enzymes could possibly serve as a useful model. Chemical reactions with citric acid cannot distinguish the two carboxy méthylène (CH 2 COOH) groups in the molecule. Ogston {146) recog-

FIG. 1. The shape of magnesium citrate as determined by X-ray crystallography by Johnson (106). This molecule was redrawn according to Glusker (74). In this configuration the pro-S portion of the molecule is at the bottom, just as it is drawn in Fig. 3. O = carbon; ® = oxygen; 0 = hydrogen.

232

PAUL A. SRERE Pro-3R 1

Pro-3S

2

3 4 5 OH HOOCH2C. T XH2COOH I COOH FIG. 2. Visualization of citric acid according to Hanson and Rose (86).

nized the asymmetry inherent in this type (Caabc) of compound and pointed out that enzymes may be able to distinguish the two identical groups. The elaboration of the stereochemistry of citric acid has been well documented and reviewed (11, 153), and I will present here only a summary of the most recent data and nomenclature. B

coo OH

H/

c

COO

COO"

Pro-3i?

ooc Pro-3S

Pro-3S KR

Pro-3S

CH2COO"

FIG. 3. Four representations of the citrate trianion. (A), (B), and (C) are oriented the same. (A) Space-filling model. (B) Three dimensional drawing, thin straight lines in the plane of the page. Solid triangular lines are above the plane and dotted lines are below the plane of the page. (C) A Newman projection of citrate looking down the 3 ^ 4 bond. Carbon 6 is the unnumbered carboxyl carbon on carbon No. 3. (D) A Fisher projection of citrate numbered and named according to Englard and Hanson (59).

THE CITRATE ENZYMES

233

Hanson and Rose (86) have visualized the asymmetry of citric acid as seen in Fig. 2. The left-hand CH 2 COOH is not superimposable on the right-hand carboxymethylene group. They are thus not identical and can be distinguished. There are three prochiral centers in citric acid, and they are labeled by Englard and Hanson (59) as shown in Fig. 3. Unfortunately, this new numbering system, consistent with the R-S rules, reverses a numbering system which has been widely used, so that great care must be taken in identifying the carbons of the molecule. Since commercially available labeled citrates are still labeled by the old system, I will use both sys­ tems, but I will always state when old or new numbering is being used. In both numbering systems, carbons 3 and 6 retain the same number. In the old system, carbons 1 and 2 were derived from the carboxyl and methyl groups of acetate, respectively. In the new numbering system carbons 4 and 5 are derived from the methyl and carboxyl groups of acetate, respectively. Hendrickson and Srere (92) have presented still another way of visual­ izing the asymmetry of citrate from models of its metal chelates. Metal chelates of citrate have the hydroxyl group, the middle carboxyl, and one terminal carboxyl coordinated to the metal ion. Such a complex is asymmetric and can exist in two isomerie forms, depending on which terminal carboxyl group participates in the complex. It can easily be seen from the molecular models (Fig. 4) of these two isomers that they are readily distinguishable and are nonsuperimposable mirror images of each other. These two isomers can be designated as having the R and S configurations according to the Cahn-Ingold-Prelog system (31).

FIG. 4. Molecular models of citrate-metal chelates (92). (A) θ-metal citrate. (B) R-metal citrate.

234

PAUL A. SRERE

C. Biochemistry of Citrate Citrate is a normal constituent of all living cells but is found in espe­ cially high concentrations in certain citrus fruits (211), bone (48), semen (222), and mammary glands (222). The high concentration in these tissues ( > 1 0 roM) has not been explained although there has been some speculation that its presence in bone is related in some way to its ability to chelate Ca2+. Another unexplained biological phenomenon is the ability of certain molds to ferment glucose almost quantitatively to citrate. Even though this process is of great commercial importance, no well-docu­ mented mechanism is available to explain the phenomenon. The importance of citric acid in terminal stages of carbohydrate metabolism was established by Krebs and Johnson (120) with the eluci­ dation of the citric acid cycle. In addition to this role three other meta­ bolic functions for citrate are (185) : a source of reducing power; a source of acetyl groups for biosynthetic pathways; and a controlling (activating or inhibiting) substance for a number of enzymes. Aconitase is the only other enzyme catalyzing a chemical conversion of citrate, although for certain metabolic purposes it is useful to con­ sider the isocitrate dehydrogenases as citrate enzymes since through them citrate serves as a hydrogen donor for biosynthetic reactions. The oxidation of citrate in the mitochondria leads to the production of energy, not to the production of reducing hydrogen needed for biosynthetic purposes. Since the reducing potential of mitochondrial metabolism is largely converted to energy and since reductive syntheses usually involve NADPH and occur in the cytosol it seems probable that citrate is metabolized via aconitase and isocitrate dehydrogenase found in the cytosol to supply hydrogen needed for cytosolic syntheses. Another major function for citrate lies in its ability to supply acetyl groups for biosynthetic purpose in a reaction catalyzed by ATP citrate lyase. Most acetyl-CoA in eukaryotic cells is generated in the mito­ chondria. A mechanism is needed to transfer it to the cytosol. The ATP citrate lyase is found exclusively in the cytosol and its function is prob­ ably related to its ability to form cytosolic acetyl-CoA and oxaloacetate. The citrate synthase and the ATP citrate lyase may act together as an acetyl group shuttle. The function of this system is consistent with the intracellular and tissue distribution of the two enzymes. High concen­ trations of ATP citrate lyase occur in those tissues that have a high capacity for acetyl group utilization (177). These roles will be discussed later in this article. The important observation that citrate can act as an activator for

