The Enzymes, Vol VIII: Group Transfer, Part A (Nucleotidyl Transfer, Nucleosidyl Transfer, Acyl Transfer, Phosphoryl Transfer), 3rd Edition

The Enzymes VOLUME V I I I GROUP TRAXSFER Part A NUCLEO T I D Y L TRANSFER NUCLEOSIDYL TRANSFER ACYL TRANSFER PHOSPHOR...

Author: Paul D. Boyer

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The Enzymes VOLUME V I I I

GROUP TRAXSFER Part A NUCLEO T I D Y L TRANSFER NUCLEOSIDYL TRANSFER ACYL TRANSFER PHOSPHORYL TRANSFER Third Edition

CONTRIBUTORS R. P. AGARWAL

HENRY PAULUS

S. J. BENKOVIC

JACK PR.EISS

DAVID P. BLOXHAM R . GAURTH HANSEN

K. J. SCHRAY R. K. SCOPES E. R. STADTMAN

F. J. KAYNE EDWIN G. KREBS HENRY A. LARDY

JEREMY W. THORNER PAOLO TRUFFA-BACHI

J. F. MORRISON

RICHARD L. TURNQUIST

S. HARVEY MUDD

P. ROY VAGELOS DONALD A. WALSH

I,. NODA R. E. PARKS, JR.

D. C. WATTS

ADVISORY BOARD ARTHUR KORNBERG

FRITZ LIPMANN

HENRY LARDY

EARL STADTMAN HERBERT TABOR

THE ENZYMES Edited by PAUL D. BOYER Molecular Biology Institute and Department of Chemistry University of California Los Angeles, California

Volume VIII

GROUP TRANSFER Part A NUCLEOTIDYL TRANSFER NUCLEOSIDYL TRANSFER ACYL TRANSFER PHOSPHORYL TRANSFER

THIRD EDITION

A C A D E M I C P R E S S New York and London A Sitbsirlinry of Harcourl Brace Jovanovich, Publishers

1973

COPYRIGHT 0 1973, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED. NO PART OF THIS PUBLICATION MAY BE REPRODUCED OR TRANSMITTED IN ANY FORM OR BY ANY MEANS, ELECTRONIC OR MECHANICAL, INCLUDING PHOTOCOPY, RECORDING, OR ANY INFORMATION STORAQE AND RETRIEVAL SYSTEM, WITHOUT PERMISSION IN WRITING FROM THE PUBLISHER.

ACADEMIC PRESS, INC. 111 Fifth Avenue, New

York,New York 10003

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

Library of Congress Cataloging in Publication Data Main entry under title. The Enzymes. Includes bibliographical references. CONTENTS: v. 2. Kinetics and mechanism-v. 3. Hydrolysis: peptide bonds.-v. 4. Hydrolysis: other C-N bonds, phosphate esters.- [etc.] 1. Enzymes. I. Boyer, Paul D., ed. [DNLM: 1. Enzymes. QU 135 B791el QP601.ES23 574.1’925 75 - 117 107 ISBN 0-12-122708-1 (V.8)

PRINTED IN THE UNITED STATES OF AMERICA

Contents List of Coiitributors Preface

.

.

.

. . . ,

Contents of Other Volumes

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. .

,

ix xi xii

1. Adenylyl Transfer Reactions

E. R. STADTMAN

.

.

. . . . . . . . . . . . .

I. Introduction . . . . . . . 11. Carboxyl Group Activation . . . . . . . 111. Biosynthesis of Phosphodiester Derivatives of Adenosine IV. Synthesis of Adenosine Diphosphate Derivatives . . V. Sulfate Activation . . . . . . . . . VI. Synthesis of Imidol Adenylate Derivatives . . . VII. Adenylylation of Functional Groups of Proteins . .

. . .

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.

2 6 20 30

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51 55 62

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73 75 77 117

35 37 40

2. Uridine Diphosphoryl Glucose Pyrophosphorylase

RICHARD L. TURNQUIST AND R. GAURTH HANSEN I. Introduction . . . 11. Metabolic Function . 111. Propehes . . .

. . . . . . . . .

. . . . . .

. .

.

.

. . . . . . . . . . . . ,

3. Adenosine Diphorphoryl Glucose Pyrophosphorylase

JACK PREISS I. Introduction . . . . . . . . . . . . 11. Classification of ADPglucose Pyrophosphorylases . . . 111. Kinetic Properties of the ADPglucose Pyrophosphorylases IV. Physical Properties of the ADPglucose Pyrophosphorylases V

. . .

vi

CONTENTS

4

.

The Adenosyltransferases

S. HARVEY MUDD I. Introduction . . . . . I1. Methionine Adenosyltransferase I11. B.?. Adenosyltransferasc . . IV . Conclusion . . . . .

5

.

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . .

121 123 144 152

Acyl Group Transfer (Acyl Carrier Protein)

P. ROYVAGELOS I . Introduction . . . . . I1. Acyl Carrier Protein . . . I11. Malonyl CoA-ACP Transacylasc IV . Acetyl CoA-ACP Transacylase V. P-Ketoacyl ACP Synthetase . 6

.

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

155 156 176 185 188

Chemical Basis of Biological Phosphoryl Transfer

S. J . BENKOVIC AND K . J . SCHRAY I . Introduction . . . . . . . . . I1. Hydrolysis of Acyclic Phosphate Esters . . I11. Nucleophilic Reactions a t Acyclic Phosphorus IV . Pentacovalency and Pseudorotation . . . V . Catalysis of Phosphoryl Transfer or Ligand Loss VI . Enzymic Catalytic Mechanisms . . . .

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7

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201 202 208 214 219 232

Phosphofructokinase

DAVID P . BLOXHAM AND HENRY A . LARDY I. Introduction . . . . . . . . . I1. Purification . . . . . . . . . I11. Assay of Phosphofructokinase Activity . . IV . Catalytic Properties . . . . . . . V. Structural Properties . . . . . . VI . Regulatory Properties of P F K . . . . VII . Role of Specific Groups in Enzymic Activity . VIII . The Role of P F K in the Control of Glycolysis 8

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.

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240 241 243 244 253 261 269 274

Adenylate Kinase

L . NODA I . Biological Aspects . I1. Molecular Properties I11 Catalytic Properties .

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. . . . . . . . . . . .

279

288 297

vii

CONTENTS

9

.

Nucleoside Diphosphokinases

R . E . PARKS. JR., AND R . P. AGARWAL I . Introduct.ion

.

.

I1. Molecular Properties

.

I11. Catalytic Properties . IV . Functions in the Cell

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

307 313 320 331

10. 3-Phosphoglycerate Kinase

R . K . SCOPES I . Introduction . . . . . . . . . I1. Biological Behavior of Phosphoglycerate Kinase I11. Isolation and Molecular Properties . . . IV. Reaction Kinetics . . . . . . . V. Conclusion . . . . . : . . .

.

11

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.

.

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335 336 340 346 351

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

353 355

. . . . . .

. . . . . .

Pyruvate Kinase

F. J . KAYNE I . Introduction . . . I1. Molecular Properties I11. Catalytic Properties .

.

12

364

Creatine Kinase (Adenosine 5’-TriphosphateCreatine Phosphotransferasel

D . C. WATTS I . Introduction . . . . . . . . . . I1. Structure . . . . . . . . . . I11. Purification. Assay. and Enzyme Stability . . I V . Substrate Specificity . . . . . . . V. The Activating Metal Ion . . . . . . VI . Enzyme Kinetics . . . . . . . . VII . Chemical Investigations of the Enzyme Mechanism 13

.

. . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . . .

384 386 395 403 409 412 431

. . . . . . . . . . . . . . . . . . . . . . . . . . .

457 464 466

.

Arginine Kinase and Other Invertebrate Guanidino Kinases

J . F. MORRISON I . Introduction . . . . . . I1. Determination of Enzymic Activity I11. Molecular Properties . . .

viii

CONTENTS

IV . Catalytic Properties . V . Reaction Mechanism VI . Equilibrium . . .

.

14

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

471 482 485

Glycerol and Glycerate Kinases

JEREMY W . THORNER AND HENRY PAULUS I. Introduction . . I1. Glycerol Kinases I11. n-Glycerate Kinascs

15

.

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

487 485

501

Microbial Aspartokinases

PAOLO TRUFFA-BACHI I. Introduction . . . . . . . . . . . . I1. Escherichia coli Aspartokinascs . . . . . . . I11. Other Coliform Bacteria . . . . . . . . . IV . Aspartokinases Regulated by Concerted Feedback Inhibition V. Ithodopseiidomoitas spheroides . . . . . . . VI . Saccharomyces cerevisiae . . . . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . .

. . . .

. . . .

555 557 566

Author Index

. . . . . . . . . . . . . . . .

583

Subject Index

. . . . . . . . . . . . . . . .

617

509 513 544 544

552 553

. Protein Kinases

16

DONAL A . WALSHA N D EDWIN G . KREBS I . Introduction . . . . . . . . I1. Substrate-Specific Protein Kinascs . . 111. Cyclic Nucleotide-Regulated Protein Kinases IV . Nonclassified Protein Kinases . . .

. . . .

. . . .

. . . .

. . . .

575

List

of Contributors

Numbers in parentheses indicate the pages on which the authors’ contributions begin.

R. P. AGARWAL (307), Division of Biological and Medical Sciences, Brown University, Providence, Rhode Island

S. J. BENKOVIC (201), Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania DAVID Y . BLOXHAM (239), Institute for Enzyme Research, University of Wisconsin, Madison, Wisconsin R. GAURTH HANSEN (51), Department of Chemistry and Biochemistry, Utah State University, Logan, Utah F. J. KAYNE (353), Johnson Research Foundation, Department of Biophysics and Physical Biochemistry, The School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania EDWIN G. KREBS ( 5 5 5 ) , Department of Biological Chemistry, School of Medicine, University of California, Davis, California HENRY A. LARDY (239), Institute for Enzyme Research, University of Wisconsin, Madison, Wisconsin

J. F. MORRISON (457), Department of Biochemistry, The John Curtin School of Medical Research, The Australian National University, Canberra, Australia S. HARVEY AlUDD (121), Laboratory of General and Comparative Biochemistry, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland L. NODA (279), Department of Biochemistry, Dartmouth Medical School, Hanover, New Hampshire ix

LIST OF CONTRIBUTORS

X

R. E. PARKS, JR. (307), Division of Biological and Medical Sciences, Brown University, Providence, Rhode Island HENRY PAULUS (487), Department of Biological Chemistry, Harvard University Medical School, Boston, Massachusetts JACK PREISS (73), Department of Biochemistry and Biophysics, University of California, Davis, California K. J. SCHRAY (201), Department of Chemistry, Lehigh University, Bethlehem, Pennsylvania R. K. SCOPES (335), La Trobe University, Bundoora, Victoria, Australia E. R. STADTMAN (l), Laboratory of Biochemistry, National Heart and Lung Institute, National Institutes of Health, Bethesda, Maryland .JEREMY W. THORNER" (487), Department of Biological Chemistry, Harvard University Medical School, Boston, Massachusetts PAOLO TRUFFA-BACHI (509), Service de Biochimie Cellulaire, Institut Pasteur, Paris, France RICHARD L. TURNQUIST (51), Department of Biochemistry, Utah State University, Logan, Utah P. ROY VAGELOS (155),Department of Biological Chemistry, Washington University School of Medicine, St. Louis, Missouri DONAL A. WALSH (555), Department of Biological Chemistry, School of Medicine, University of California, Davis, California

D. C. WATTS (383), Biochemistry and Chemistry Department, Guy's Hospital Medical School, London, England

* Present

address : Department of Biochemistry, Stanford University School of Medicine, Stanford, California.

Preface Volumes VIII and I X of this treatise deal with the important and versatile enzymes that catalyze the transfer of a variety of chemical groups. The most extensively studied are enzymes catalyzing phosphoryl group transfer from ATP to various acceptors, known by their trivial name as kinases; thirteen separate chapters on kinases appear in the two volumes. Volume VIII also includes a general chapter dealing with the chemical basis of phosphoryl transfer. Intramolecular phosphoryl transfer was covered earlier in Volume VI with the isomerases; and phosphoryl transfer by phosphatases was covered in Volume IV with the hydrolases. Another class of transferases of prime metabolic importance are those catalyzing transfer of one carbon groups. These are covered in Volume IX. Prominent consideration is given the catalytic versatility of folic acid. Aspartate transcarbamylase also falls within this group because of the elegant researches on its separate catalytic and regulatory subunits that introduced new principles into enzymology. One carbon group transfer also includes some molecular information about methylation of DNA; rapid growth is anticipated in this area. As representative of the widespread occurrence of transfer of glycosyl moieties, some key enzymes of carbohydrate metabolism and biosynthesis appear in these volumes. Other miscellaneous group transfer reactions complete the coverage. As with previous volumes, the principal criterion for inclusion as a separate chapter is considerable information a t the molecular level about either the enzyme or the process catalyzed. Response of the best-qualified authors in the field continues to be gratifying. For nearly all the chapters, the author is the first choice of the Advisory Board and the Editor. It is to these authors that the reader is indebted for the excellent coverage. Also it is a pleasure to extend appreciation to the members of the Advisory Board for their invaluable assistance in planning the volumes. The continued fine professional work by the staff of Academic Press is clearly evident in the product produced.

PAUL D. B O Y ~ R xi

Contents of Other Volumes Volume I: Structure and Control

X-Ray Crystallography and Enzyme Structure David Eisenberg Chemical Modification by Active-Site-Directed Reagents Elliott Shaw Chemical Modification as a Probe of Structure and Function Louis A . Cohen Multienzyme Complexes Lester J . Reed and David J . COX Genetic Probes of Enzyme Structure Milton J . Schlesinger Evolution of Enzymes Emil L . Smith The Molecular Basis for Enzyme Regulation D . E . Koshland, JT. Mechanisms of Enzyme Regulation in Metabolism E. R . Stadtman Enzymes as Control Elements in Metabolic Regulation Daniel E . Atlcinson Author lndex-Subject

Index

Volume II: Kinetics and ,Mechanism

Steady State Kinetics W . W . Cleland Rapid Reactions and Transient States Gordon B . Hammes and Paul R . Schimmel xii

CONTENTS OF OTHER VOLUMES

Stereospecificity of Enzymic Reactions G . PopjQk Proximity Effects and Enzyme Catalysis Thomas C . Bruice Enzymology of Proton Abstraction and Transfer Reactions Irwin A . Rose Kinetic Isotope Effects in Enzymic Reactions J. H . Richards Schiff Base Intermediates in Enzyme Catalysis Esmond E . Snell and Samuel J . Di Mari Some Physical Probes of Enzyme Structure in Solution Serge N . Timasheff Metals in Enzyme Catalysis Albert S. Mildvan Author Index-Subject

Index

Volume 111: Hydrolysis: Peptide Bonds

Carboxypeptidase A Jean A . Hartsuck and William N . Lipscomb Carboxypeptidase B J. E . Folk Leucine Aminopeptidase and Other N-Terminal Exopeptidases Robert J . DeLange and Emil L . Smith Pepsin Joseph S . Fruton Chymotrypsinogen : X-Ray Structure J . Kraut The Structure of Chymotrypsin D . M . Blow Chymotrypsin-Chemical George P . Hess

Properties and Catalysis

xiii

xiv

CONTENTS OF OTHER VOLUMES

Trypsin B . Keil Thrombin and Prothrombin Staffan Magnusson Pancreatic Elastase B. S. Hartley and D. M . Shotton Protein Proteinase Inhibitors-Molecular Aspects Michael Laskowski, Jr., and Robert W . Sealock Cathepsins and Kinin-Forming and -Destroying Enzymes Lowell M . Greenbaum Papain, X-Ray Structure J . Drenth, J . N . Jansonius, R. Koekoek, and B. G. Wolthers Papain and Other Plant Sulfhydryl Proteolytic Eneymes A . N . Glazer and Emil L. Smith Subtilisin: X-Ray Structure J . Kraut Subtilisins : Primary Structure, Chemical and Physical Properties Francis S . Markland, Jr., and Emil L. Smith Streptococcal Proteinase Teh-Yung Liu and S. 1).Elliott The Collagenases Sam Seijter and Elvin Harper Clostripain William M . Mitchell and William F. Harrington Other Bacterial, Mold, and Yeast Proteases Hiroshi Matsubara and Joseph Feder Author Index-Subj ect Index