235

T H E CITRATE E N Z Y M E S

acetyl-CoA carboxylase (132), an enzyme necessary for lipid synthesis, uncovered a third biological role for this compound. Citrate alters the activity of at least two other enzymes, phosphofructokinase (147) and isocitrate dehydrogenase (162). The phosphofructokinase reaction is known to be a rate-limiting step in glycolysis, hence this interaction may have important implication in the control of metabolism. A detailed examination of possible metabolic interactions of citrate has been pre­ sented in a symposium on citrate metabolism (77). Thus, in those instances where its metabolism has been examined, citrate is seen to be a multifunctional component whose concentration is undoubtedly under stringent regulation. This article will attempt to present all that is known concerning three enzymes involved in the direct formation and breakdown of citrate. It is possible that studies on the enzymes will not only be useful in elucidating their mechanisms and those of other similar aldolases, but perhaps will be useful in understand­ ing the metabolic roles of citrate. D. Common Enzyme Mechanisms 1. ENOLASE ACTIVITY

All three citrate lyases must be able to catalyze at least two separate reactions. One reaction is an enolase activity CH3COO- = -CH2COO- + H+ The other is a ligase activity H+ + -CH2—COO" + 0=C—COO" I

CH2COOI

CH2—COO- = HO—C—COO-

I 2—COOCH It has been postulated that in acetyl-CoA the thioester bond enhances the electrophilic nature of the carbonyl group and makes the proton on the methyl group more acidic and thereby easier to enolize than the proton on an acetate molecule (104)- Since citrate lyase must catalyze a similar enolization on acetate, a reaction that probably occurs on the acetyl thioester which constitutes part of the active site (see below). In any case this proton removal is considered difficult in chemical aldol reactions and often represents a rate-determining step. The citrate enzymes cannot by themselves catalyze an exchange be­ tween the protons of the methyl group of the acetyl moiety and the water of the medium. Eggerer (51) showed, however, that L-malate (S-malate) was able to induce an enolase activity, measured as an ex-

236

PAUL A. SRERE

change of T 2 0 into acetyl-CoA in pig heart citrate synthase. This activ­ ity was less than 1% of the overall enzymatic activity, but the specificity of the inducing substance [ü-malate (Ä-malate) and other compounds did not work] indicated that this exchange was related to the enzyme activity. Eggerer postulated as had Bove et al. (18) and Marcus and Vennesland (131) that the carboxyl group of oxaloacetate (or the inducer) acts as a base to aid in the enolization reaction. Srere (187) re­ peated Eggerer's experiments by following the exchange from D 2 0 into acetyl-CoA and found that in addition to L-malate, α-ketoglutarate was also able to induce an enolase activity in citrate synthase. On the basis of experiments which indicated that oxaloacetate, L-malate, and a-ketoglutarate could bind to citrate synthase and protect it against ureainduced unfolding (186), Srere postulated an enzyme conformation change to explain the enolase activity. Subsequent studies on the stereo­ chemistry of this reaction favors this explanation over the one postu­ lated by Eggerer. When oxaloacetate and acetyl-CoA are present with citrate synthase, no exchange of the methyl protons of acetyl-CoA can be found, an observation indicating that the condensation step is much more rapid than the back reaction of the enolization step. The fact that the removal of the proton is the rate-limiting step is also indicated by the experiments of Kosicki and Srere (119) which showed a kinetic isotope effect of about 1.4 when D 3 acetyl-CoA was used as the substrate. Larger kinetic isotope effects have been calculated by Klinman and Rose (109) to explain the distribution of 3 H in the products when the stereochemistry of the acetyl protons removal was studied (see below). No data are available concerning the enolase activity of citrate lyase. Using NMR techniques, we have been unable to demonstrate a catal­ ysis of exchange between acetate protons and D 2 0 either in the presence of enzyme alone or in the presence of L-malate. The enolase activity, if any, must be investigated using the more sensitive tritium exchange techniques. The ATP citrate lyase catalyzes a rapid exchange between the pro­ tons of the methyl group of àcetyl-CoA and protons of the medium when oxaloacetate is present (46). This exchange has been followed by using 3 H-labeled acetyl-CoA and measuring the production of 3 H-labeled H 2 0 and by following the exchange of 2 H 2 0 for the protons in the methyl group of acetyl-CoA using NMR techniques described earlier (187). Comparable results were obtained with both methods. The rate of ex­ change is about % ^max of the enzyme in the direction of citrate forma-

THE CITRATE ENZYMES

237

tion. The exchange is not affected by any combination of the remaining substrates, ADP, Pi, or Mg2+. Neither L-malate, D-malate, nor a-ketoglutarate can replace oxaloacetate as a cosubstrate in this reaction. The oxaloacetate exchange could be due to a rate-limiting enolization fol­ lowed by a citrate formation (either citryl-CoA or citryl enzyme) and breakdown or perhaps a conformation change in the enzyme similar to that occurring in the citrate synthase reaction. The condensation of "enol" acetate (the carbanion) with oxaloacetate is a common reaction in chemistry and except for the stereochemistry should present no diffi­ culties. As early as 1891, Claisen and Hori (35) showed the chemical synthesis of aconitic derivatives by the aldol condensation of oxalo­ acetate esters and acetic acids. 2. STEREOCHEMICAL FEATURES

Metabolic experiments using rat tissues {127, 154) supported Ogston's (1 46) contention that citrate was labeled and metabolized in an asym­ metrical manner. At the enzyme level Stern and his co-workers {145, 209, 210) showed that citrate labeled in the carboxyls derived from oxaloacetate was handled asymmetrically by crystalline pig heart citrate synthase. Srere and Bhaduri {196) established that chicken liver ATP citrate lyase had the same stereochemistry as pig heart citrate synthase. Wheat and Ajl (241) showed that the citrate synthase and citrate lyase of Escherichia coli had identical stereospecificity. Stern et al. (207) showed pig heart citrate synthase, and Klebsiella aerogenes citrate lyase had the same stereospecificity. Although such experiments indicated that these enzymes had the same stereospecificity, they did not indicate what the absolute stereospecificity of citrate formed was. The absolute stereochemistry of citrate biosyn­ thesis was established by Hanson and Rose (86). Their results indicate that the carboxymethylene group derived from acetyl-CoA was in the si attack position of citrate. That is, the acetyl moiety is added to the si side of the oxaloacetate molecule (Fig. 5). With the exception of citrate synthase from certain anaerobes (78-80, 142, 206), it is assumed that all other citrate synthases have a si attack specificity. Citrate (re) synthases, that is enzymes in which the acetylCoA is added to the re face of oxaloacetate, have been found only in certain anaerobic bacteria. These represent the only known divergence from the stereospecificity of all citrate enzymes. Designating these enzymes as either re attack or si attack enzymes is of course only a shorthand method of nomenclature. Hanson and

238

PAUL A. SRERE

Hirschmann (85) have suggested that an unequivocal naming of these enzymes would be useful and would name them as follows E C number

Systematic name

4.1.3.6

Citrate oxaloacetate-lyase (pro-3-£-CH 2 COO- -> acetate) Citrate oxaloacetate-lyase (pro-3-£-CH 2 COO- -> acetyl-CoA) or (pro-3-Ä-CH 2 COO- -» acetyl-CoA) Citrate oxaloacetate-lyase (pro-3-Ä-CH 2 COO- -+ acetyl-CoA, A T P dephosphorylating)

4.1.3.7

4.1.3.8

Trivial name Citrate (pro-S-S) lyase Citrate (si) synthase Citrate (re) synthase A T P citrate (pro-S-S) lyase

In this article EC 4.1.3.6 will be referred to only as citrate lyase and EC 4.1.3.8 will be called ATP citrate lyase since only pro-3-S enzymes are known. In the case of EC 4.1.3.7 where both re and si attack enzymes 4

3 2

COO -

Hs \'c/ H/\/l c

/

ooc

re Face

OOC

\ /HR c \/\H s C

\ coo

si Face

FIG. 5. The faces of oxaloacetate. Top: Space-filling model of oxaloacetate (re face is u p ) . Bottom: Representation of both faces of oxaloacetate.

239

THE CITRATE ENZYMES "°0CvC=0 "OOCH2C^

τ^ / x o o I

"OOCCH2 "OOCv\ /OH T

I iD

2

3

K

H COO 5 FIG. 6. Inversion of configuration of protons on acetate during condensation with oxaloacetate.

are known, then citrate (si) synthase will be referred to mainly as citrate synthase while the re attack citrate synthase will always be designated as citrate (re) synthase. The other aspect of stereospecificity, that is the stereorelationship between the C-C and C-H bond making events, has been elegantly eluci­ dated in several laboratories (52, 109, 157). Using specifically labeled B- 2 H, 3 H acetate and S-2H,3H acetates, Eggerer et al. (52), Rétey et al. (157), and Klinman and Rose (109) showed that all the citrate enzymes, citrate (si) synthase, citrate (re) synthase, ATP citrate lyase and citrate lyase proceeded with inversion of configuration at the methyl group of the acetyl moiety (Fig. 6). This stereochemical relation rules out the possibility of the carboxyl group of oxaloacetate acting as an internal base for the enolization reaction, a process that requires retention of configuration. Rose (159) has pointed out that the aldolases which cata­ lyze an enolization without a cosubstrate proceed with retention of con­ figuration. In the case of the citrate enzymes, malate synthase, and isocitritase, a cosubstrate is required for enolization and inversion is common to all. Rose (159) also notes that a two-base mechanism, one aiding proton removal from one side, the other donating a proton, be­ comes a likely mechanism for this reaction. It is of interest to consider the stereochemistry of the two citrate de­ rivatives, fluorocitrate and hydroxycitrate. There are, of course, many isomers of each of these compounds, but we shall consider just two. Fanshier et al. (64) showed that when fluoroacetyl-CoA is used in place of acetyl-CoA as a substrate in the reaction catalyzed by pig heart citrate synthase, a single isomer of fluorocitrate is produced. This pro­ duct is inhibitory in the aconitase reaction. This isomer was subjected to crystallographic analysis by Carrell et al. (32) and shown to be (2Ä,3Ä)-2-fluorocitrate (chemical numbering) or (pn)-(4i?)-4-fluorocitrate (parent numbering).* If the steric course of the reaction * The numbering system devised to number citrate on the basis of isotope substitution leads to an apparent inconsistency with accepted chemical numbering practices when used with chemical derivatives of citrate. Thus chemically the fluorocitrate is correctly numbered as (2Ä,3i2)-2-fluorocitrate. However, since the

240

PAUL A. SRERE

coo"

COO"

H—C—H OH OOC—C-OH

V coo

F—C—H COO"

coo

COO

"OOC

(A)

OH

(/w)-(4A)-4-Fluorocitrate FIG.