CONTENTS OF OTHER VOLUMES

xv

Volume IV: Hydrolysis: Other C-N Bonds, Phosphate Esters

Ureases F . J . Reithel Penicillinase and Other p-Lactamases Nathan Citri Purine, Purine Nucleoside, Purine Nucleotide Aminohydrolases C. L. Zielke and C. H . Suelter Glutaminase and y-Glutamyltransferases Standish C . Hartman L-Asparaginase

John C . Wriston, Jr. Enzymology of Pyrrolidone Carboxylic Acid Marian Orlowski and Alton Meister Staphylococcal Nuclease X-Ray Structure F . Albert Cotton and Edward E . Haxen, Jr. Staphylococcal Nuclease, Chemical Properties and Catalysis Chrisfian B . Anfinsen, Pedro Cuatrecasas, and Hiroshi Taniuchi Microbial Ribonucleases with Special Reference to RNases T,, T,, N,, and U, Tsuneko Uchida and Fuji0 Egami Bacterial Deoxyribonucleases I . R. Lehman Spleen Acid Deoxyribonuclease Giorgio Bernardi Deoxyribonuclease I M . Laskowski, Sr. Venom Exonuclease M . Laskowski, Sr. Spleen Acid Exonuclease Alberto Bernardi and Giorgio Bernardi

xvi

CONTENTS OF OTHER VOLUMES

Nucleotide Phosphomonoesterases George I. Drummond and Masanobu Yamamoto Nucleoside Cyclic Phosphate Diesterases George I. Drummond and Masanobu Yamarnoto E. coli Alkaline Phosphatase Ted W . Reid and Irwin B. Wilson Mammalian Alkaline Phosphatases H. N. F m l e y Acid Phosphatases Vincent P. Hollander Inorganic Pyrophosphatase of Escherichia coli John Josse and Simon C. K. Wong Yeast and Other Inorganic Pyrophosphatases Larry G. Butler Glucose-6-Phosphatase, Hydrolytic and Synthetic Activities Robert C. Nordlie Fructose-l,6-Diphosphatases S. Pontremoli and B. L. Horecker Bovine Pancreatic Ribonuclease Fredem’c M . Richards and Harold W . Wyckoff Author Index-Subject Index

Volume V: Hydrolysis (Sulfate Esters, Carboxyl Esters, Glycosides), Hydration

The Hydrolysis of Sulfate Esters A. B . Roy Arylsulfatases R. G. Nicholls and A. B. Roy Carboxylic Ester Hydrolases Klaus Krisch Phospholipases Donald J . Hanahan

CONTENTS OF OTHER VOLUMES

Acetylcholinesterase Harry C. Froede and Irwin B. Wilson Plant and Animal Amylases John A. Thoma, Joseph E. Spradlin, and Stephen Dygert Glycogen and Starch Debranching Enzymes E . Y . C. Lee and W . J . Whelan Bacterial and Mold Amylases Toshio Takagi, Hirolco Toda, and Toshizo Isenzura Cellulases D. R. Whitaker Yeast and Neurospora Invertases J . Oliver Lampen Hyaluronidases Karl Meyer Neuraminidases Alfred Gottschallc and A. S. Rhargava Phage Lysozyme and Other Lytic Enzymes Akira Tsugita Aconitase Jenny Pickworth Glusker p-Hydroxydecanoyl Thioester Dehydrase Konrad Bloch Dehydration in Nucleotide-Linked Deoxysugar Synthesis L. Glaser and H . Zarkowsky Dehydrations Requiring Vitamin B,, Coenzyme Robert H . Abeles Enolase Finn Wold Fumarase and Crotonase Robert L. Hill and John W . Teipel

xvii

xviii

CONTENTS O F OTHER VOLUMES

6-Phosphogluconic and Related Dehydrases W . A. Wood Carbonic Anhydrase S. Lindslcog, L. E . Henderson, K . K . Kannan, A . Liljas, P. 0. Nyman, and B . Strandberg Author Index-Sub j ect Index Volume VI: Carboxylation and Decarboxylation 1Nonoxidative) , lromeriration

Pyruvate Carboxylase Michael C. Scrutton and Murray R. Young Acyl-CoA Carboxylases Alfred W . Alberts and P. Roy Vagelos Transcarboxylase Harland G . Wood Formation of Oxalacetate by COz Fixation on Phosphoenolpyruvate Merton F . Utter and Harold M . Kolenbrander

Ribulose-1,5-Diphosphate Carboxylase Marvin I. Siegel, Marcia Wishnick, and M . Daniel Lane Ferredoxin-Linked Carboxylation Reactions Bob B. Buchanun Amino Acid Decarboxylases Elizabeth A . Boelcer and Esmond E . Snell Acetoacetate Decarboxylase Irwin Fridovich Aldose-Ketose Isomerases Ernst A . N o l t m n n Epimerases Luis Glaser Cis-Trans Isomeriaation Stanley Seltzer Phosphomutases W . J . Ray, Jr., and E. J . Peck, Jr.

CONTENTS OF OTHER VOLUMES

xix

Amino Acid Racemases and Epimerases Elijah Adams Coenzyme BIz-Dependent Mutases Causing Carbon Chain Rearrangements H . A. Barker

B,, Coenzyme-Dependent Amino Group Migrations Thressa C . Stadtman IsopentenylpyrophosphateIsomerase P . W . Holloway Isomerization in the Visual Cycle Joram Heller A5-3-Ketosteroid Isomerase Paul Talulay and Ann M . Benson Author Index-Subject

Index

Volume VII: Elimination and Addition, Aldol Cleavage and Condensation, Other C-C Cleavage, Phosphorolysis, Hydrolysis (Fats, Glycosides)

Tryptophan Synthetase Charles Yanofsky and Irving P. Crawford Pyridoxal-Linked Elimination and Replacement Reactions Leodis Davis and David E . Metzler The Enzymic Elimination of Ammonia Kenneth R. Hanson and Evelyn A . Havir Argininosuccinases and Adenylosuccinases Sarah Ratner Epoxidases William B. Jakoby and Thorsten A . Fjellstedt Aldolases B. L. Horecker, Orestes Tsolas, and C. Y . Lai Transaldolase Orestes Tsolas and B . L. Horecker

xx

CONTENTS OF OTHER VOLUMES

2-Keto-3-deoxy-6-phosphogl~conic and Related Aldolases W . A. Wood Other Deoxy Sugar Aldolases David Sidney Feingold and Patricia A n n Hoffee 6-Aminolevulinic Acid Dehydratase David Shernin 6-Aminolevulinic Acid Synthetase Peter M . Jordan and David Shemin Citrate Cleavage and Related Enzymes Leonard B. Spector Thiolase Ulrich Gehring and Feodor Lynen Acyl-CoA Ligases Malcolm J . P . Higgins, Jack A. Kornblatt, and Harry Rudney a-Glucan Phosphorylases-Chemical and Physical Basis of Catalysis and Regulation Donald J. Graves and Jerry H . Wang Purine Nucleoside Phosphorylase R. E . Parks, Jr., and R. P . Agarwal Disaccharide Phosphorylases John J. Mieyal and Robert H . Abeles Polynucleotide Phosphorylase T . Godefroy-Colbum and M . Grunberg-Manago The Lipases P . Desnuelle p-Galactosidase Kurt Wallenfels and Rudolf Weil Vertebrate Lysozymes Taiji Imoto, L. N . Johmon, A . C . T . North, D . C . Phillips, and J . A. Rupley Author Index-Subject

Index

Adenylyl Transfer Reactions E. R . STADTMAN I . Introduction . . . . . . . . . . . . . I1. Carboxyl Group Activation . . . . . . . . . . A . Activation of Fatty Acids and Amino Acids and General Considerations . . . . . . . . . . . B . Nucleic Acid-Independent Peptide Synthesis . . . . C Acylation of the NE-Lysyl Residues of Enzymes . . . D . Adenylylation of Luciferin and Dehydroluciferin . . . I11. Biosynthesis of Phosphodiester Derivatives of Adenosine . . . A . RNA Synthesis . . . . . . . . . . . B. Adenyl Cyclase . . . . . . . . . . . C . Adenylylation of Amino Glycoside Antibiotics . . . IV . Synthesis of Adenosine Diphosphate Derivatives . . . . . A . General Features . . . . . . . . . . B. Synthesis of ADPglucose . . . . . . . . C . Adenine-Myonic Acid Dinucleotide and Adenylyl . . . . . . . . . Diphosphoglycerate V . Sulfate Activation . . . . . . . . . . . . VI . Synthesis of Imidol Adenylate Derivatives . . . . . . A Argininosuccinate Synthesis . . . . . . . . B . Synthesis of GMP . . . . . . . . . . . VII . Adenylylation of Functional Groups of Proteins . . . . . A . Adenylylation and Deadenylylation of E . Cali Glutamine Synthetase . . . . . . . . . . . B . Adenylylation of the LysineSensitive Aspartokinase of E coli . . . . . . . . . . . . C . Adenylyl Transfer Functions of DNA Ligase . . . . D . Covalent Modification of RNA Polymerase . . . .

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1

2 6 6 11 17 19 20 20 26 27 30 30 32 33 35 37 38 39 40 40 44 45 48

2

E. R. STADTMAN

1. Introduction

The high group transfer potential of ATP makes it a substance of singular importance in energy metabolism. The standard free energy of hydrolysis of ATP to produce AMP and inorganic pyrophosphate, Eq. (1), or to produce ADP and orthophosphate, Eq. ( 2 ) , is a measure of its adenylyl group transfer potential, or phosphoryl group transfer potential, respectively (1).

+ HzO + AMP + PPi ATP + HzO ADP + Pi ATP

(1) (2) ATP and the products of both of these reactions exist in solution as equilibrium mixtures of several polyanionic species, each of which can form a complex with divalent cations. Therefore, the observed equilibrium constant, Koba,of Eqs. (1) and (2) -+

are functions of the acid dissociation constants of each ionic species present and also of the stability constants of their complexes with divalent cations (9-7).The observed standard free energy of hydrolysis, AGEbs, for these reactions is therefore dependent upon the hydrogen ion and divalent cation concentrations. Data summarized in Table I show the effect of pH and pMg2+ on the G& of Eqs. (1) and (2). These data were obtained by interpolation from the contour diagrams of Alberty ( 4 ) which relate pH, pMg2' and AGzbs. At pH 6.0, the AG:b, of both reactions are about the same, except at the highest Mg2' concentration (0.1M) which favors Eq. ( 1 ) . However, a t higher pH values, the AGzbBfor Eq. (1) is significantly greater than that for Eq. (2) and the disparity increases with increasing Mg2' concentration. It is evident from these data that under physiological conditions, ATP has a higher adenylyl group transfer potential than phosphoryl group transfer potential; i.e., it is a better adenylylating agent than a phos1. F. Lipmann, Advan. Enzymol. 1, 99 (1941). 2. A. Schuegraf, S. Ratner, and R. C. Warner, JBC 235, 3597 (1960). 3. H. G. Wood, J. J. Davis, and H. Lochmuller, JBC 241, 5692 (1966). 4. R. A. Alberty, JBC 244, 3290 (1969). 5. R. A. Alberty, JBC 243, 1337 (1968). 6. R. C. Phillips, P. George, and R. J. Rutman, JBC !244, 3330 (1969). 7. M . R. Atkinson and R. K. Morton, Comp. Biochem. 2, 1 (1960).

1.

3

ADENYLYL TRANSFER REACTIONS

TABLE I EFFECT OF pH AND Mga+ CONCENTRATION ON THE FREE ENERQY OF HYDROLYSIS OF THE B,y- AND +~-PYROPHOSPHATE BONDSOF ATP

Mgl+ concn.

ATP

+ Hz0

-+

ADP

+ Pi

ATP

+ Hz0

-+

AMP

+ PPi

(M)

pH 6.0

p H 7.0

pH 8.0

pH 6.0

pH 7.0

pH 8.0

10-1 10-2 10-8 10-7

8.3 8.2 8.7 8.9

9.3 8.5 8.8 9.6

10.5 9.6 9.6 10.6

9.8 8.4 8.4 8.5

12.0 10.0 9.7 9.7

14.0 12.7 12.0 11.3

The data were derived by interpolation from contour diagrams relating AG;,,, pH and pMg, published by Alberty (4).

phorylating agent (2-7). Accordingly, when ATP is used as a source of energy for highly endothermic biological processes, mechanisms involving the adenylyl group transfer function are more suitable than those involving the phosphoryl group transfer function. This is clearly illustrated by two well-established mechanisms for the formation of acetyl-CoA from acetate and CoA, Eq. (3), which is strongly endergonic, AGL. = 8.5 kcal mole-'. Acetate

+ CoA -+ acetyl-CoA + H20

(3) In mechanism A, the phosphoryl group of ATP is transferred to acetate forming an energy-rich acetyl-P derivative, Eq. (3a). The acetyl moiety of this intermediate is then transferred to CoA to generate the energy-rich acetyl-CoA derivative, Eq. (4). In the overall reaction ATP is cleaved to ADP and Pi, and the energy of this cleavage ( z -8.9 kcal mole-') is utilized for the synthesis of acetyl-CoA, Eq. ( 5 ) . AGk

Mechanism A ATP acet.ate + acetyl-P ADP Acetyl-P CoA + acetyl-CoA P,

+

Sum: ATP

+

+

(kcal mole-') +3.1 -3.0

+

+ acetate + CoA + acetyl-CoA + ADP + Pi

Mechanism B ATP acetate + acetyl-AMP PPi Acetyl-AMP CoA -+ acetyl-CoA AMP

+

Sum: ATP

+

+

+ acetate + CoA

---*

+

acetyl-CoA

+ PPi f AMP

(34 (4)

+o. 1

(5)

+3.6 -5.7

(6)

-2.1

(8)

(7)

Mechanism B is similar except that the adenylyl group rather than the phosphoryl group of ATP is transferred in the first step, Eq. (6),

4

E. R. STNITMAN

to yield acetyl-AMP which serves as the acetyl donor in the formation of acetyl-CoA, Eq. (7). It is noteworthy that a t pH 7.3 and pMg 2.0, the AG:bs for Eqs. (3a) and (6) are +3.1 and +3.6 kcal mole-', respectively ( 7 a ) . Thus, in the activation of acetate, the adenylyl transfer reaction is slightly less favorable than the phosphoryl transfer reaction. However, the thermodynamic advantage of mechanism B is that acetylAMP has a much greater group reaction potential, -14.6 kcal mole-' ( 7 a ) ,than does acetyl-P, -12.0 kcal mole-' ( 7 a ) . Synthesis of acetylCoA by mechanism B is therefore considerably more exergonic than by mechanism A. In addition, the thermodynamic advantage of mechanism B can be augmented through removal of the product, PPi, by the action of pyrophosphatase, Eq. (9), which is abundantly present in most, if not, all cells. PPi

+ Hz0

-+

2 Pi

(9)

At pH 7.3 and pMg 2.0, the AG:,,, for Eq. (9) is -6.5 kcal mole-' ( 4 ); therefore, the overall coupled process, Eq. ( l o ) , is highly exergonic (AG& = -8.6 kcal mole-'). ATP

+ acetate + CoA + H20

-+

acetyl-CoA

+ AMP + 2 Pi

(10)

The extra driving forces available from the coupling of Eqs. ( 8 ) and (9)

7a. Different experimental conditions (pH, pMg) were used in the studies of reactions needed to calculate the A G L for hydrolysis of acyl adenylates; therefore, only approximate values are obtainable. The AGL. values reported here were derived from the accompanying tabulation. ~~

Reaction

+

+

(a) ATP acetate -+ acetyl-P ADP (b) Acetyl-P CoA -+ acetyl-CoA Pi (c) ADP Pi + ATP Hz0 (d) ATP HzO -+ AMP PPi (el Acetyl-CoA imidazole --t acetyl-imidazole CoA imidaeole (f) Acetyl-imidazole AMP -+ acetyl-AMP ( g ) ATP acetate -+ acetyl-AMP PPi (h) Acetate AMP --t acetyl-AMP HzO (i) Acetyl-CoA AMP --* acetyl-AMP CoA (j) Acetate Pi -+ acetyl-P H20

+ +

+

+

+

+

+

+

+

+ +

+ + +

+

+

+

+

AGL (kcal mole-')

Ref.