7 (A).

of fluoroacetyl-CoA is the same as that for acetyl-CoA (inversion) then Klinman and Rose (109) have pointed out that only the pro-S-H of fluoroacetyl-CoA is activated in the enzymatic condensation. The hydroxycitrate of Garcinia isolated by Lewis (1&4) and his coworkers has been shown by Watson et al. (281) to be a potent inhibitor of rat liver ATP citrate lyase. The crystallographic studies by Glusker fluorocitrate was made biosynthetically from fluoroacetyl-CoA in a si attack on oxaloacetate, then the fluorine would be in a position that we have previously designated as carbon number 4. We would then use a parent number (pn) and call this compound (pn)-(4Ä)-4-fluorocitrate as opposed to use of the chemical number (25,3/S)-2-fluorocitrate. Even though we do not know the biosynthetic path of hydroxycitrate formation we could extend this system to it, and (2S,3S)2-fluorociträte. Even though we do not know the biosynthetic path of hydroxy­ citrate formation we could extend this system to it, and (2S,3S) -2-hydroxycitrate is equivalent to (pn)-(4£)-4-hydroxy ci träte.

THE CITRATE ENZYMES

241

1

COO'

2 Y/ 3

COO"

I

H—C—H

c

\

/

OH

OOC—C—OH

c'

\ / \xx>

4

H—C—OH

I .

c

coo"

5

coo

coo" "

O O C

S/TY°H

HO^O>^H CI^COO"

(B)

(/m)-(4S)-4-Hydroxycitrate

FIG. 7. (A) (p. 240). Representations of (pw)-(4.R)-4-fluorociträte, the biosynthetic product of fluoroacetyl-CoA and oxaloacetate with pig heart citrate synthase. (From Carrell et al. (32)). (B) (above). Representations of Garcinia hydroxycitrate according to Glusker et al. (75) and and Boll et al. (17) and Lewis (124).

et al (75), the NMR and infrared (IR) studies by Boll et al (17) have shown that the Garcinia hydroxycitrate is (pn)-(4S)-4-hydroxycitrate.* These two compounds are shown in the Fig. 7. It can be seen that at the 4 position (méthylène derived from acetate) the stereochem­ istry is opposite, and in spite of the change in nomenclature (due to the change in the R-S priorities with fluorine substitution) the 3 position of both has the same stereochemistry. Thus (pn)-(4Ä)-4-fluorocitrate, a substrate for citrate synthase, has its 4-H s (of citrate) proton while the (pn) - (4S) -4-hydroxycitrate, an inhibitor for ATP citrate lyase, has no 4-H s proton available. Since the stereochemistry of both enzymes are the same, then we can say that the 4-H s proton is necessary for the enzymatic reaction to occur. * See note, μ239·

242

PAUL A. SRERE

II. Citrate Lyase (Citritase) A . Distribution The least studied of the citrate enzymes is citrate lyase, the one catalyzing the simplest reaction. This enzyme has been detected only in bacteria even though an enzyme which cataylzes a similar reaction, isocitritase, has been found in yeast and molds. This enzyme has been isolated and studied from Klebsiella aerogenes (formerly Aerobacter aerogenes) by Dagley and Dawes (4I) and by SivaRaman {169), Esherichia coli by Wheat and Ajl (240) and Bowen and SivaRaman (21), and in Streptococcus faecalis by Gunsalus and his co-workers (73, 172). In these bacteria the enzyme is inducible. In E. coli, glucose is needed in addition to citrate for growth while in K. aerogenes growth occurs on citrate alone. Stern and his co-workers have shown that Na + is necessary for the formation of citrate lyase (143, 144)- Harvey and Collins (89) have shown, however, that in S. diacetilactis the enzyme is constitutive. They showed that strains of Leuconostoc and one strain of S. liquefaciens also contained the enzyme constitutively. O'Brien et al. (I4I) showed that Salmonella typhimurium grown aerobically on citrate contained citrate lyase (pro-SS). B. Physical Properties Purified citrate lyase has been obtained by SivaRaman from K. aerogenes (169). This enzyme is reported by Mahadik and SivaRaman (129) to have a molecular weight of 575,000 and to be composed of eight subunits of identical size. The s2o,w value for K. aerogenes citrate lyase obtained by Bowen and Rogers (19) was 16.2 S, and a value of 4 X 10~7 cm2 sec -1 was reported for D20,w, thus yielding a molecular weight of 318,000. They also used the Archibald technique of molecular weight determination and obtained a value of 314,000. We (168) have measured a sedimentation coefficient of 14.2 S for the enzyme from K. aerogenes. According to Mahadik and SivaRaman (129) the enzyme is dissociated to tetramere and dimers when dialyzed against EDTA at low ionic strength. In addition, pMB causes a dissociation of the enzyme. The enzyme contains about 72 SH groups per mole. C. The Reaction 1. EQUILIBRIUM

The equilibrium constant of the reaction as calculated by Burton (30) is about 1 M. A number of attempts have been made to determine the

THE CITRATE ENZYMES

243

equilibrium constant of the enzyme-catalyzed reaction. Smith, Stamer, and Gunsalus {172) using an enzyme from S. faecalis determined a value of 0.64 M (27°C), while Harvey and Collins (90) have reported a value of 0.064 M (30°C) for the equilibrium constant of the reaction catalyzed by an enzyme from S. diacetilactis. Tate and Datta (220) have discussed the complexity and difficulty in obtaining an accurate measurement of the equilibrium constant of the citrate lyase reaction. They pointed out that one must consider the fact that both citrate and oxaloacetate form stable chelates with Mg2+ as well as the fact that oxaloacetate exists as a tautomerie mixture of enol and keto forms, each of which forms chelates. In addition, buffers commonly used (Tris and phosphate) often interact with metal ions. With the en­ zyme from E. coli and K. aerogenes additional problems of enzyme in­ stability, inhibition by oxaloacetate, and enzyme inactivation during reaction are introduced. Tate and Datta's measurements (220) have been performed under a variety of conditions and they have obtained an ap­ parent equilibrium constant of about 0.3 M at 25°. 2. METAL REQUIREMENT

A number of divalent metals were reported by Dagley and Dawes (42) to be effective with the enzymes from E. coli and K. aerogenes. These include Mg2+, Mn2+, Fe2+, Co2+, Zn2+, and Ni2+. The ions Cu2+, Hg2+, Ca2+, Sr2+, and Ba2+ were inactive in the reaction, but Ca2+ has been shown to be a competitive inhibitor for Mg2+. Harvey and Collins (90) also showed that Ca2+ was a competitive inhibitor for Mg2+ with the enzyme from S. diacetilactis. They concluded in addition that the rate of reaction was dependent upon the concentration of 1:1 Mg-citrate complex. Ward and Srere (230) using the enzyme from S. diacetilactis showed by pulsed NMR techniques that Mn2+ formed a binary complex with the enzyme but could not show ternary complex formation with any of the other substrates. The Kais for the E-Mn complex was cal­ culated to be 0.4 X 10-4 M which is to be compared to a Km for Mn2+ of 0.9 X IO-4 M. In addition it was shown that Ca2+ competes for the Mn2+ binding site on the enzyme. They concluded that for this enzyme a metal-enzyme complex acted upon free citrate. Studies by Blair et al. (13) on the stability of the enzyme from K. aerogenes also indicated the presence of metal-binding sites on the en­ zyme. These workers found that the enzyme is reversibly inactivated at low Mg2+ concentrations. They found also that there was in addition an irreversible component of the inactivation. In agreement with other re­ sults with Ca2+, they found that this ion was also able to protect the enzyme from inactivation.

244

PAUL A. SRERE

3. REACTION-INACTIVATION

Wheat and Ajl (241) using a partially purified enzyme from E. coli observed that the rate of the reaction decreased to zero during the course of the reaction. They concluded that the loss in activity was the result of inhibition of the enzyme by OAA. Product inhibition of the enzyme by OAA had been observed and studied with the enzymes obtained from E. coli, K. eterogenee, and Streptococcus faecalis. It was further suggested that the enol form of OAA reported to be the product in the reaction {219) is the cause of this apparent inactivation (20, 42, 60, 220). In the course of their studies with the citrate lyase obtained from K. aerogenes, Singh and Srere (168) employed the MDH-coupled assay system as well as a direct spectrophotometric assay of OAA formation. The decrease in enzyme activity occurred whether or not OAA is allowed to accumulate in the system. If inhibition by OAA is the cause of this phenomenon, one would expect some difference in the rate of decrease of enzyme activity when OAA is allowed to accumulate, compared to the system where OAA is being continuously removed. When Mn2+, Mg2+, and Co2+ were used as the metal ions, the rate of inactivation was first order with values of about 1 min -1 . Zn2+ which functions well in the catalytic reaction does not cause a concomitant in­ activation of the enzyme. This is in agreement with the earlier reports of Gruber and Moellering (81, 138), who reported stabilization of the enzyme by Zn2+. The mechanism of reaction-inactivation is not under­ stood, although the oxaloacetate portion of the citrate molecule becomes attached to the enzyme during inactivation (168). The enzyme from S. diacetilactis behaves normally with Mg2+ and Mn 2+ in that it is not inactivated during the course of its reaction (90), nor is it inhibited by oxaloacetate as is the case with the citrate lyase from the E. coli, K. aerogenes, and S. faecalis. 4. PRESENT STATUS

No other substrate has been reported for the citrate lyase reaction, and Wheat and Ajl (241) showed its stereospecificity to be the same as that for citrate synthase. Very few studies are available concerning in­ hibition of the enzyme. Daron and Gunsalus (44) report that NaF in­ hibits the reaction, but this is probably due to a chelation of the Mg2+ by F". Inhibition by pCMB is reported by Mahadik and SivaRaman (129). Srere et al. (204) have shown that the enzyme from K. aerogenes is inhibited by hydroxylamine and that the hydroxylamine inactivation could be reversed by treatment of the enzyme with acetic anhydride.