+3.1 -3.0

8 9

-11.0 +5.7 0.0 +3. 6a +14.6O +5.7a +12.00

4 10 11

Values calculated as follows: (g) = sum of Eqs. (a) through (f); (h) = ( g ) plus reverse of (d); (i) = (e) (f); (j) = (a) (c). 0

+

+

1.

ADENYLYL TRANSFER REACTIONS

5

may be of critical importance in some biosynthctic processcs, viz., in the activation of sulfate (see Section V). I n view of the above considerations it is not surprising that the adenylyl transfer capacity of ATP plays a prominent role in biological processes. In addition to numerous biosynthetic reactions in which only the energy of ATP bond cleavage is utilized, as in the synthesis of acetylCoA by mechanism B, the adenylyl transfer potential of ATP is used directly in the synthesis of compounds containing adenylyl moieties, viz., various coenzymes (12) and nucleic acids. Furthermore, the adenylyl transfer function is implicated in the allosteric alteration of enzyme structure which plays a dominant role in cellular regulation and also in the detoxification of various antibiotics. A thorough discussion of adenylyl group transfer reactions would therefore embody a large segment of biochemistry, including certain aspects of the synthesis of fatty acids, proteins, nucleic acids, polysaccharides, coenzymes, sulfur amino acids, and cellular regulation. Such an exhaustive treatment in a single chapter is precluded by limitations of space; furthermore, it is not justified in view of the fact that entire chapters have been devoted to some of these specific areas of ATP metabolism in past volumes of “The Enzymes” and others are the subject of separate chapters in a forthcoming volume of “The Enzymes” which is to be concerned exclusively with ATP-linked syntheses and processes. It is therefore inappropriate here to attempt an in-depth coverage of all aspects of adenylyl transfer phenomena. Instead an abbreviated discussion of various types of adenylyl transfer reactions will be made for the purpose of drawing attention to the central importance of such reactions in intermediary metabolism. For purposes of discussion these reactions can be grouped into the following categories : (a) reactions involving activation of carboxyl groups, (b) reactions to form phosphodiester derivatives of adenosine monophosphate, (c) reactions to form adenosine pyrophosphate esters, (d) reactions involving imidol group modification, (e) reactions concerned with activation of sulfate and other polyanionic inorganic com-

8. I. A. Rose, M. Glunberg-Manago, S. R. Korey, and S. Ochoa, JBC 211, 737 (1954). 9. H. U. Bergmeyer, G . Hols, H. Klotzch, and G. Lang, Biochem. Z. 338, 114 (1963). 10. E. R. Stadtman, in “The Mechanism of Enzyme Action” (W. D. McElrog and B. Glass, eds.), p. 581. Johns Hopkins Press, Baltimore, Maryland, 1954. 11. W. B. Jencks, BBA 24, 227 (1957). 12. A. Kornberg and W. W. Pricer, Jr., JBC 204, 329 (1953).

6

E. R. STADTMAN

pounds, and ( f ) reactions involving covalent alteration of functional groups of proteins.

II. Carboxyl Group Activation

A. ACTIVATIONOF FATTY ACIDSAND AMINOACIDSAND GENERAL CONSIDERATIONS Studies with animal enzyme systems led eventually to the discovery that the synthesis of acetyl-CoA is coupled to the cleavage of ATP to form AMP and PPi (13,14). Berg (15-17) purified a similar enzyme system from yeast and studied the mechanism of the reaction. He concluded that acetyl-CoA formation involves a two-step process in which acetyl-AMP is an intermediate, i.e., by mechanism B. In the meantime evidence has been obtained indicating that acyl-AMP intermediates are also involved in the biosynthesis of acyl-CoA derivatives of long chain fatty acids (3,lS-21), benzoate and phenylacetate (22-26), oxalate (27),and cholate (28,29).Moreover, aminoacyl adenylates are intermediates in the biosynthesis of all aminoacyl tRNA’s (30-35) and in the RNA-independent synthesis of carnosine (36-39) and antibiotic polypeptides (.@-&). Acyladenylates are also involved in the conversion of pantoate and p-alanine to pantothenic acid (46) and in the covalent attachment of biotin (47) and lipoic acid (48-50)to the c-amino groups of specific lysyl residues in biotin- and lipoate-containing enzymes. All of these biosynthetic events can be described by a generalized twoF. Lipmann, M. E. Jones, s. Black, and R. M. Flynn, JACS 74, 2384 (1952). M. E. Jones, S. Black, R. M. Flynn, and E. Lipmann, BBA 12, 141 (1953). P. Berg, JBC 222, 991 (1956). P. Berg, JBC 222, 1015 (1956). P. Berg, JBC 222, 1025 (1956). H. R. Mahler, S. J. Wakil, and R. M. Bock, JBC 204, 453 (1953). H. Beinert, .D. E. Green, P. Hele, H. Hift,, R. W. von Korff, and C. V. Ramakrishaan, JBC 203, 35 (1953). 20. W. P. Jencks, “Methods in Enzymology,” Vol. 5, p. 467, 1962. 21. W. P. Jencks and F. Lipmann, JBC 225, 207 (1957). 22. C. H. Lee Peng, BBA 22, 42 (1956). 23. M. Whitehouse, H. Moeski, and S. Gurin, JBC 225, 813 (1957). 24. K. Moldave and A. Meister, JBC 229, 463 (1957). 25. K. Moldave and A. Meister, BBA 24, 645 (1957). 26. K. Moldave and A. Meister, BBA 25, 434 (1957). 27. J. Giovanelli, BBA 118, 124 (1966). 28. W. H. Elliot, BJ 62, 427 (1956). 13. 14. 15. 16. 17. 18. 19.

1. ADENYLYL

7

TRANSFER REACTIONS

step mechanism (51) similar to that initially proposed by Berg (16) to account for acetyl-CoA formation [Eqs. (11) and (12)]. E

i + i i

+R

-0-

E.R -0-

1

ATP

-0-Ad

r ; p

E-RC-0-

b-

+X

~

4-Ad

+ PPC

(11)

0

e

R -X

+ AMP + E

(12)

0-

The overall coupled reaction is given by Eq. (13) 0 R-

e

-0-+X+ATP=R

8

-X+AMP+PPi

in which X is the ultimate acyl acceptor. As shown in Table 11, X may 29. H. Cantrenne, JBC 189, 227 (1951). 30. M. P. Stulberg and G. D. Novelli, “The Enzymes,” 2nd ed., Vol. 6, p. 401, 1962. 31. G . D. Novelli, Annu. Rev. Biochem. 36, Part 11, 449 (1967). 32. J. E. Allende and C. C. Allende, “Methods in Enzymology,” Vol. 20, Part C, p. 210, 1971. 33. R. B. Loftfield, Prog. NucE. Acid Res. Mol. Biol. 12, 87 (1972). 34. A. Meister, “Methods in Enzymology,” Vol. 6, p. 757, 1963. 35. A. Meister, “Biochemistry of the Amino Acids,” 2nd ed., Vol. 1. Academic Press, New York, 1965. 36. G. D. Kalyankei and A, Meister, JACS 81, 1575 (1959). 37. G. D. Kalyankei and A. Meister, Fed. Amer. SOC.Exp. Biol. Fed. Proc., 18, 256 (1957). 38. G. D. Kalyandei and A. Meister, JBC 234, 3210 (1959). 39. G . D. Kalyankei and A. Meister, “Methods in Enzymology,” Vol. 17B, p. 109, 1971. 40. H. Kleinkauf, R . Roskoski, Jr., and F. Lipmann, Proc. Nat. Acad. Sci. U . S . 68, 2069 (1971). 41. F. Lipmann, in “Chemical Evolution and the Origin of Life” (R. Bunet and C. Ponnamperum, eds.), p. 381. North-Holland Publ., Amsterdam, 1971. 42. F. Lipmann, Science 173, 875 (1971). 43. R. Roskoski, Jr., W. Gevers, H. Kleinkauf, and F. Lipmann, Biochemistry 9, 4839 (1970). 44. H. Kleinkauf and W. Gevers, Cold Spring Harbor Symp. Quant. Biol. 34, 805 (1969). 45. W. Gevers, H. Kleinkauf, and F. Lipmann, Proc. Nat. Acad. Sci. U . S. 63, 1335 (1969). 46. W. K. Maas and G . D. Novelli, ABB 43, 236 (1953). 47. M. D. Lane, K. L. Rominger, D. L. Yang, and F. Lynen, JBC 239, 2865 (1964). 48. L. J. Reed, F. R. Leach, and M. Koike, JBC 232, 123 (1958). 49. L. J. Reed, Proc. Int. Symp. Enzyme Chem., 1967, p. 71 (1958). 50. L. J. Reed, M. Koike, M. E. Levitch, and F. R. Leach, JBC 232, 143 (1958). 51. W. D. McElroy, M. Deluca, and J. Travis, Science 157, 150 (1967).

8

E. R. STADTMAN

be CoA, tRNA, an enzyme sulfhydryl group, the €-amino group of a lysyl residue of a protein, or an amino acid. The enzyme, E, has both adenylyl and acyl transfer functions. It catalyzes (a) transfer of the adenylyl group from ATP to the carboxyl group of an acid (acyl activation) and (b) transfer of an acyl group from the acyl adenylate to the acceptor X. Since reaction (11) is slightly endergonic [AGE,,. = +3.6 kcal mole-' (7a)l the enzyme also catalyzes transfer of the adenylyl group from acyl adenylates to PPi [reverse of Eq. ( l l ) ] . When X is CoA, tRNA, or a protein sulfhydryl group, Eq. (12) is also readily reversible; however, when X is an amino group, Eq. (12) is essentially irreversible under physiological conditions. Because the acyl-AMP intermediates do not dissociate from their specific enzymes, direct proof for their formation is not generally available. Nevertheless, as pointed out by Berg (16),their existence is inferred by the following characteristics: 1. I n the absence of acceptor X, the enzyme catalyzes an exchange between PPi and ATP [as a result of reversibility of Eq. ( l l ) ] . 2. Exchange of AMP into ATP does not occur unless the carboxylic acid and acceptor X are both present [as a result of reversibility of Eq. (13) 1. 3. I n the absence of X, the enzyme catalyzes the synthesis of ATP from added acyl-AMP and PPi; this is the result of reversibility of Eq. (11) and also because synthetic acyl-AMP is a substrate for the enzyme even though acyl-AMP generated by Eq. (11) does not readily dissociate from the enzyme. 4. I n the absence of the carboxylic acid, the enzyme catalyzes the formation of acyl-X from added acyl-AMP and X. 5. I n the absence of X, but in the presence of hydroxylamine, the enzyme catalyzes the formation of acyl hydroxamate from ATP and the carboxylic acid; this is because NHzOH reacts nonenzymically with acyl-AMP to form acyl hydroxamate and AMP. 6. Finally, as first shown by Boyer ( 5 l a ) in the overall Eq. (13), one atom of oxygen is transferred from the carboxyl group of the acyl donor acid to the phosphoryl group of AMP. ATP

+ RA

-180-

e

-+

R

L i4 - A d 180-

A-

-0-Ad

-1Q-

A'SO

'80

R-

i

'SO

180

+X

I1

-+

R-C-X

+ PP,

(14)

0

I1 + "O-P--O--Ad

A-

(15)

1. ADENYLYL

TRANSFER REACTIONS

9

Whereas this oxygen transfer is undoubtedly the best evidence for the formation of acyl-AMP intermediate, it is more difficult to determine experimentally. Therefore, other criteria, usually carboxylic acid-dependent incorporation of 32PPi into ATP or hydroxamate formation in the presence of NH,OH, have been taken as presumptive evidence for the formation of an acyl-AMP intermediate (52-109). 51a. P. D. Boyer, 0. J. Koeppe, and W. W. Luchsinger, JACS 7 4 356 (1956). 52. T. Okamoto and Y . Kawade, BBA 145, 613 (1967). 53. L. T. Webster, Jr., JBC 238, 4010 (1963). 54. L. T. Webster, Jr., and F. Campagnari, JBC 237, 1050 (1962). 55. P. R. Krishnaswamy and A. Meister, JBC 238, 405 (1960). 56. K. K. Wong and K. Moldave, JBC 235, 694 (1960). 57. J. E. Allende, C. C. Allende, M. Gatica, and M. Matamala, BBRC 16, 342 (1964). 58. A. T. Norris and P. Berg, Proc. Nut. Acad. Sci. U. S. 52, 330 (1964). 59. M. Yarus and P. Berg, JMB 28, 479 (1967). 60.C. H. Grosjean and J. Vanhembeeck, Arch. Znt. Physiol. Biochim. 75, 359 (1967). 61. P. Rouget and F. Chapeville, Eur. J . Biochem. 4, 310 (1968). 62. J. Waldenstrom, Eur. J . Biochem. 5, 239 (1968). 63. D. I. Hirsh, JBC 243, 5731 (1968). 64. H. Bluestein, C. C. Allende, J. E. Allende, and G. Cantoni, JBC 243, 4693 (1968). 65. C. C. Allende, H. Chaimovich, M. Gatica, and J. E. Allende, JBC 345, 93 (1970) . 66. S. Chousterman, F. Sonico, N. Stone, and F. Chapeville, Eur. J . Biochem. 6, 8 (1968). 67. U. Lagerkvist and J. Waldenstrom, JBC 240, PC2264 (1965). 68. H. Kleinkauf, W. Gevers, and F. Lipmann, Proc. Nut. Acad. Sci. U . S. 62, 226 (1969). 69. W. P. Jencks, “Methods in Enzymology,” Vol. 6, p. 762, 1963. 70. W. P. Jencks, “The Enzymes,” 2nd ed., Vol. 6, p. 373, 1962. 71. A. Kornberg, Advan. Enzymol. 18, 191 (1957). 72. A. Meister, Proc. Int. Pharmacol. Meet., l s t , 1961 Vol. 6, p. 77 (1962). 73. A. Mehler, “‘Methods in Enzymology,” Vol. 20, Part C, p. 203, 1971. 74. J. E. Allende and C. C. Allende, “Methods in Enzymology,” Vol. 20, Part C, p. 210, 1971. 75. W. Gevers, H. Kleinkauf, and F. Lipmann, Proc. N u t . Acad. Sci. U . S. 60, 269 (1968). 76. A. Bock, Arch. Mikrobiol. 68, 165 (1969). 77. S. K. Mitra and A. H. Mehler, JBC 242, 5490 (1967). 78. I. Hirschfield and H. J. P. Bloemers, JBC 244, 2911 (1969). 79. R. B. Loftfield and E. A. Eigner, BBA 130, 426 (1966). 80. W. Seifert, G. Nass, and W. Zillig, JMB 33, 507 (1968). 81. H. Hayashi, J. R. Knowles, J. R. Katze, J. LaPointe, and D. Soll, JBC 245, 1401 (1970). 82. R. Stern and A. H. Mehler, Biochem. Z . 342, 400 (1965). 83. R. D. Marshall and P. C. Zamecnik, BBA 198, 376 (1970). 84. J. R. Katze and W. Konigsberg, JBC 245, 923 (1970).

10

E. R. STADTMAN

In a number of instances, formation of an acyl-AMP intermediate has been confirmed by direct isolation of the acyl-AMP-enzyme complex (58-56).Webster (54) utilized Sephadex gel filtration to isolate the acetyl-AMP-enzyme complex formed when acetate :CoA ligase was incubated with ATP and acetate in the absence of CoA. The isolated complex reacted either with PPi to form ATP and acetate or with CoA to form acetyl-CoA and AMP (54).In the meantime, the technique has been used to isolate the acyl-AMP-enzyme complex of other fatty acids and various amino acids (6748) (see Table 11). Although it is evident that under in vitro conditions some aminoacyl-AMP-enzyme complexes are formed, Loftfield (33) cautioned that the two-step mechanism [Eqs. (11)-(13) ] for the synthesis of aminoacyl tRNA may not represent the physiological mechanism. He summarized a number of facts inconsistent with this mechanism. For example, (1) The association constant for the tRNA-enzyme complexes are such that at physiological concentrations of the macromolecules, essentially all of the enzyme is in the form of enzyme-tRNA complex; and (a) in some 85. R. Rigler, E. Cronvall, R. Hirsch, U. Packmann, and H. G. Zachau, PEBS Lett. 11, 32d (1970). 86. H. Heider, E. Gottschalk, and F. Cramer, Eur. J. Biol. 20, 144 (1971). 87. M. Rouge, BBA 171, 342 (1969). 88. E. C. Preddie, JBC 244, 3958 (1969). 89. G. Lemairc, R. van Rapenbusch, C. Gross, and B. Labouesse, Eur. J . Biol. 10, 336 (1969). 90. G. Lemaire, M . Dorirzi, G. Sportorno, and B. Labouesse, Bull. SOC.Chim. B i d . 51, 495 (1969). 91. D. R. Joseph and K. H. Muench, JBC 246, 7610 (1971). 92. D. R. Joseph and K. H. Muench, JBC 246, 7602 (1971). 93. J. M. Clark and J. P. Eyzaquire, JBC 237, 3698 (1962). 94. R. Calendar and P. Berg, Biochemistry 5, 1681 (1966). 95. R . Calendar and P. Berg, Biochemistry 5, 1690 (1966). 96. R. F. B. Diller and G. M. Tener, Can. J. Biochem. 49, 822 (1971). 97. H. L. James and E. T. Bucovar, JBC 244, 3210 (1969). 98. M. P. Duntscher, JBC 242, 1123 (1967). 99. W. R. Folk, Biochemistry 10, 1728 (1971). 100. R. L. Hendrickson and B. S. Hartley. BJ 105, 17 (1967). 101. C. J. Burton and B. S. Hartley, BJ 108, 281 (1968). 102. M. P. Stulberg, JBC 242, 1060 (1967). 103. M. H. J. E . Kosakowaski and A. Bock, EUT.J. Biol. 12, 67 (1970). 104. D. V. Santi, P. V. Danenberg, and P. Slatterly, Biochemistry 10, 4804 (1971). 105. M. H. Makman and G. L. Cantoni, Biochemistry 4, 1434 (1955). 106. F. Fasiolo, N. Bedfort, Y. Boulanger, and J. P. Ebel, BBA 217, 305 (1970). 107. J. Preiss and P. Handler, JBC 233, 493 (1958). 108. T. S. Papas and A. H. Mehler, JBC 245, 1888 (1970). 109. T. P. Bennett, JBC 244, 3182 (1969).

1. ADENYLYL

TRANSFER REACTIONS

11

instances (with arginine, glutamine, and glutamate) in vitro synthesis of enzyme- (aminoacyl-AMP) complexes cannot be demonstrated in the absence of the acceptor tRNA. Whereas these and the other arguments summarized by Loftfield are well taken, all such arguments can be explained by assuming that in addition to its role as a substrate, tRNA may have an allosteric role in provoking enzyme comformations that facilitate the reaction of ATP and amino acid with the enzyme to yield the acyl-AMP-enzyme derivative. Such a role of the acceptor molecule has been clearly established in the case of argininosuccinate biosynthesis (see Section VI,A). I n any event, it seems unlikely that the physiological mechanism differs significantly in principle from that outlined above. Distinction between a two-step reaction and the alternative concerned mechanism can be very subtle indeed. The above discussion is not intended to represent an exhaustive analysis of the huge body of information that is available on those enzymes concerned with the activation of carboxylic acids. The biosynthesis of acyl-CoA compounds has been extensively reviewed by Jencks (20, 69, 70), Kornberg ( 7 1 ) , Meister (72), and McElroy et al. (51);this was the topic of a chapter in an earlier edition of “The Enzymes” (70) and is to be the subject of a separate chapter in the next volume of “The Enzymes.” Similarly, studies on aminoacyl tRNA synthetases have been reviewed by Novelli (SI), Meister (34, 35), Loftfield (W), Mehler (73), Allende and Allende ( 7 4 ) , and Stulberg and Novelli (30); this subject too, was reviewed in an earlier edition of “The Enzymes” (30) and will comprise a separate chapter in a forthcoming volume of “The Enzymes.” Therefore, the adenylyl transfer function associated with these processes will not be considered further here. Also, since relatively little detailed information is available on the enzymes catalyzing activation steps 3-7 in Table 11, these will not be considered further.

B. NUCLEICACID-INDEPENDENT PEPTIDE SYNTHESIS In their studies on the biosynthesis of gramicidin S and the tyrocidines by enzyme preparations from Bacillus brevis, Lipmann et al. (40-45, 68, 75, 110) discovered a novel mechanism for nucleic acidindependent synthesis of peptides in which aminoacyl adenylates are intermediates. Gramicidin S is a cyclic decapeptide composed of two 110. R. Roskoski, Jr.? H. Kleinkauf, W. Gevers, and F. Lipmann, Biochemitry 9, 4846 (1970).

TABLE I1 CARROXYL GROUPACTIVATION

0

ATP

II

+ R-C-4-

0

+X

-+

AhIP

/I + R--CX + PPi ~~

Activation Step

1 2

3 4 5 6

7 8 9

10 11

0

e

Acyl donor (R 4) Acetate" Long chain fatty acids Benzoate Phenylacetate Oxalate Cholate &Ahnine Luciferin Pantoate Alanine Arginine

Acyl acceptor (X) CoA CoA CoA CoA CoA CoA Histidine CoA 8-Alanine tRNAAla tRNAA*g

Enzyme (E) Acetate: CoA l i m e (AMP) Fatty acid: CoA lyase (AMP) Benzoate: CoA ligase (AMP) Phenylacetate: CoA ligase (AMP) 0xalate:CoA.ligase (AMP) Cholate: CoA ligase (AMP) p-Alanine: histidine ligase (AMP) Luciferin CoA ligase (AMP) Pantoate:&alanine ligase (AMP) Almine: tRNAA" ligase (AMP) Arginine: tRNAAreligase (AMP)

Ref. 13-17,64 5, 18-22

2.4-26,29 24-86 27 28,29 36,239 57, 128-130 46 76 62,77,78

H

F

; tl c3

Fx

Y 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 a

Glycine Histidine0 Isoleucinea Leucinea Lysinea Seri;rea Threoninea Tryptophan Tyrosinea Valinea Aspartate Cysteine Glutamate Glutamine Methionine Phenylalanine Proline Desamido-DPN Lipoate Biotin

Glycine: tliNAGly ligase (AMP) Histidine: tRNAA" ligase (AMP) Isoleucine: tRNA1" ligase (AMP) Leucine: tRNALeU ligase (AMP) Lysine: tltNALyaligase (AMP) Serine: tRNAser ligase (AMP) Threonine: tRNAThr ligase (AMP) Tryptophan: tRNAny ligase (AMP) Tryosine: tRNATyr ligase (AMP) Valine:tRNAV8'ligase (AMP) Aspartate: tRNAAoP ligase (AMP) Cysteine: tRNACY8ligase (AMP) Glutamate: tRNAG1" ligase (AMP) Glutamine: tRNAG1nligase (AMP) Methionine: tRNAMet ligase (AMP) Phenylalanine: tIZNAPhe ligase (AMP) Proline: tRNAProligase (AMP) Desamido-DPN :glutamine-amido ligase (AMP) Lipoate: N*-enzymeligase (AMP) Biotin: N*-enzymeligase (AMP)

62 87 5840, 79 61, 80, 81 62, 82, 83

G

64,8447 67, 63, 65 88-92

e rn P

1S, 58,93-95 52, 67 96 97 98 99

2 8*: r

2c1 3 rn

M P c3

8ti: u)

loo, 101

102-106 108 107

@-50 47

Cases in which enzyme-acyl-AMP intermediates have been definitely established.

CL

w

14

E. R. STADTMAN

identical sequcnrcs of fiw different amino acids joined together as follows: L-Leti-o-P he-IrPro-IrVai-cOrn

t

LOrn-LVal-LPro-D-Phe-LLeu

I

Tyrocidine is a decapeptide composed of up to nine different amino acids joined together in the sequence: 1 2 . 1 4 5 o-PkycPro-bPhe-D-Phe-bAy tL.kIrOrn-L-Val-cTyr-L-Gln.L 1

0

9

8

7

6

The tyrocidines are, in fact, a mixture of decapeptides resulting from a lack of enzyme specificity permitting substitutions of tryptophan for phenylalanine in positions 3 and 4, and of either tryptophan or phenylalanine for tyrosine in position 7, leading to variants (111, 112). A soluble enzyme system capable of catalyzing the synthesis of gramicidin S from ATP and a mixture of the component amino acids was developed in several laboratories (75,113-117).This system has since been highly purified and was separated by Sephadex G-200 chromatography into two complimentary fractions (75, 1 1 4 ) . The heavier Fraction I (MW 280,000) catalyzes amino acid-dependent ATP-32PP+ exchange in the presence of any one of the four amino acids proline, valine, ornithine, or leucine (75, 115). The lighter Fraction I1 (MW lO0,OOO) catalyzes ATP-32PPi exchange only in the presence of phenylalanine (75, 115, 116). I n addition, Fraction I1 catalyzes phenylalaninedependent ATP-AMP exchange ( 7 5 ) .Neither fraction by itself catalyzes peptide formation, but together they catalyze gramacidin S formation in the presence of ATP and all five amino acids (75, 1 1 4 ) . In the presence of Mg?', fraction I1 by itself catalyzes a reaction between L-phenylalanine and ATP to form an enzyme-bound D-phenylalanyl-AMP derivative [Eq. (16) ] and in addition catalyzes transfer of the D-phenylalanyl group from the aminoacyl-AMP intermediate 111. B. Mack nnd E. L. Tatum, Proc. Not. Acad. Sci. U . S. 52,876 (1964). ' . Sakamoto, T. Suzuki, and K. Kurnhnshi, BBA 169, 520 112. K. Fujikawn, 1 (1968). 113. K. Kurahashi, Abstr., Znt. Congr. Biochem. 6th, 1901, p. 37 (1963). 114. S. Tomino, M. Ynmada, H. Itoh. and K. Kurahashi, Biochemistry 6, 2552 (1967). 115. H.Itoh, M. Ynmada, S. Ionino. and K. Kurahashi, J . Biochem. ( T o k y o ) 64, 259 (1968). 116. M. Yamada. and K. Kurahashi, J . Biochem. ( T o k y o ) 63, 59 (1968). 117. T.L. Berg, L. 0. Frplholm, and S. G. Laland, BJ 96, 43 (1965).

1.

15

ADENYLYL TRANSFER REACTIONS

to a specific sulfhydryl group on thc same protcin to form a protein bound D-phenylalanyl t,hiolester derivative [Eq. (17) 3 (75). The latter can react with a second equivalent of ATP and L-phenylalanine to form a protein derivative containing one equivalent each of phenylalanyl thiolester and phenylalanyl-AMP [Eq. (18)] (75).

+ ATP

+ L-Phenylalanine

D-Phenylalanyl-AMP

-

D-Phenylalanyl-AMP + PPi

-

S-D-Phenylalanyl

+

(16)

AMP

S-D-Phenylalanyl

+ ATP +

L-Phenylalanine

SH + 2ATP

+

2 L-Phenylalanine

-

S-O- Phenylalanyl

D-Phenylalanyl-AMP

+ ZPP, + AMP

(19)

As illustrated in Fig. 1, Fraction I catalyzes a completely analogous series of reactions with any one or all of the other four amino acids L-leucine, L-proline, L-valine, and L-ornithine. The reaction with each substrate is completely independent of the others and when all substrates are present simultaneously (in the absence of Fraction 11) two equivalents of each substrate are bound to Fraction I, one of each in thiolester linkage to amino acid specific protein sulfhydryl groups and one of each as the aminoacyl adenylate derivative. These results suggest that Fraction I is a multienzyme complex composed of a t least four separate enzymes, each specific for a different one of the four amino acids (75). I n addition to an amino acid specific sulhydryl group on each of the four different amino acid activating enzymes, Fraction I also contains an acyl carrier protein possessing a pantetheine group, presumably bound to the protein in phosphodiester linkage. As noted above, no polymerization of the aminoacyl thiolester derivatives occurs with either Fraction I or Fraction I1 alone. Polymerization is initiated by transfer of the Dphenylalanyl moiety of Fraction I1 to form a dipeptidyl (n-phenylalanyl-

16

E. R. STADTMAN r

r‘ S-Prolyl

S-Prolyl

Pm Prolyl- AMP

SH

S-Valyl a

S-Omithyl

Valyl- AMP

PPj

+

+ Ornithine Ornithyl-AMP

a

4

Ornithine ATP d l Om

L

SH

S-Ornithyl

Omithyl- AMP

s-Leucyl

Leucyl- AMP

a

L

Leucyl- AMP

FIG. 1. Amino acid activation by the multienzyme complex Fraction I. The linear array of four oblong segments depicts the multienryme complex comprising Fraction I. Each segment represents a separate enzymic activity that catalyzes the activation of a single amino acid with the indicated specificity. The sites designated ‘ I >, a on each segment represents the site of attachment of the aminoacyl adenylate derivative. After formation of the aminoacyl adenylate a t site “a,” the aminoacyl group is transferred to the sulfhydryl group to form the corresponding enzyme bound aminoacyl thiolester derivative. Although not shown, the multienzymc complex (Fraction I) also contains an acyl carrier protein which is the site of polypeptide synthesis, as described in the text.

L-prolyl) -thiolester derivative bound to the proline activating enzyme of Fraction I ( 4 2 ) . This is followed by a thiol transesterification reaction in which the dipeptidyl group is transferred to the sulfhydryl group of the pantetheine moiety of the acyl carrier protein in Fraction I. The dipeptide is subsequently transferred to the amino group of the next aminoacyl thiolester of Fraction I to form a tripeptidylthiolester derivative bound to the L-valine amino acid activating enzyme in Fraction I. This is followed by a second thiol transesterification in which the tripeptidyl group (D-Phe-L-Pro-L-Val-) is transferred to the pantetheine sulfhydryl group. The tripeptidyl-S-pantetheine group then swings to the L-ornithine activating enzyme where the process is repeated and a tetrapeptide is formed. As Lipmann points out (&), pantetheine performs a switch or translocation function involving alternating condensation and transthiolation leading ultimately to the formation of the thiolester derivative of the pentapeptide D-Phe-L-Pro-L-Val-L-Orn-L-

1. ADHNYLYL

17

TRANSFER REACTIONS

Leu. By a mechanism that is not understood, two such pentapeptidyl thiolester derivatives finally condense to form the cyclic decapeptide gramicidin S. The synthesis of tyrocidine by extracts of B. brevis occurs by an analogous mechanism ; however, since this antibiotic is composed of nine different amino acid residues and contains no repeating sequences, the process of alternate condensation and transthiolation continues until the complete decapeptide is formed. The enzyme system catalyzing tyrocidine synthesis has been resolved into three separate protein fractions: a light fraction (MW 100,000) which possesses phenylalanine activating enzymic activity, a fraction of intermediate size (MW 230,000) which specifically activates L-proline, and a heavy component (MW 460,000) which activates all of the other amino acids of tyrocidine and additionally contains the pantetheine-linked acyl carrier protein (42).

C. ACYLATION OF

THE

NE-LYSYLRESIDUESOF ENZYMES

1. Lipoate Activating Enzyme

Reed and his associates (49, 118, 119) have shown that lipoic acid is covalently bound in amide linkage to a €-amino group of a lysyl residue of a-ketoacid dehydrogenases. The enzyme-bound lipoate is concerned with acyl generation, acyl transfer, and electron transfer functions involved in the oxidation of a-ketoglutarate and pyruvate by their respective multienzyme complexes (48-50, 120). Attachment of the lipoyl group to the lipoate reductase-transacetylase component of this complex is catalyzed by an activating enzyme system that has been partially purified from E. coli and Streptococcus jaeculis (48, 118, 119, 121). The enzyme system from 8.faecalis was resolved into two protein components, E, and E,, which together are needed to catalyze the attachment of lipoic acid to the dehydrogenase. The E, fraction apparently catalyzes an adenylyl transfer reaction [Eq. (20)] since, in the presence of ATP, lipoate, Mg2+, and hydroxylamine, the enzyme catalyzes the 0

El

+ ATP + CHr-CH&H(CH2)4 AH

I

SH

e

0

0

II I

0-+ EI.CHYCH&H(CH~)~-0-P-O-Ad

I

SH

AH

0-

+ PPi

(20)

118. L. J. Reed, Advan. Enzymol. 18, 319 (1957). 119. H. Nawa, W. T. Brady, M. Koike, and L. J. Reed, JACS 82, 896 (1960). 120. L. J. Reed, Vitam. Horm. ( N e w Yo&) 20, 1 (1962). 121. M. Koike, L. J. Reed, and W. R. Carroll, JBC 235, 1924 (1960).