245

THE CITRATE ENZYMES

Enzyme which is reaction-inactivated is also activated with acetic an­ hydride. The enzyme has been recently shown to contain stoichiometric amounts of phosphopantothenate (196a). It would seem that an acetyl group is necessary for the formation of an active site on the enzyme, possibly on a pantetheine SH. We have found that crude enzyme extracts can also be activated with acetic anhydride so that the possibility exists that an acetylation-deacetylation mechanism exists for the control of this enzyme activity. It does not, however, answer the question of how the cell handles the prob­ lem of reaction-inactivation. Buckel, Buschmeier, and Eggerer (29a) have recently shown that an acetyl group on the enzyme is necessary for the activity of citrate lyase. They have been able to label the enzyme with stereospecifically r e ­ labeled citrate and show that the removal of the label with NH 2 OH, SH compounds or oxaloacetate leads to inactive enzyme. They have proposed that the enzyme contains the acetyl group as a thioester and that the mechanism proceeds E-S ~ acetyl + citrate = E-S ~ citryl -f acetate E-S ~ citryl = E-S ~ acetyl + oxaloacetate Sum: Citrate = acetate + oxaloacetate

Thus these workers make the very important proposal that the citrate lyase performs at an "enzyme level" what occurs at a "substrate level" in the citrate synthase reaction. They feel that loss of the acetyl group accounts for both reaction-inactivation and oxaloacetate inhibition. Although it is probable that a metal enzyme is involved in the enzyme reaction it is not at all clear whether or not a metal-citrate complex is the substrate for the reaction. III. Citrate Synthase A. Distribution Citrate synthase has been found in every cell (except for certain E. coli mutants) examined for it. In eukaryotic cells it occurs chiefly in the mitochondria, but if the cells contain glyoxisomes it is also found in those organelles (23, 36, 37). In mitochondria it is located within the inner membrane-matrix fraction, and there is some evidence that it is loosely attached to the inner membrane (193). It is presumed to be syn­ thesized in the cytosol, as are other enzymes of the Krebs cycle (107, 108). Its metabolic role in mitochondria is as part of the Krebs cycle

246

PAUL A. SRERE

IÌH- · i i f a P a Ä ·.· ·'.·■·'·»,'WAS,

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FIG. 8. Crystals of citrate synthase. (A) Eschenchia coli. (B) Rat liver. (C) Azotobacter. (D) Rat kidney. All magnified about X 600.

. *

THE CITRATE ENZYMES

247

and as such the control of its activity has been postulated as a key regu­ latory step of the cycle. Glyoxysomal citrate synthase functions in the glyoxylate shunt in which 2 acetyl units are converted to succinate. It is not known whether in those cells containing both organelles the citrate synthase in both is the same protein. In aerobic prokaryotic cells citrate synthase undoubtedly functions also as part of the Krebs cycle. In bacteria which contain the glyoxylate shunt pathway, it must also serve as part of that metabolic process. There is no indication as to the intracellular location of citrate synthase in bacteria, or whether or not the identical enzyme serves in both path­ ways. In a citrate synthaseless mutant of E. coli, Ashworth et al. (4) showed that the structural gene for the enzyme mapped close to the galactokinase gene. In bacteria, especially anaerobes, a second important function of the Krebs cycle is in the synthesis of glutamic acid. Thus in anaerobes or facultative anaerobes which lack a-ketoglutarate dehydrogenase com­ plex, the remaining two arms of the Krebs cycle will function as biosynthetic pathways. Citrate synthase was first described in pigeon liver by Stern and Ochoa {207a) and the first pure preparation of the enzyme was the crystalline enzyme prepared from pig heart by Ochoa et al. {H5). Srere and Kosicki {199) developed a rapid procedure yielding far more pig heart citrate synthase. The enzyme is available commercially, and as a consequence much of the research has been on the pig heart enzyme. At present we have purified and crystallized the enzyme from E. coli {62, 167), Azotobacter {14), rat heart {139), rat liver {139), rat kidney {134), rat brain {134), pigeon breast muscle {149, 191), and moth flight muscle {191, 197). Figure 8 shows some of these crystals. Other workers have reported on the preparation of pure enzymes from E. coli {236), yeast {14S, 149), and beef heart {171). In addition studies have appeared on partially purified enzymes from beef liver {105), mango fruit {202), lemon mitochondria {15, 16), Clostridium acidi-urici {80), Garcinia leaves {47), cauliflower buds {163), citrus fruit {203), Bacillus subtilis {68), Rhodopseudomonas {57), Rhodo spirillum {57), and trout liver {95). There are studies on the enzyme in extracts from various aerobic and facultative anaerobic bacteria {239), from strictly anaerobic bac­ teria {78), higher plants {93), fungi {67, 110, 156), and several animal tissues {26, 105). There are apparent variations among these citrate synthases concern­ ing their size, structure, and function, and a more organized survey of the enzyme in a phylogenetic scale of cells is necessary before common characteristics can be recognized.

248

PAUL A. SRERE

B. Physical Properties There is wide variation of the size of synthases from various cell types thus far examined. Table I shows the molecular weights reported for different citrate synthases. The enzymes from animals have similar molecular weights, but the only consistency recognizable among them might be a monomer size of about 50,000. Weitzman and Dunmore (237) have classed citrate synthases as large or small on the basis of gel filtration experiments and claim that allosteric properties are seen only with the "large" enzymes. Reports of allosteric effects of indole acetate accompanied by size changes of the enzyme failed to give a size of the normal or affected enzyme (163). The best-studied citrate synthase, from pig heart, has the molecular properties as determined by us shown in Table II (166). Wu and Yang (250) have obtained similar results in their extensive studies of the en­ zyme. They determined a diffusion coefficient of 5.8 X 10~7 cm2/sec and an intrinsic viscosity of 3.95 ml/gm for the native enzyme as well as determining experimentally a specific volume of 0.73 ml/gm. Their cal­ culations of molecular weight of the native enzyme averaged 1.0 ± 0.05 X 105. The optical properties of the enzyme reported by them indi­ cated a higher helix content than do the results of Srere (186). The data of Wu and Yang were obtained at ten times or higher protein concentra­ tions than those of Srere. The results of these studies indicate that pig heart citrate synthase is a compact, symmetrical, globular molecule with TABLE I MOLECULAR W E I G H T S AND SUBUNITS OF VARIOUS CITRATE SYNTHASES

Source Pig heart Escherichia coli

R a t heart R a t tissue Pigeon breast Moth muscle Azotobacter Mango

M W X 105

Subunits

Method 0

Reference

1.0 0.96 2.1 2.8 2.3 2.5 1.0 1.0 1.0 1.4 3 5 0.65

2 2 4

SE SE SE GF EM GF SE SE SV SE SE GF GF

250 166 167 62 161 249 139 139 191 191 14

— 4 4 2 2

?