18

E. R. STADTMAN

formation of lipoylhydroxamate, PPi, and AMP (4.8, 49). The second enzyme fraction, E,, is apparently concerned with transfer of the lipoyl group from El-lipoyl-AMP to the acceptor r-NH2-lysyl group of the a-ketoacid dehydrogenase. The intermediate role of lipoyl-AMP is further supported by the fact that synthetic lipoyl-AMP can replace ATP and lipoate in the overall reaction. In this case, both E, and E2 are required, suggesting that the El-lipoyl-AMP complex is an obligatory intermediate in the lipoyl transfer part of the reaction. Proteolytic degradation of the pyruvate and a-ketoglutarate dehydrogenase complexes containing bound 35S-lipoic acid led to the isolation of lipoyl-containing peptides having the common amino acid sequenceAsp-Nf-lipopyl-Lys (122) .

2 . Biotin Activating Enzyme The enzyme methylmalonyl-CoA-pyruvate carboxytransferase (123) is one of a large class of biotin-dependent enzymes whose active form contains biotin attached in amide linkage to an Nf-lysyl group of the enzyme (47,124-1.26).A biotin activating enzyme, biotin: Nf-apotranscarboxylase ligase (AMP), that catalyzes the ATP-dependent attachment of biotin to the apotranscarboxylase [Eq. (21)] was highly purified from extracts of Propionibacteriunz shermanii by Lane and Lynen (127). ATP

-

+ Biotin + Ncapotranscarboxylase M d + biotinyl-Ng-transcarboxylase + AMP + PPi

(21) The formation of a biotinyl-AMP-enzyme complex as an intermediate in the overall reaction is supported by the following observations: (a) The purified biotin:“-apoenzyme ligase catalyzes the ( + ) - biotindependent ATP-PPi exchange; (b) it catalyzes the synthesis of ATP from PPi and ( + ) - biotinyl-5’-AMP, and the synthesis of ( + ) - biotinyl-Nf-transcarboxylase from apotranscarboxylase and ( ) - biotinyl5’-AMP; (c) ( + ) - biotinyl-5’-AMP can replace ( + ) - biotin, ATP, and a divalent cation in holocarboxylase synthesis; and (d) the kinetics of these processes are consistent with the conclusion that biotinyl-AMP is an intermediate (47).

+

122. K. Diago and 1,. J. Reed, JACS 84, 666 (1962). 123. R. W. Swick and H. G. Wood, Proc. Nat. Acad. Sci. U . S. 46, 28 (1960). 124. H. G. Wood, H. Lochmuller, C. Riepertinger, and F. Lynen, Biochem. 2.337, 247 (1963). 125. M. D. Lane and F. Lynen, Proc. N a t . Acad. Sci. U . S. 49, 379 (1963). 126. D. P. Kosow and M. D. Lane, BBRC 7, 439 (1962). 127. M. D. Lane, D. L. Young, and F. Lynen, JBC 239, 2858 (1961).

1. ADENYLYL

19

TRANSFER REACTIONS

D. ADENYLYLATION OF LUCIFERIN AND DEHYDROLUCIFERIN McElroy et a2. (128-131) have demonstrated that highly purified firefly luciferase catalyzes adenylylation of luciferin (LH,) or dehydroluciferin (L) by Eqs. (22) and (23). The enzyme-bound adenylylated derivative of luciferin (E-LH,-AMP) reacts with molecular oxygen to

R

produce light and unknown products, whereas the E-L-AMP derivative cannot produce light (128). It was determined that the equilibrium constant for the dissociation of E-L-AMP to yield E + L-AMP [Eq. (24)] is 5 X and that the equilibrium constant for reaction (25) is 4 X 104 (128). From these values and the known free energy of hydrolysis of ATP to yield AMP and PPi [Eq. (26)], Rhodes and McElroy (128) estimated that the free energy of hydrolysis of the dehydroluciferyladenylate bond is 13.1 kcal mole-’ [i.e., the sum of reactions (24), (25), and (26)l. 128. 129. 130. 131.

W. W. M. W.

C. Rhodes and W. D. McElroy, JBC 233, 1528 (1958). D. McElroy, “Methods in Enzymology,” Vol. 6, p. 775, 1963. DeLuca, G. W. Wirtz, and W. D. McElroy, Biochemistry 3, 935 (1964). B. McElroy and H. H. Seliger, Advan. Enzymol. 25, 119 (1963).

20

E. R. STADTMAN AGO

+

L A M P E F! E-LAMP PPi E-L-AMP E ATP ATP He0 AMP PPi

+ +

+ +L + L-AMP + H 2 0 E AMP + L

(kcal mole-') 4-12.6 +7.2

-7.7 ~~

-

(24) (25) (26)

(27) Rhodes and McElroy assumed the free energy of hydrolysis of reaction (26) to be the same as that for the hydrolysis of ATP to yield ADP and Pi. As pointed out earlier (Table I) this assumption is not valid except under a very specific set of conditions. Under the conditions of their experiments (pH 7.1 and 5 X 10-3M MgZ+),the AGL. for Eq. (26) is probably somewhat greater than the value of -7.7 kcal assumed in the above calculations (see Table I) and the AGzb, for Eq. (24) would be correspondingly greater also (probably -14 to -15 kcal mole-'). I n any case the free energy of hydrolysis of dehydroluciferyladenylate is similar to the value of -14.6 kcal calculated for the hydrolysis of acetyladenylate (?'a). Probably because of its high affinity for luciferase, L-AMP is a, potent inhibitor of light emission provoked by LH2-AMP. This inhibitory effect can be overcome by the addition of CoA, owing to the capacity of luciferase to catalyze reversible transfer of the dehydrolucerifyl moiety of L-AMP to CoA [Eq. (28)] (13.2). E-L-AMP

+ CoA

E

-13.1

+ L-CoA + AMP

(28)

This reaction, and an ATP-dependent hydrolysis of enzyme-bound LAMP in the presence of pyrophosphatase (128), may have regulatory roles in the light emission process. For a further discussion of these possibilities, as well as other aspects of this extremely interesting enzyme system, the reader is referred to excellent reviews by McElroy et al. (61, 131).

111. Biosynthesis of Phosphodiester Derivatives of Adenosine

A. RNA SYNTHESIS During the last decade a vast literature has accumulated describing the isolation and properties of enzymes catalyzing the synthesis and degradation of polynucleotides. Since this work has been exhaustively reviewed in numerous articles, it is mentioned here only to draw attention to the fact that some of the reactions concerned with RNA metab132. R. Airth, W. C. Rhodes, and W. D. McElroy, BBA 27, 619 (1958).

1. ADENYLYL

21

TRANSFER REACTIONS

olism involve adenylyl group transfer mechanisms. Table I11 lists some of the more important enzymes studied to date. For more detailed discussion of these processes the reader is referred to several comprehensive reviews (71, 133-1 60). 1. RNA Polymerase

The DNA-dependent incorporation of adenylyl groups from ATP into RNA is catalyzed by RNA polymerases from many sources (133154). Although under appropriate conditions the purified enzymes can catalyze the synthesis of homopolymers composed exclusively of adenylic acid residues (i.e., poly A) (165, 156), their normal function is concerned with transcription of DNA templates. They are, therefore, able to catalyze ribonucleotidyl transfer from all four nucleoside triphosphate according to the general equation nATP

-

+ nUTP + nGTP + ~ C T PMns+,DNAMga+ RNA + 4 nppi

(29)

133. A. M. Michelson, “The Chemistry of Nucleosides and Nucleotides.” Academic Press, New York, 1963. 134. J. P. Richardson, Prog. Nucl. Acid Mol. Biol. 9, 75 (1969). 135. S. B. Weiss, “Methods in Enzymology,” Vol. 12, Part B, p. 555, 1968. 136. J. S. Krakow and W. J. Horsley, “Methods in Enzymology,” Vol. 12, Part B, p. 566, 1968. 137. J. S. Krakow and S. Ochoa, “Methods in Enzymology,” Vol. 6, p. 11, 1963. 138. J. Hurwitz, “Methods in Enzymology,” Vol. 6, p. 23, 1963. 139. M. Grunberg-Manago, “The Enzymes,” 2nd ed., Vol. 5, p. 257, 1961. 140. T. Kimhi and U. Z. Littauer, “Methods in Enzymology,” Vol. 12, Part. B, p. 513, 1968. 141. F. Lipmann, Advan. Enzyme Regul. 9, 5 (1971). 142. A. Sibitani, Prog. Biophys. Mol. Biol. 16, 17 (1966). 143. C. C. Richardson, Annu. Rev. Biochem. 38, 795 (1969). 144. E. K. F. Bautz, in “Molecular Genetics” (J. H. Taylor, ed.), Part 2, p. 213. Academic Press, New York, 1967. 145. E. P. Ceiduschek and R. Haselkorn, Annu. Rev. Biochem. 38, 647 (1969). 146. E. P. Geiduschek, in “Aspects of Protein Biosynthesis” (C. B. Anfinsen, ed.), Part A, p. 43. Academic Press, New York, 1970. 147. G . Schmidt, Annu. Rev. Biochem. 33, 667 (1964). 148. J. Hurwitz and J. T. August, Prog. Nucl. Acid Res. 1, 59 (1963). 149. R. L. Erikson and R. M. Franklin, Bacteriol. Rev. 30, 267 (1968). 150. R. M. Franklin, “Methods in Enzymology,” Vol. 12, Part B, p. 572, 1968. 151. J. Hurwitz, J. J. Furth, M. Anders, P. J. Ortiz, and J. T. August, Cold Spring Harbor Symp. Quant. Biol. 26, 91 (1961). 152. S. B. Weiss, Proc. Nut. Acad. Sci. U . S. 46, 1020 (1960). 153. U. Maitra and J. Hurwitz, Proc. Nut. Acad. Sci. U . S. 54, 815 (1965). 154. U. Maitra, Y. Nakata, and J. Hurwitz, JBC 242, 4908 (1967). 155. A. Stevens, JBC 244, 245 (1969). 156. M. Chamberlin and P. Berg, Proc. Nut. Acnd. Sci. U. S. 48, 81 (1962).

TABLE I11 ADENYLYLTRANSFER REACTIONS OF NUCLEIC ACIDMETABOLISM

Es. No. (29)

Enzyme RNA polymerase

Reaction catalyzed.

+ nUTP + nGTP + nCTP

nATP

oligonucleotide

(31)

Polynucleotide phosphorylase

nXDP

(XMP)n Mg*+

(32)

ATP(CTP): tRNA nucleotidyltransferase

tRNA

DNA

RNA

Mna+ or Mg*+

Ref.

+ 4 nPPi

133-138, 166-166

+ nPj

. . . pX + 2 CTP + ATP

4

133,139, 1.40

tRNA

. . . pXpCpCpA + 3 PPi

16.4-167

Mgr+

(33)

ATP: RNA adenylyltransferase

RNA

+ nATP +RNA-(AMP)n + nPPi

(34)

Nucleoside triphosphate: DNA nucleotidyltransferase

DNA

+ XTP +DNA-XMP + PPi

16.2,168, 175

Me+

18.4-188 ~

~~

X refers to any one of the following bases: adenine, cytidine, uridine, and guanine;pX in Eq. (32) refers to the fourth 5'-nucleotidyl residue from the terminal end of a fully active tRNA molecule.

PJ p

1. ADENYLYL

23

TRANSFER REACTIONS

The specific site of 5'-AMP incorporation are determined by complimentary sites on the DNA template. The adenylylation reaction itself involves a transfer of the adenylyl group from ATP to the unesterified (2,'-hydroxy group of the terminal nucleoside residue of the growing polynucleotide chain, Eq. (30), (153, 154). Analogous reactions are catalyzed by RNA-dependent RNA polymerase induced in phage-infected strains of E. coli (1.49, 157) and in virus-infected animal cells (1.69) *

x

x

x

x

x

x 0

II -0-P-0-Ad

(30)

1

P

0 MnZ+

+ PPi

2. Polynucleotide Phosphorylase Reversible transfer of adenylyl groups from ADP into polyribonucleotides is catalyzed by polynucleotide phosphorylase (139,1.40). The enzyme catalyzes both homopolymer and heteropolymer formation from the nucleoside diphosphates according to Eq. (31).

where X can be any single one or a mixture of the bases: adenine, uridine, guanine, or cytidine (139,150, 158, 169). The polymerization does not require a DNA template, but it is stimulated by oligoribonucleotides which serve as primers of the reaction (158-162).As in the case of RNA polymerase, polymerization involves sequential addition of the 5'-nucleotidy1 groups to the unesterified Cs'-hydroxyl group of the terminal nucleoside residue [cf. reaction (30)]. The extent of adenylylation is not template directed but is related to the relative concentration of ADP to other nucleoside diphosphates and of primers in the reaction mixture. In contrast to the reaction catalyzed by RNA polymerase [Eq. (3011, reaction (31) is freely reversible (139).This fact and the random nature of the nucleotide polymerization catalyzed by polynucleotide phos157. C. Weisaman, L. Simon, and S. Ochoa, R o c . Nut. Acad. Sci. U. S. 49, 407 (1963). 158. M. F. Singer, L. A. Hepple, and R. J. Hilmoe, JBC 235, 738 (1960). 159. M. F. Singer, R. J. Hilmoe, and L. A. Hepple, JBC 235, 751 (1960). 160. M. F. Singer, R. J. Hilmoe, and M. Grunberg-Manago, JBC 235, 2705 (1960). 161. M. F. Singer, L. A. Hepple, and R. J. Hilmoe, BBA 26, 447 (1957). 162. J. T. August, P. J. Ortiz, and J. Hurwitz, JBC 237, 3786 (1962).

24

E. R. STADTMAN

phorylase as well as other considerations have led to the conclusion that the normal function of the phosphorylase is to catalyze degradation of RNA rather than its synthesis. It is noteworthy that under physiological conditions phosphorolysis of polynucleotides to yield nucleoside diphosphate derivatives [reverse of Eq. (31) ] is 1-2 kcal mole-1 more exergonic than is pyrophosphorolysis to yield nucleotide triphosphates [reverse of Eq. (30)]. For this reason, phosphorolytic cleavage of RNA is more suitable for degradative purposes than is pyrophosphorolysis. However, the lack of reversibility of the RNA polymerase reaction (148,163) cannot solely result from its exergonic character since, as will be described later, pyrophosphorolytic cleavage of similar phosphodiester bonds is readily catalyzed by other nucleotidyltransferases (16‘4, 166). 3. ATP-Specific Polynucleotide Adenylyltransferases

In addition to the RNA polymerase and polynucleotide phosphorylase which for obvious reasons are able to catalyze 5’-nucleotide transfers from all four nucleoside triphosphates and nucleoside diphosphates, enzymes have been isolated from many sources that show a high degree of specificity for ATP as the nucleotidyl donor. Among the better characterized enzymes are the ATP (CTP) :tRNA nucleotidyltransferases (164-167) and the poly A polymerases (162,168-182). 163. U. Maitra and J. Hurwitz, JBC 242, 4897 (1967). 164. J. Preiss, M. Diekmann, and P. Berg, JBC 236, 1748 (1961). 165. V. Daniel and U. Z. Littauer, “Methods in Enzymology,” Vol. 12, Part U, p. 579, 1968. 166. V. Daniel and U. Z. Littauer, JBC 238, 2102 (1963). 167. J. J. Furth, J. Hurwitz, R. Krug, and M. Alexander, JBC 236, 3317 (1961). 168. S. J. S. Hardy and C. G. Kurland, Biochemistry 5, 3668 (1966). 169. K. J. Payne and J. A. Boezi, JBC 245, 1378 (1970). 170. M. E. Gottesman, Z. N. Canellakis, and E. S. Canellakis, BBA 81,34 (1962). 171. H. G. Klemperer, BBA 72, 416 (1963). 172. M. Edmonds and R. Abrams, JBC 237, 2636 (1962). 173. A. J. E. Colvill and M. Terzi, BBA 155, 394 (1968). 174. R. H. Burdon and R. M. S. Smellie, BBA 61, 633 (1962). 175. N. W. Wilkie and R. M. S. Smellie, BJ 109, 229 (1968). 176. P. R. Venkataraman and H. R. Mahler, JBC 238, 1058 (1963). 177. V. Daniel and U. Z. Littaner, J M B 11, 692 (1965). 178. K. J. Payne and J . A. Boezi, JBC 245, 1378 (1970). 179. L. I. Hecht, M. L. Stephenson, and P. C. Zamecnik, Proc. N o t . Acad. Sci. U. S. 45, 505 (1959). 180. E. Herbert and E. S. Canellakis, BBA 47, 85 (1961). 181. E. Herbert and E. S. Canellakis, “Methods in Enzymology,” Vol. 6, p. 28, 1963. 182. A. J. E. Colvill and M. Terzi, BBA 155, 394 (1968).

1.

25

ADENYLYL TRANSFER REACTIONS

a. ATP ( C T P ):tRNA Nucleotidyltransjerases. These enzymes catalyze synthesis of the -pCpCpA sequence on the 3' end of transfer RNA (16&167), Eq. (32), in which pX represents the fourth nucleotidyl group tRNA

. . . pX + 2 CTP + ATP

tRNA

. . . pXpCpCpA + 3 PPi

(32)

from the end of the fully active tRNA molecule. Since the pCpCpA sequence is a common end terminal sequence of all active tRNA molecules (164, 176), the physiological role of this enzyme in the final steps of tRNA synthesis seems assured. This function is further supported by the fact that tRNA molecules stripped of one or more of the terminal 3' residues (pCpCpA) are the only nucleotidyl group acceptors for the enzyme (164) and, in the case of the enzyme from E. coli, by the fact that C T P and A T P are the only active nucleotidyl donor substrates (164, 165). The enzyme from rat liver (165) utilizes UTP as a uridylyl donor provided CTP is absent. The enzyme has been purified from E . coli (164, 165) and from rat liver (165-177). Synthesis by the terminal pXpCpCpA sequence involves stepwise addition of each of the two cytidylyl groups followed by the adenylyl group. ATP is not required for addition of the cytidylyl groups to pX, nor is CTP required for the addition of an adenylyl group to . . . pXpCpC (164, 165, 17'7). Although it is formally similar to reaction (30) catalyzed by RNA polymerase, reaction (32) is freely reversible (164-166). I n the presence of inorganic pyrophosphate the purified enzyme catalyzes sequential pyrophosphorolysis of the terminal . . . pCpCpA nucleotidyl groups to form 2 moles of C T P and 1 mole of ATP (164, 165). Pyrophosphorolysis of other phosphodiester bonds of tRNA is not catalyzed (164-166). Since reaction (32) is freely reversible, it is obvious that the failure of RNA polymerase to catalyze pyrophosphorolysis of RNA cannot be attributed to an unfavorable equilibrium of the reaction.

b . Poly A Polymerase ( A T P :R N A Adenylyltransferase). Enzymes that catalyze multiple transfers of adenylyl groups from ATP to the 3' terminus of an RNA primer have been isolated from numerous sources (16.2, 168-175). As shown by Eq. (33) in Table 111, the products formed are inorganic pyrophosphate and RNA derivatives containing polyriboadenylic acid chains of up to 100 nucleotide residues covalently bound to the 3 terminus. The enzyme from E. coli catalyzes exchange of PPi into the P,y-phosphoryl groups of several different nucleoside triphosphates, but the polymerization reaction is relatively specific for ATP (162). Although the physiological significance of this enzyme has not been established, the fact that its activity is markedly reduced

26

E. R. STADTMAN

following infection of E . coli by T2,T4,T,,and T, phages (183) has led to the suggestion that its normal function is concerned with host ribosomal RNA synthesis which is known to cease following phage infection.

c. Nucleoside Triphosphate : D N A Nucleotidyltransferase. An enzyme from calf thymus nuclei catalyzes the addition of a single 5’-nucleotidyl group from any one of the ribonucleoside triphosphates (GTP, CTP, ATP, and UTP) to the 3’-hydroxyl group of the terminal deoxyribonucleotide residue of primer DNA, Eq. (34). DNA

-

+ X T P Mga+ DNA-(XMP) + PPi

(34) Although ribonucleotidyl-DNA exhibits primer activity for ATP and UTP polymerases (184-188) and for the polymerization of ribonucleotides by polynucleotide phosphorylase (180), these compounds have not been demonstrated to occur in nature. The physiological significance of reaction (34) is therefore in doubt.

B. ADENYLCYCLASE In the foregoing examples of phosphodiester synthesis, an adenylyl group from ATP is transferred to the 3‘-hydroxyl group of a nucleotide residue a t the terminus of HNA or an oligonucleotide. A somewhat similar, but unique, example of adenylylation is seen in the conversion of ATP to 3’,5’-CAMP (189). In this case a single molecule of ATP serves both as the adenylyl donor and the adenylyl acceptor. As in nucleic acid metabolism, the adenylyl group of ATP is transferred from ATP to a 3‘-hydroxyl group to form a phosphodiester and inorganic pyrophosphate, Eq. (35). The importance of the reaction in the regulation of metabolism has been amply demonstrated (190).Adenylyl cyclase is ubiquitous and has been isolated from a number of sources. 183. P. J. Ortiz, J. T. August, M. Watanabe, A. M. Kaye, and J. Hurwitz, JBC 237, 3786 (1962). 184. J. S. Krakow, H. 0. Kammen, and E. S. Canellakis, BBA 53, 52 (1961). 185. E. S. Canellakis and Z. N. Canellakis, in “Informational Macromdecules” (H. J. Vogel, V. Bryson, and J. 0. Lampen, eds.), p. 107. Academic Press, New York,

1963. 186. H. G. Klemperer, J. S. Krakow, and E. S. Canellakis, BBA 61, 43 (1962). 187. H. G. Klemperer and H. 0. Kammen, BBRC 6, 344 (1962). 188. M. E. Gottesman and E. S. Canellakis, JBC 241, 4339 (1966). 189. E. W. Sutherland and T. W.Rall, JBC 232, 1065 (1958). 190. T. W. Rall, M. Rodbell, and P. Condliff, eds., “The Role of Adenyl Cyclase nnd 3’,5’-AMP in Biological Systems.” Fogarty International Proceedings No. 4 ( 1969).

1. ADENYLYL

27

TRANSFER REACTIONS 0

0

I

I

I1 II CH,-0-P-O-P-O-P-O-

Adenine

G HO

9

0-

0 II I

9

on

(35)

Since various polynucleotides are cleaved by phosphorolysis and pyrophosphorolysis to yield nucleoside diphosphates and triphosphates, it is evident that the 3’,5‘-phosphodiester linkage possesses a high group transfer potential (191).This has been confirmed by the demonstration that the cyclase from Brevibacterium liquefaciens (192,193) catalyzes pyrophosphorolysis of cAMP to form ATP, i.e., the reverse of reaction (35). At pH 7.3,pMg 3-3.5, and a t 25”, the observed equilibrium constant for reaction (35) is Kobs

=

(CAMP)(PPi) = o.Mr, M (ATP)

the corresponding AG:,, is +1.6 kcal mole-’. From this value and the observed standard free energy of hydrolysis of ATP to yield 5’-AMP and PPi (AG:,, = -10.3 kcal mole-’), the observed standard free energy of hydrolysis of cAMP to yield 5’-AMP was calculated to be -11.9 kcal mole-‘ (192).Thus, the 3’-phosphodiester bond of cAMP is about 3 kcal mole-1 more “energy rich” than the P,y-pyrophosphate bond of ATP (free energy of hydrolysis, AG:,, = 8.8 kcal mole-’) under similar conditions (192).

C. ADENYLYLATION OF AMINOGLYCOSIDE ANTIBIOTICS During the last two decades numerous drug resistance factors (R factors) have been found in strains of Enterobacteriaceae th a t confer upon these organisms a capacity to grow normally in the presence of various 191. F. Liprnann, Adwan. Enzyme Regul. 9, 5 (1971). 192. 0. Hayaishi, P. Greengard, and S. Colowick, JBC 246, 5840 (1971). 193. K. Taksi, Y. Kurashina, C. Suzuki, H. Okarnoto, A. Uedi, and 0. Hayaishi, JBC 246, 5843 (1971).

28

E. R. STADTMAN

drugs and antibiotics (194). Genetic studies established that these R factors are extrachromosomal elements that replicate independently of the host chromosome and can be transmitted to essentially all members of the Enterobacteriaceae either by bacterial conjugation or transduction (194). Among these R factors are those that specify the elaboration of enzymes catalyzing the adenylylation of specific hydroxyl groups on the amino glycoside antibiotics. At least two different adenylyltransferases have been detected in appropriate R+strains. Both are constitutive periplasmic enzymes. One of these is the ATP: streptomycin- (spectinomycin) adenylyltrans-

p-ccNH

HA,

C

gNH

WLC+NH

(-+-c:::z I

I

NH

NH,

HO

HO

OH OH

HOQo

CH,NH f F

H

+ ATP

CHO

H

OH

-

O

q

o

OH

+ PPi

CHO

(36)

(Fj CH,NH

j

no

P

O=P-0-Ad

Streptomycin

I

0Adenylyl-streptomycin

7

H&-NH H O

O

F

-0-P-O-Ali H,C-NH H 0 3 x C H 3

Ha

H HO NCH,

+ ATP

-

+ PPf

NC H,

HO

o

HO

o

Spectinomycin

0

0

Adenylyl- spectinomycin

FIQ.2. Adenylylation of streptomycin and spectinomycin. 194. J. E. Davies and

R. Rownd, Science 176, 758 (1972).

(37)

1. ADEINYLYL

29

TRANSFER REACTIONS

ferase that catalyzes inactivation of streptomycin by adenylylation of the 3’-OH group on the L-glucosamine residue (195) of this antibiotic, Eq. (36) in Fig. 2. The same enzyme also catalyzes inactivation of spectinomycin, probably by attachment of an adenylyl group to the hydroxyl group of the D-threo-methylamino alcohol moiety (194-198), Eq. (37) in Fig. 2.

R =

ATP

& t

-

R

/

/O

CH,OH

NH* OH

HO

I

O=P -0-Ad

iPPi

I

+ ATP-

H&NH

OH

0Adenylyl-kanamycin A

Kanamycin A

HO &OH

w

0

/ OI

h

H,CNH O H

-0-P-0-Ad

II

Centamicin

Adenylyl-gentamycin

FIG.3. Adenylylation of kanamycin A and gentamycin. The arrows marked “a,” “b,” and “c” indicate sites of acetylation, phosphorylation, and adenylylation, respectively. 195. T. Yamada, D. Tipper, and J. Davies, Nature (London) 219, 288 (1968). 196. R. Benveniste, T.Yamada, and J. Davies, Infection Immunity 1, 109 (1970). 197. H.Umesawa, S. Takasawa, M. Okanishi, and R. Utahara, J. Antibiot., Ser. A 21, 81 (1968). 198. D. H. Smith, J. A. Janjigian, N. Prescott, and P. W. Anderson, Infection Immunity 1, 120 (1970).

30

E. R. S'I'ADTMAN

All spectinomycin- and streptomycin-resistant strains that have been examined contain the adenylyltransferase that is specific for these antibiotics (194). Adenylylation is therefore the only known mechanism for the inactivation of streptomycin or spectinomycin. A different enzyme catalyzes the adenylylation of gentamicin, and is probably also responsible for adenylylation of kanamycin (194, 199). It is proposed that a hydroxyl on the garosamine rings (see Fig. 3) is the most probable site of adenylylation (199) ; however, the hydroxyl group on the deoxystreptomycin ring is also a candidate for the group that undergoes modification. In contrast to streptomycin, or spectinomycin, which can only be inactivated by adenylylation (194), gentamicins and kanamycins may also be inactivated by either acetylation or phosphorylation. It is noteworthy that different R factors arc responsible for the elaboration of enzymes catalyzing these reactions (194); moreover, the sites of esterification are different from the adenylylation site (see Fig. 3 ) (194). Neither the streptomycin nor the gentamycin-adenylyltransferase has been purified to homogeneity ; however, preliminary results based on gel filtration studies with partially purified preparations indicate that they have molecular weights of about 30,000 and 12,500, respectively. Both activities are stimulated by NH,' (and amine buffers) and are subject to CAMP mediated catabolite repression (200).

IV. Synthesis of Adenosine Diphosphate Derivatives

A. GENERALFEATURES Transfer of an adenylyl group from ATP to the phosphoryl group of monophosphate esters [Eq. (38)] is an important mechanism for the synthesis of various coenzymes (Table IV) . 0

0

ATP

+ R-0-

B-0-

A-

I1

-+

R-0-P-0-P-0-Ad

0

II

A- A-

+ PPi

(38)

The first reaction of this type, which is also the first example of an adenylyl transfer to be recognized, was discovered in yeast extracts by 199. R. Benveniste and J. Davies, FEBS Lett. 14, 293 (1971). 200. W. Shaw, personal communication.

c3

s

TABLE I V REACTIONS O F ATP

Eq. No.

WITH PHOSPHATE

ESTERSTO

P

Enzyme

Reaction

(40)

ATP: NMM-adenylyltransferae

Nicotinate ribonucleotide

(41)

ATP :FMN-adenylyl transferase

FMN

(42)

ATP: Pantetheine-P adenylyltransferase

(43)

ATP: glucos+l-P adenylyltransferase

(44p

ATP:myonic acid denylyltransferase

+ ATP

0

ATP :2,3-diphosphoglycerate adenylyltrmsferase

FAD

~

+ ATP

Ref. deamido-DPN

+ PPi

+ ATP dephospho-Coh + PPi ~ - ~ - G l ~ c o ~ e+ - l ATP - P 5 ADPa-D-glucose + PPi Myonic acid + ATP + adenine myonic acid Pantetheine-4'-P

dinucleotide (45)a

2 2 s

ADENOSINE L)IPHOSPHATF:L)ERIV*4TIVES

PHODUCE

+ PPi

+

2,3-Diphosphoglycerate ATP + adenylyl2,3diphosphoglycerate PPi

+

+ PPi

m

12, 202,

b

203

z

206-207

2

208 209-21 2 218 219

These reactions have not been demonstrated; they are inferred from the structure of the isolated adenosine diphosphate derivative.

W

c

32

E. R. STADTMAN

Kornberg (12).The enzyme catalyzes the reversible synthesis of D P N by the reaction: Nicotinamide mononucleotide + ATP DPN + PPi (39) Subsequent studies showed that under physiological conditions nicotinic acid mononucleotide rather than nicotinamide mononucleotide [Eq. (40) in Table IV] is probably the normal substrate for the enzyme (201-204). I n the meantime analogous reactions with F M N and pantetheine-4‘-P [Eqs. (41) and (42) in Table IV] were shown to be involved in the synthesis of FAD (205-207) and CoA (208), respectively. The properties of these enzyme systems were summarized earlier by Kornberg (71)and by Imsande and Handler in an earlier volume (203) of “The Enzymes”; hence, they will not be discussed further here.

B. SYNTHESIS OF ADPGLUCOSE The reaction between ATP and glucose l-P to yield ADPglucose [Eq. (43) in Table IV] was discovered in wheat extracts by Espada (209) and later found in extracts of an Arthrobacter sp. by Shen and Preiss (210).Highly purified preparations of the enzyme have been obtained from plant (209, 211) and bacterial sources (210,212). The enzyme from both sources exhibits an absolute requirement for Mg2+ and a relatively high specificity for ATP as the nucleotidyl donor. No other nucleoside triphosphate will replace ATP as a substrate for the enzyme from corn grain (211), whereas with the bacterial enzyme the rates of synthesis of CDPglucose, IDPglucose, UDPglucose, and ADPglucose are 6.5, 0.6, 20, 0.1, and 0.5%, respectively, of that of ADPglucose (210, 212). The assumption that ADPglucose formation involves transfer of the adenylyl group from ATP is based on analogy to a similar reaction in which UDP-glucose is formed, Eq. (46). 201. J. Preiss and P. Handler, JBC 233, 493 (1958). 202. J. Preiss and P. Handler, JBC 233, 488 (1958). 203. J. Imsande and P. Handler, “The Enzymes,” 2nd ed.. Vol. 5. p. 281, 1961. 204. J. Preiss and P. Handler, JACS 79, 4246 (1957). 205. A. W. Schrecker and A. Kornberg, JBC 182, 795 (1950).. 206. C. DeLuca and N. 0. Kaplan, BBA 30, 6 (1958). 207. C. DeLuca, “Methods in Enzymology,” Vol. 6, p. 342, 1963. 208. M. B. Hoagland and G. D. Novelli, JBC 207, 767 (1954). 209. J. Espada, JBC 237, 3577 (1962). 210. L. Shen and J. Preiss, JBC 240, 2334 (1965). 211. J. Espada, “Methods in Enzymology,” Vol. 8, p. 259, 1966. 212. L. Shen and J. Preiss, “Methods in Enzymology,” Vol. 8, p. 262, 1966.