2 6 ? ?

H

202

a SE, sedimentation equilibrium; GF, gel filtration; E M , electron microscopy; SV, sedimentation velocity.

249

THE CITRATE ENZYMES TABLE II MOLECULAR P R O P E R T I E S OF P I G H E A R T CITRATE SYNTHASE*

S20

IN] Mw (sed. equil.) M (N) v [amino acid] 00

bo 0219 0208

E\%m (280 nm) α

Native

in 7 M GuHCl

6.2 X 10"13sec — 0.96 X 105 — 0.74 ml/gm — 4 deg cm2/dmole - 2 3 0 deg cmVdmole -11,400 deg cm2/dmole -11,800 deg cmVdmole 17.8

— 38.5 ml/gm 0.46 X 105 0.45 X 105 — — 405 deg cmVdmole 0 — — —

These data are from Singh et al. (166) and Srere {186).

a helical content between 30 and 60%. It is also apparent that it contains two subunits which according to all physical measurements are identical. The results of Wu and Yang (250) indicate that each subunit is asym­ metrical and they propose a side-by-side interaction between them to give a symmetrical protein. Antibodies against a crude pig heart prep­ aration was shown by Broder and Srere (25) to neutralize the activity of several other synthases. C. Chemical Properties of the Enzyme and Active Site The amino acid composition of the pig heart enzyme has been reported by Wu and Yang (250) and by Singh et al. (166) with fair agreement between the values. No phosphorus, lipid, or carbohydrate was detected by Singh et al. (166), and they could account for the total dry weight of the protein as amino acids and cations. The studies of Singh et al. (166) showed that tryptic digests of the pig heart enzyme yields 41 ninhydrin-positive spots. On the basis of the 32 total argininyl and 48 lysyl residues one expects 81 spots if a single polypeptide chain is involved. Therefore, these data are in agreement with the physical data and indicate that the enzyme is made up of two identical subunits. Preliminary amino acid analyses have been performed on the pigeon breast and moth flight muscle citrate synthases (194)· The percentage of the various amino acids in these two enzymes is similar to that of the pig heart enzyme. A peptide map of the pigeon breast enzyme is quite similar to that of the pig heart enzyme (Fig. 9) (61). Amino acid analyses of E. coli citrate synthase has been reported by Wright and

250

PAUL A. SKEBE

Θ| Θ

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PYRIDINE=Ac=HfO» 100.4=900 pH6.5 2 4 0 0 * 2 5 0 « A . 70HIN nBuOH'AoHjO»* 1=5 DESCENDING I8MA 74-75

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o to pig heart citrate synthase with a KO of 100 μΜ. Using this spin label these authors were able to show that the pig heart enzyme con­ tained 1 binding site/48,000 gm indicating 2 active sites per mole of enzyme. By titrating the spin-labeled complex with acetyl-CoA a dis­ sociation constant for this compound of 140 μΜ was obtained. This figure is 30 times higher than Km and since the kinetic pattern with this en­ zyme suggests that ΚΌ = Km it is probable that the Km represents a constant for the binary complex of oxaloacetate-enzyme but not for the free enzyme. The stereochemistry of the fluorocitrate made from fluoroacetyl-CoA has been examined by crystallographic methods (32). The data suggest that the isomer formed is (pn)-(4Ä)-4-fluorocitrate as illustrated earlier in Fig. 7A. Fluorooxaloacetate is the only compound aside from oxaloacetate that can condense with acetyl-CoA in the citrate synthase reaction (63). Srere has tried a number of keto acids (glyoxalate, pyruvate, monoethyl oxaloacetate, α-ketoglutarate, ketomalonate, α-ketobutyrate) as possible substrates, but they were inactive with the pig heart citrate synthase (186). Fluorooxaloacetate is competitive with both oxaloacetate (Kx = 125 μΜ) and acetyl-CoA (Ki = 1.8 m l ) according to Fanshier et al. (63). Its Km in the reaction is 140 μΜ, and F m a x with it as substrate is 0.1 that seen with oxaloacetate. It has been shown by Englard (58) and later by Annett and Kosicki (2) that ketooxaloacetate is the reactive species in the pig heart citrate synthase-catalyzed reaction. Englard followed deuterium incorporation from a D 2 0 medium into citrate to establish this point while Annett and Kosicki used a purified oxaloacetate enolase to establish the form of the substrate. I have investigated the binary complex of pig heart citrate synthase with oxaloacetate and found it to be quite stable (182, 186). The forma­ tion of this complex is accompanied by a shift in the ultraviolet spectrum