1. ADENYLYL

33

TRANSFER REACTIONS

UTP

+ glucose-1-P -+

UDPglucose

+ PPi

(46)

It is evident that this reaction involves transfer of a uridylyl group from UTP to glucose 1-P since, in the presence of 32P-labeled PPi, isotope is incorporated into U T P (213). Whereas UDPglucose is the immediate source of glucosyl groups for the synthesis of glycogen by animals, ADPglucose is the immediate source of glucosyl groups for the synthesis of starch in plants (214, 215) and of bacterial glycogen (215217). Equation (43) (Table IV) therefore represents the first committed step in the biosynthesis of starch and glycogen in plants, green algae, and bacteria. Thus, it is not surprising that the ATP:glucose-1-P adenylyltransferases from these organisms exhibit both positive and negative control by various metabolites. The regulatory characteristics of the ATP: glucose-1-P adenylyltransferases from different sources have been summarized by Preiss (217). It is evident from these effects that in bacteria and plants accumulation of glycogen and starch is favored under conditions of ATP “excess.” These polysaccharides can therefore be regarded as storage forms of ATP energy. Preiss (217) has noted that the ATP: glucose-1-P adenylyltransferases can be classified into six groups according to their activator and inhibitor specificities and furthermore that there is a good correlation between the major pathway of carbohydrate metabolism of the organism and the nature of the primary activator of its ATP: glucose-1-P adenylyltransferase (Table V). C. ADENINE-MYONIC ACID DINUCLEOTIDE AND ADENYLYL DIPHOSPHOGLYCERATE In addition to the above compounds whose biological functions are well established, two other adenine dinucleotide derivatives of unknown function have been isolated from animal sources, but the enzymes catalyzing their formation have not been purified and the mechanism of their synthesis has not been definitely established. Adenine-myonic acid dinucleotide was isolated by de Caputto et al. (218) from rabbit 213. A. Munch-Peterson, H. M. Kalckar, and E. E. B. Smith, Kgl. Dan. Vidensk. Selsk., Bid. M e d d . 22, No. 73 (1955); quoted by H. M. Kalckar, in A d v a ~ . Enzymol. 20, 111 (1958). 214. E. Recondo and L. F. Leloir, BBRC 6, 85 (1961). 215. V. Ginsburg, Advan. Enzymol. 26, 35 (1964). 216. L. Shen, H. P. Ghosh, E. Greenberg, and J. Preiss, BBA 89, 370 (1964). 217. J. Preiss, Cum. Top. Cell. Regul. 1, 125 (1969). 218. D. P. de Caputto, W. H. Mosley, J. L. Poyer, and R. de Caputto, JBC 236, 2727 (1961).

TABLE V AND INHIBITORS OF ADPGLUCOSE PYROPHOSPHORYLASES FROM VARIOUS SOURCES ACTIVATORS Primary activator

Source ~~

~~

Serratia marcescena Aeromnas formicum

Inhibitor

Possible mode of carbon metabolism

~

Leaves of higher plants green algae SPhosphoglycerate Escherichia coli, Aerobacter aerogenes, Aerobacter cloacae, Salmonella typhimurium, Citrobacter freundii, Escherichia aurescens Arthrobacter viscosus Agrobacterium tumefaciena Rhodopseudomonas capsulata Rhoohpirillurn rubrum

Secondary activators

Fructose-di-P, TPNH, pyridoxal-P,

Fructose-6-P Pyruvate None Fructose-6-P, fructose-1,Mi-P

Fructos&P, fructose-di-P, phosphoenolpyruvate ZPhosphoglycerate, Sphosphoglyceraldehyde, phosphoenolpyruvate Pyruvate ltibose-5-P Deoxyribose-5-P None ?

Pi 5'-AMP

Calvin cycle or Hatch slack cycle Glycolysis

Pi, AMP, ADP None None 5'-AMP ADP

Does not grow on glucose; grows well on TCA intermediates Glycolysis? Glycolysis?

PJ p

E Ez

1.

35

ADENYLYL TRANSFER REACTIONS

muscle as a crystalline solid and has been tentatively assigned structure (I). I

H-C-OH

H-C-OH I

I I

I

H2C

I

0

I

I1

0-P-0-P-0 I l 0I

I

CH2

0

I

II

l

9

Myonic acid

(I)

On the basis of this structure and by analogy t o Eqs. (40)-(43) in Table I V it seems likely that adenine myonic dinucleotide is produced by reaction of ATP with myonic acid, Eq. (44) in Table IV. A compound corresponding in composition to adenylyl diphosphoglycerate was isolated from pig blood (219). Treatment of the compound with nucleotide pyrophosphatase from snake venom of Aglcistrodon blomhofi led to the formation of 5’-AMP and 2,3-diphosphoglyceric acid. Thus, the adenylyl group is presumed to be attached to pyrophosphate linkage to one of the phosphoryl groups of diphosphoglycerate. One could imagine that it is synthesized by the reaction shown in Eq. (45), Table IV.

V. Sulfate Activation

An eneynie (ATP: sulfate adenylyl transferase) that catalyzes the synthesis of adenosine-5’-phosphosulfate (APS), Eq. (47), has been 0 ATP

+ SO:- + H+

It

-+

-0-+O-

!

0

1-0-Ad + PPi

A-

(47)

found in numerous organisms (220-227). The enzyme has been partially purified from rat liver (228), yeast (228-230), and Penicillium chryso219. T. Hashimoto and H. Yashikawa, BBRC 5, 71 (1961).

36

E. R. STADTMAN

genum (231, 232); the molecular weights of the enzyme from these sources are 900,000, 100,000, and 440,000, respectively. The observed equilibrium constant for Eq. (47) a t pH 8.0 is lo-@(616).Thus, the reaction is so strongly endergonic (AG’ a t pH 8.0 = +11 kcal mole-l) that under physiological conditions only negligible amounts of adenylyl sulfate can accumulate. However, as pointed out by Robbins and Lipmann (229), the reaction can be pulled in the direction of adenylyl sulfate synthesis by hydrolysis of the product, PPi, which is catalyzed by pyrophosphatase, Eq. (9). Since the AG& for Eq. (9) a t p H 8.0 and pMg 4 is about -8.5 kcal mole-’ (4), the AGE,, for the coupled reaction (48) [i.e., Eq. (9) plus Eq. (47)] is 11-8.5 or +2.5 kcal mole-’ ATP

+ SO’,- + H20 ----+ APS + 2 Pi

(48)

Therefore, in the activation of sulfate by ATP, the advantages of an adenylylation versus a phosphorylation mechanism are obvious. Not only is the adenylyl group transfer potential somewhat higher than the phosphoryl group transfer potential of ATP (see Table I) but also hydrolysis of the pyrophosphate bond serves as an extra source of energy to drive the activation reaction. These thermodynamic considerations may account a t least in part for the fact that whereas ADP cannot replace ATP in the synthesis of adenylyl sulfate [Eq. (49)], phosphorolysis of APS t o form A D P and SO:- [reverse of Eq. (49)] is nevertheless catalyzed by the activating enzyme (228, 233). The apADP

+ 80:- ----+ APS + Pi

(49) parently higher adenylyl transfer potential of ATP compared to ADP under physiological conditions is largely the result of differences in the

220. F. Lipmann, Science 128, 575 (1958). 221. P. C. DeVito and J. Dreyfuss, J . Bacteriol. 88, 1341 (1964). 222. A. S. Levi and G . Wolf, BBA 178, 262 (1969). 223. J. F. Wheldrake, BJ 105, 697 (1967). 224. M. C. Jones-Mortimer, J. F. Wheldrake, and C. A. Pasternak, BJ 107, 51 ( 1958). 225. M. C. Jones-Mortimer, BJ 110, 589 (1968). 226. J. Dreyfuss and A. B. Pardee, J . Bacteriol. 91, 2275 (1966). 227. C. A. Adams and R. E. Johnson, Plant Physiol. 43, 2041 (1968). 228. P . W. Robbins, “Methods in Enzymology,” Vol. 5, p. 964, 1962. 229. P. W. Robbins and F. Lipmann, JBC 233, 686 (1958). 230. L. G. Wilson and R. S. Bandurski, ABB 62, 503 (1956). 231. J. W. Tweedle and I. H. Segel, JBC 246, 2438 (1971). 232. J. W. Tweedle and I. H. Segel, Prep. Biochem. 1, 90 (1971). 233. P. W. Robbins, “The Enzymes,” 2nd ed., Vol. 6, p. 469, 1962.

1.

37

ADENYLYL TRANSFER REACTIONS

relative affinities of divalent cations for the substrates and products of the reaction. Thus, in reaction 47, the affinity of Mgz+ for the product, PPi, is considerably greater than for the substrate, ATP, whereas in reaction (49) the reverse is true; the affinity of Mg2+ for the substrate ADP is greater than its affinity for the product, Pi ( 4 ) . Inasmuch as the formation of adenylyl sulfate is the first step in the metabolism of sulfate, it is not surprising that its formation is under metabolic control. In yeast and in P. chrysogenum synthesis of ATP:sulfate adenylyltransferase is repressed by methionine, and activity of the enzyme is inhibited by sulfide (234). I n E . coli the level of adenylyltransferase is repressed by cystine (236). VI. Synthesis of lmidol Adenylate Derivatives

Transfer of an adenylyl group from ATP to an imidol oxygen atom is apparently involved in the synthesis of both argininosuccinate (236, 237) and guanylic acid (238, 239). As in acyl-CoA and aminoacyl tRNA synthesis, the formation of these metabolites probably occurs by a twostep mechanism. The first step leads to an enzyme-bound imidol adenylate and PPi, Eq. (501, and the second involves replacement of the adenylyl group with an amino compound to form the metabolite and AMP, Eq. (51). Therefore, in the overall reaction the energy of the a,P-pyrophosphate bond of ATP is used to facilitate replacement of the imidol oxygen atom with an amino derivative, Eq. (52). NH

E

+ R-NH-

NH ER-NH-

A*

-OH

NH

+ ATP s E.R-NH-b-&-!’-O-Ad

A-

A-

+ R’-NHt

II

0

+E

(51)

+ Ad-0- 4-6- + PPi

(52)

A-

NH

A*

-OH

(50)

+ R-NH--CNHR’

+ Ad-0-P-6-ll Sum: R-NH-

+ PPi

NH

0

4-0-P-0-Ad .1

0

NH

+ ATP + R’-NH,

-+ R-NH-

e

-NHR’ 0

A-

38

E. R. STADTMAN

A. ARGININOSUCCINATE SYNTHESIS The formation of an imidol adenylate intermediate in the synthesis of argininosuccinate was inferred by results of experiments with lSOlabeled citrulline showing that in the course of the overall reaction, the imidol oxygen atom [indicated by 6 in Eqs. (50)-(52)] becomes incorporated into the phosphoryl group of the AMP that is produced. However, the proposed mechanism was contradicted by the fact that argininosuccinate synthetase does not catalyze the expected exchange of inorganic pyrophosphate into ATP in the presence of citrulline, Eq. (50) ; moreover, the inability to catalyze this exchange could not be attributed to an unfavorable equilibrium situation since ATP-PPi exchange does occur when both citrulline and aspartate are present (937). I n view of these results it was suggested that the synthesis of argininosuccinate might involve a concerted reaction between ATP, citrulline, and aspartate without the formation of adenylylcitrulline as a distinct intermediate (237). I n subsequent studies, however, Rochovansky and Ratner (936) showed that when substrate levels of highly purified argininosuccinate synthetase are incubated with ATP and citrulline, both adenylyl-citrulline and PPi are produced but they remain tightly bound to the enzyme (936). The failure of ATP to exchange with PPi is therefore the result of the fact that PPi does not dissociate from the enzyme to equilibrate with added "YP-labeledPPi. They showed further that whereas a-methyl aspartate cannot replace aspartate in the overall biosynthetic reaction, it does replace aspartate in stimulating the citrulline-dependent ATP-PPi exchange reaction (236). These results indicate that the stimulation of exchange by aspartate is not due to reversal of the overall biosynthetic reaction but rather to its ability to facilitate dissociation of PPi from the enzyme. Accordingly, the synthesis of argininosuccinate probably involves the sequence of reactions shown in Scheme 1. In addition to its role as an acceptor for the citrulline moiety in step c of this scheme, aspartate by binding to the enzyme apparently also facilitates dissociation of PP i from the intermediate ternary complex (step b ) .

234. 235. 236. 237. 238. 239.

P. C. DeVito and J. Dreyfuss, J . Bncleriol. 88, 1341 (1964). C. A. Pasternak, BJ 85, 44 (1962). 0. Rochovansky and S. Ratner, JBC 242, 3839 (1967). 0. Rochovansky and S. Rntner, JBC 236, 2254 (1961). R. Abrams and M. Bentley, ABB 79, 91 (1959). U. Lngerkvist, JBC 233, 143 (1958).

1.

39

ADENYLYL TRANSFER REACTIONS

-0-Ad 1 I

NH

0-

I

COOH COOH I

H-C-NH, COOH I

HC-NH-C AMP

+

LOOH FH'

B{

NH

FK

I1

(;H2i. NH

+

NH I1

I

,J:;( H-C-NH,

HC-NH, COOH

0 II

C-0-P-0-Ad I

'-

COOH

PPi

SCHEME 1

The studies of Rochovansky and Ratner are extremely important because they demonstrate that inability to observe partial reactions by exchange experiments does not preclude the existence of a sequential two-step mechanism. Thus, the failure of enzymes to catalyze detectable ATP-PPi exchange, except in the presence of all reactants, as in the case of some aminoacyl tRNA synthetases (33) and in the case of GMP synthetase (238, 239) does not exclude the formation of an adenylylated intermediate in these reactions (33, 37, 238).

B. SYNTHESIS OF G M P The synthesis of GMP from xanthylic acid, ATP, and either NH,+ or glutamine is assumed to involve a two-step mechanism (Scheme 2) analogous to that for the synthesis of arginosuccinate. The G M P synthetases from pigeon liver (239, 240) and from bone marrow (238) utilize glutamine as the preferred amino donor, whereas the enzyme from Aerobacter aerogenes (241) utilizes ammonia as the sole source of amino nitrogen. The two-step mechanism is supported by the fact that in the overall reaction the imidol oxygen of X M P is incorporated, 240. U. Lagervist, JBC 233, 138 (1958). 241. H. S. Moyed and B. Magasanik, JBC 228, 339 (1957).

40

no

E. R. STADTMAN

&)

+ ATP

-

Ad-0-P-0 I 0-

LRibose- 5‘- P Xanthylic acid

N LRitmse- 5 P f -

Adenylyl-xanthylic acid A ,../

J

Gerogenes) glutamine (liver)

?H Glutamate (liver)

+

AMP

+ H2N LRibose- 5’-P Guanosine-5‘-P

SCHEME 2

without dilution, into the phosphoryl group of AMP (238, 239). However, as with the arginosuccinate system described above, efforts to demonstrate partial reactions by pyrophosphate exchange were unsuccessful (238,239, @ l ) , as were also efforts to demonstrate overall reversibility of the reaction (239, 241). Nevertheless, as in the case of arginosuccinate synthesis, the inability to demonstrate partial reactions may result from the failure of PPi to dissociate from the enzyme; the conclusion that a concerted reaction is involved (258) is therefore probably not justified. VII. Adenylylation of Functional Groups of Proteins

A. ADENYLYLATION AND DEADENYLYLATION OF E. coli GLUTAMINESYNTHETASE Adenylylation and deadenylylation of glutamine synthetase are ultimate steps in a highly complicated casc,ade system (242) that regulates the activity of glutamine synthetase in E. coli. This system will be described in greater detail later in a separate chapter devoted to the regulation of glutamine synthetase activity (Vol. IX of “The Enzymes”). The present discussion is therefore concerned only with the nature of the adenylylation reactions themselves; a discussion of the role of these 242. A. Ginsburg and E. Stadtman, in “The Enzymes of Glutamine Metabolism” (S. Prusiner and E. R. Stadtman, eds.), p. 9. Academic Press, New York, 1973.