THE CITRATE ENZYMES

259

of the enzyme. The complex is stable in 4 M urea as judged by activity measurements, reaction of its SH groups with DTNB, ORD measure­ ments, and ultracentrifugai studies. The rate constant for the appearance of SH groups in 4 M urea could be easily measured and addition of oxaloacetate protected the enzyme from urea denaturation and SH group appearance. With this technique it was possible to calculate ΚΌ for oxaloacetate as 0.6 μΜ which is to be compared with its Km for this enzyme of 1.5 μΜ. A similar study using fluorooxaloacetate indicates a ΚΌ of 100>M which compares well with its Ki of 125 μΜ (68). Interac­ tions between other oxaloacetate analogs and the enzyme could be studied using the protection technique described above. a-Ketoglutarate, transaconitate, and D-isocitrate were good protectors, while aconitate, tricarballylate, oxalate, citrate, and L-malate were fair protectors and L-isocitrate, maleate, and D-malate were very poor protectors. This method also indicated that 1 mole of oxaloacetate or fluorooxalo­ acetate was bound per protecting (active) site. It is reasonable to assume a conformation change occurs in the enzyme and that this change brings an enzyme residue close to the pro-S H of acetyl-CoA aiding in proton abstraction. The facts that two other protectors, L-malate and a-ketoglutarate, induce an enolase activity in the enzyme and that an inversion gf stereochemistry of the protons on the acetyl group occurs make this mechanism very attractive. Kosicki and Srere (119) showed that substitution of deuteroacetyl-CoA for acetyl-CoA resulted in a kinetic isotope effect, fcH/fcD, of 1.4 at all pH values. The results of Klinman and Rose (109) using acetyl-CoA containing H, D, and T in the acetyl methyl group show an intramolecu­ lar isotope effect kK/kT of 4.4 which they have indicated to be equal to a fcH/fcD of 3.0. They find, however, that the enrichment of T in citrate does not agree with these values and postulate additional modes of racemization of the H isotopes on that carbon. In any event, these data, along with the exchange data of Eggerer (51), clearly imply that proton removal is a rate-limiting step in the overall reaction. This is consistent with the known chemistry of aldol condensations. Kosicki and Srere also showed (119) that if the citrate synthase reac­ tion is allowed to proceed in D 2 0, then ku/kD = 2.8 at pH 8.1. This ef­ fect is pH (pD) dependent varying from 3.8 at pH 7.5 to 0.8 at pH 10. These effects were also seen when the reverse reaction was measured in D 2 0 at various pH values. These data were interpreted as indicating that the active site contained at least two basic groups, one protonated and one dissociated in its active form. Melander (136) has reported similar behavior in D 2 0 in acid-catalyzed reactions. Citryl-CoA was a logical and early postulated intermediate in the

260

PAUL A. SRERE

citrate synthase reaction. Stern {205) reported that citryl-CoA was not cleaved by the enzyme and later Srere and Kosicki {200) reported that a preparation of citryl-CoA inhibited citrate synthase reaction. Eggerer and Remberger (54) were the first to show that citryl-CoA could be hydrolyzed by the enzyme, a result that was later confirmed by Srere (181). The rate of hydrolysis was slower than the overall reaction, but citryl-CoA was a chemically synthesized material consisting of a mixture of R- and S-citryl-CoA, and it was possible that the i2-citryl-CoA was an inhibitor of the reaction. Srere (181) was able to demonstrate such inhibition with the synthetic mixture. Eggerer was able to show that the citryl-CoA could be aldol-cleaved to yield acetyl-CoA and oxaloacetate. Since no exchange of 14C-labeled acetyl-CoA into a pool of citryl-CoA could be detected, Eggerer concluded that the citryl-CoA was tightly bound to the enzyme during the reaction. Eggerer et al. (56) also showed that S-malyl-CoA could be hydrolyzed by pig heart citrate synthase, but Ä-malyl-CoA was not hydrolyzed. Since the reaction is a reversible one and the direct formation of citrylCoA from citrate and CoASH is not a plausible first step, a number of other postulates have been put forth. Cornforth (38) and later Arigoni and Eschenmoser (3) postulated β-lactone formation as an intermediate step while Eggerer (54) has considered the possibility that an anhydride is an intermediate in the reaction. Experiments using H 2 18 0 designed to test these postulates have been performed by Suelter and Arrington (212) Wunderwald and Eggerer {252). Suelter and Arrington studied the disposition of 18 0 from H 2 18 0 when citrate synthase was allowed to cata­ lyze the condensation in this medium. They found that 1 atom of 18 0 was incorporated into citrate in the carboxyl group derived from acetylCoA. Wunderwald and Eggerer found that the hydrolysis of malyl-CoA by citrate synthase (as well as malate synthase) in the presence of H 2 18 0 also found one atom of 18 0 in the carboxyl group of malate which had originated from acetyl-CoA. These data do not decide between or sup­ port either the lactone or anhydride theory although they do allow cer­ tain versions of them to be possible. The results with H 2 18 0 seem to rule against Schiff base formation between the enzyme and the two substrates. We (194) have shown with citrate synthase that the enzyme can be in­ cubated with NaBH 4 alone or in the presence of either or both of its substrates, without loss of activity. Wunderwald and Eggerer (252) have reported similar results with malate synthase. Buckel and Eggerer (29) have studied the chemical hydrolysis of citryl-CoA. They have concluded that an intramolecular nucleophilic attack by the carboxylate anion yields CoA and citryl anhydride, the latter of which is rapidly hydrolyzed. They postulated that neighboring

261

THE CITRATE ENZYMES HC:B

HB. I

X!

H H \c /

HCB

CSCoA

 A

^ \ _^ C^ c o °

-

°

"OOCT

\