1.

41

ADENYLYL TRANSFER REACTIONS

reactions in cellular regulation will be included in the forthcoming chapter. Suffice it to mention here that the adenylylation of glutamine synthetase leads to marked changes in a number of the enzyme’s characteristics, among which are pH optimum, divalent ion specificity, stability, catalytic potential, and susceptibility to feedback inhibition by various end products of glutamine metabolism (243-245). In general, adenylylation leads essentially to inactivation of the enzyme under most physiological conditions (246). Adenylylation of glutamine synthetase (GS) is catalyzed by a specific adenylyltransferase (ATase) and involves transfer of one adenylyl group from ATP to a specific acceptor site on each subunit of the enzyme (247-249). Since glutamine synthetase contains twelve identicsl subunits, up t o twelve equivalents of adenylyl groups can be bound to each enzyme molecule, Eq. (53) (247). AG:b (kcal mole-’)

12 ATP

M@+

+ GS +GS-(AMP)ig + 12 PPi ATm

+ GS-(AMP)ig W GS + 12 ADP Sum: 12 ATP + 12 Pi -+ 12 ADP + 12 PPi 12 Pi

-1.0

(53)

-1.0

(54)

(55) Following exhaustive proteolysis of adenylylated enzyme, Shapiro and Stadtman (650) isolated in good yield a single adenylylated decapeptide having the approximate composition Asp, Glu, Pros Gly, Leul Tyr,. Proof that the adenylyl group is attached in phosphodiester linkage to the tyrosyl residue was obtained by showing that treatment of the peptide with snake venom phosphodiesterase resulted in the release of 5’-AMP and the appearance of an equivalent amount of unesterified tyrosyl hydroxyl groups (250). In the meantime, Heinrikson and Kingdon (251, 252) established the amino acid sequence of a 21 amino acid tryptic peptide containing the adenylylated tyrosyl residue. Starting from the amino terminal end of this peptide the amino acid sequence is -2.0

243. E. R. Stadtman, A. Ginsburg, J. E. Ciardi, J. Yeh, S. B. Hennig, and B. M. Shapiro, Aduan. Enzyme Regul. 8, 99 (1970). 244. B. M. Shapiro and E. R. Stadtman, Annu. Rev. Microbiol. 24, 501 (1970). 245. E. R. Stadtman, Harvey Lect. 65, 97 (1971). 246. D. Mecke, K. Wulff, and H. Holzer, BBA 128, 559 (1966). 247. H. S. Kingdon, B. M. Shapiro, and E. R. Stadtman, Proc. Nat. Acad. Sci. U. S. 58, 1703 (1967). 248. B. M. Shapiro, H. S. Kingdon, and E. R. Stadtman, Proc. Nat. Acad. Sci. U.S. 58, 612 (1967). 249. K. Wulff, D. Mecke, and H. Holzer, BBRC 28, 740 (1967). 250. B. M. Shapiro and E. R. Stadtman, JBC 243, 3769 (1968). 251. R. L. Heinrikson and H. S. Kingdon, JBC 245, 138 (1970). 252. R. I,. Heinrikson and H. S. Kingdon, JBC 246, 1099 (1971).

42

E. R. STADTMAN

AMP

1

Ile-His-Pro-Gly-GluAla-Met-Lys-Asp-Asn-Leu- yrAsp-Leu-Pro-Pro-GluGly-Glu-Ala-Lys

It is noteworthy that each subunit of glutamine synthetase contains 15 tyrosyl groups (248) but only one of these can be adenylylated. Heinrich et al. (253) identified inorganic pyrophosphate as a product of the adenylylation reaction, and Mantel and Holzer (254) showed that the reaction is reversible. The equilibrium varies significantly with pH and divalent ion concentration. At pH 7.35 and 30°, the observed

equilibrium constant varies from 2.0 to 30 as the Mgz+ concentration is varied from 3 to 50 mM. When the Mgz+ concentration is held constant, pMg = 2.0, the values for Kobsa t pH 6.6, 7.36, and 7.74 are 2.4, 8.5, and 23.9, respectively. The interpolated Kobsa t pH 7.0 and pMg 2 is 5.7, which corresponds to a AGL. = 1.0 kcal mole-l. From this and the fact that at pH 7.0, pMg 2.0, the AGEbsfor hydrolysis of ATP to yield AMP and PPi is about 10 kcal mole-1 (Table I ) , it is estimated from reactions (56) and (57) that the AGzb8 for hydrolysis of the AMP-Otyrosyl bond of adenylylated glutamine synthetase [ i.e., reverse of Eq. (58)] is about -9.0 kcal mole-l.

+

ATP GS PP, AMP

+

+

G S A M P PP, ATP H20

+

AGZb. (kcal mole-') -1.0

+lO.O

(56) (57)

____

+ GS

+ H20

(58) The adenylyl-0-tyrosine residue in glutamine synthetase therefore has a high group transfer potential comparable to that of ATP. Whereas deadenylylation of glutamine synthetase by the reverse of Eq. (53) is thermodynamically feasible, this is probably not an important physiological mechanism. Instead, deadenylylation is catalyzed by a more complex enzyme system (255, 256) involving phosphorolysis of the adenylyl-0-tyrosyl bond to form ADP and unadenylylated enzyme, Eq. (54). From the chemical point of view, Eq. (54) and the reverse of Eq. (53) are formally very similar. Both are adenylyl transfer reactions in Sum: AMP

GS-AMP

+9.0

253. C. P. Heinrich, F. A. Battig, M. Mantel, and H. Holzer, Arch. Mikrobiol. 73, 104 (1970). 254. M. Mantel and H. Holzer, Proc. N a t . Acad. Sci. U.8.e5, 660 (1970). 255. B. M. Shapiro, Biochemistry 8, 659 (1969). 256. W. B. Anderson and E. R. Stadtman, BBRC 41, 604 (1970).

1.

ADENYLYL TRANSFER REACTIONS

43

which the adenylyl moiety of the enzyme-bound adenylyl-0-tyrosyl group is transferred to a phosphoric acid group to form a phosphodiester derivative. In Eq. (53) inorganic pyrophosphate is the adenylyl group acceptor, whereas in Eq. (54) orthophosphate is the acceptor. I n view of this chemical similarity it is not surprising that both reactions are catalyzed by one and the same adenylyltransferase (25‘7). It follows therefore that the separate functions of this ATase must be rigorously regulated to prevent useless coupling between reactions (53) and (54) which would lead simply to phosphorolysis of A T P to form ADP and PPi with concomitant slight loss of energy [Eq. (55)]. Indeed, a highly sophisticated mechanism has evolved which permits rigorous regulation of the adenylylation and deadenylylation reactidns (242, 255-260), but a detailed discussion of this regulatory system is deferred to a later volume of “The Enzymes.” Suffice it to say that regulation of the two functions is mediated by a regulatory protein, PI, (255, 257, 261, 262, 263), whose capacity to specify adenylylation or deadenylylation activity of ATase is regulated by its interconversion between uridylylated and unmodified forms (259-263). This interconversion in turn is catalyzed by two separate enzymes (259) whose activities are under strict metabolic control by UTP, ATP, glutamine, a-ketoglutarate, NH,, glutamate, and various glycolytic and Krebs cycle intermediates (246, 255-264). The ATase has been isolated as a homogeneous protein (264-2681, but determined values for its molecular weight have varied from 115,000 (265) to 130,000 (266) to 145,000 (264). Hennig et al. (266) showed 257. B. M. Shapiro and E. R. Stadtman, BBRC 30, 32 (1968). 258. W. B. Anderson, S. B. Hennig, A. Ginsburg, and E. R. Stadtman, Proc. N a t . Acad. Sci. U . S. 67, 1417 (1970). 259. J. H. Mangum, G. Magni, and E. R. Stadtman, unpublished data. 260. E. R. Stadtman, M. Brown, A. Segal, W. A. Anderson, S. B. Hennig, A. Gins-

burg, and J. H. Mangrum, in “Proceedings of the Second International Symposium on Metabolic Interconversion of Enzymes” (0.Wieland, E. Helmreich and H. Holzer, eds.), p. 231. Springer-Verlag, Berlin and New York, 1972. 261. W. B. Anderson and E. R. Stadtman, A B B 143, 428 (1971). 262. E. R. Stadtman, A. Ginsburg, W. B. Anderson, A. Segal, M. S. Brown, and J. E. Ciardi, in “Molecular Basis of Biological Activity” (K. Gaede, B. L. Horecker, and W. J. Whelan, eds.), p. 127. Academic Press, New York, 1972. 263. M. Brown, A. Segal, and E. R. Stadtman, Proc. N a t . Acad. Sci. U . S. 88, 2949 (1971). 264. E. Ebner, D. Wolf, C. Gancedo, S. Elsasser, and H. Holzer, Eur. J. Biochem. 14, 535 (1970). 265. D. Wolf, E. Ebner, and H. Hinze, Eur. J. Biochem. 25, 239 (1972). 266. S. B. Hennig. W. A, Anderson, and A. Ginsburg, Proc. Nut. Acad. Sci. U . S. 87, 1761 (1970).

44

E. R. STADTMAN

that an apparently homogeneous preparation of 130,000molecular weight could be dissociated into two dissimilar subunits of 60,000 and 70,000 molecular weights. Dissociation was accompanied by a complete loss of ability to catalyae deadenylylation of glutamine synthetase [Eq. (54)] and a slight increase in capacity to catalyze adenylylation [Eq. (53)1. The adenylylation capacity resided exclusively in the 70,000 molecular weight subunit (266, 267). The latter subunit form has been isolated as a homogeneous protein (267). The influence that the 60,000 molecular weight subunit might have a regulatory role concerned with the deadenylylation function of ATase remains to be determined. The possibility that ATase is an adenylyl group carrier in the transfer of the adenylyl group from ATP to glutamine synthetase was suggested by the observation that some ATase preparations catalyze glutamine-stimulated exchange of PPi into ATP in the absence of glutamine synthetase (258, 286, 268). However, the exchange capacity is lost with purification of the enzyme by some procedures (269) and upon dissociation of the 130,000molecular weight aggregated form (258, 266, 267; moreover, kinetic studies indicate that the adenylylation of glutamine synthetase involves an ordered mechanism in which a ternary complex between ATase, ATP, and glutamine synthetase is an obligatory intermediate (269). These results are not consistent with a pingpong mechanism in which an AMP-adenylyltransferase complex is an intermediate. Nevertheless, as clearly shown by Rochovansky and Ratner ($36) in their studies with argininosuccinate synthetase, absence of PPi-ATP exchange does not necessarily exclude the formation of an adenylylated enzyme intermediate. Exchangeability by different enzyme preparations could reflect variations in protein conformation that affect the ease with which PPi can dissociate from the postulated AMPadenylyltransferase intermediate, and the apparent requirement for an interaction with glutamine synthetase could be related to its ability to facilitate dissociation of PPi from the ATase rather than to its involvement in a concerted reaction.

B. ADENYLYLATION OF OF E . coli

THE

LYSINE-SENSITIVE ASPARTOKINASE

During growth on glucose minimal media, activity of the lysinesensitive aspartokinase (AK 111) of E. coli increases rapidly during 267. S. B. Hennig and A. Ginsburg, ABB 144, 611 (1971). 268. M. D. Denton and A. Ginsburg, Fed. Proe. Fed. Amer. SOC. Ezp. Biol. 27, 783 (1968). 269. R. M. Wohlhueter, E. Ehner, and D. Wolf, JBC 247, 4213 (1972).

1. ADENYLTL

TRANSFER REACTIONS

45

exponential growth, reaches a maximum in early stationary phase when glucose is depleted, and then rapidly declines (270). This suggests that the level of AK 111 is under metabolic control other than by simple repression and derepression. Furthermore, AK I11 isolated from stationary cultures of E . coli has abnormally high absorbancy in the region of 260 nm. Compared to the enzyme isolated from cells in midexponential growth (970) this spectral difference is similar to that exhibited between adenylylated and unadenylylated forms of glutamine synthetase (271). It was therefore suggested that adenylylation of the AK 111 might occur in the stationary phase of batch cultures (270). I n support of this suggestion, Niles and Westhead (270) showed that incubation of purified aspartate kinase with either [ P ~ ~ P ] A T Por [3H]ATP and a relatively crude soluble fraction of E . coli extract led to the covalent attachment of the labeled moiety of ATP to the aspartate kinase. Snake venom phosphodiesterase released the labeled material from the enzyme indicating that the derivative (probably an adenylyl group) is bound in phosphodiester or phosphoramidate linkage to the enzyme. Unfortunately, under the in vitro conditions used, only a small amount of protein-bound ATP derivative was produced (0.03-0.05 equivalent per mole of enzyme). This has precluded positive identification of the covalently bound group as an adenylyl group as well as identification of the specific site of its attachment. Nevertheless, these preliminary findings invite much speculation on the possible role of adenylylation reactions in the regulation of aspartate kinase activity in E. coli.

C. ADENYLYLTRANSFER FUNCTIONS OF DNA LIGASE DNA ligases, the so-called “joining enzymes,” catalyze repair of single-strand breaks in native DNA. This repair involves joining together in phosphodiester linkage the exposed 5’-phosphoryl group and the 3‘-hydroxyl group of the nicked DNA duplex. It is therefore an endergonic process and is driven by energy made available from cleavage of the energy-rich adenosine pyrophosphate bonds in either ATP or D P N (272-276). As illustrated in Fig. 4, the pyrophosphate 270. E. G. Niles and E. W. Westhead, Biochemistry (1972) (in press). 271. B. M. Shapiro and A. Ginsburg, Biochemistry 7 , 2153 (1968). 272. C. C. Richardson, Y. Masamume, T. R. Live, A. JacqueminSablon, B. Weiss, and G . C. Fareed, Cold Spring Harbor Symp. Quant. Biol. 33, 151 (1968). 273. C. C. Richardson, Annu. Rev. Biochem. 38, 795 (1969). 274. B. M. Olivera, Z. Q. Hall, Y. Anraku, J. R. Chien, and I. R. Lehman, Cold Spring Harbor Symp. Quant. Bwl. 33, 27 (1968).

46

E. R. STADTMAN

bond energy is made available by transfer of an adenylyl group from ATP or D P N to a specific lysyl residue on the DNA ligase, thus forming an enzyme-bound energy-rich adenosine monophosphoramidate derivative (277) [reaction (I),Fig. 41. The adenylyl group is then transferred from phosphoramidate linkage to the 5'-phosphoryl group a t the broken end of the DNA strand, thereby producing an adenine dinucleotide derivative of DNA (272-276) [reaction (11), Fig. 41. Finally, the newly formed pyrophosphate bond of this new dinucleotide is cleaved by attack of the 3'-hydroxyl group of the nicked DNA, displacing the adenylic acid moiety and regenerating the phosphodiester bond [reaction (III), Fig. 41. In essence the DNA ligase is a bifunctional enzyme. [The r-amino group of one of its lysyl residues serves as a carrier for transport of energy-rich adenylyl groups from either ATP or D P N to the 5'-phos-

((b) DPN

+ N'-Ligase

s

2 Ad-0-P-N'-Llgase I

+ NMN

(Cata'yzed native enzyme)

0

3 L -7

i

FIQ.4. Reactions involved in the repair of single-strand breaks in n DNA duplex.

275. M. Gellert, J. W. Little, C. K. Oshinsky, and S. Zimmerman, Cold S p h i g Harbor Symp. Quant. Biol. 33, 21 (1968). 276. P. Sadowski, B. Ginsberg, A. Yudelevich, 1,. Feiner, and J. Hunvitz, Cold Spring Harbor Symp. Quant. Biol. 33, 165 (1968). 277. R. I. Gumport and I. R. Lehman, Proc. Nat. Acad. Sci. U.S. 88, 2559 (1971).

1.

ADENYLYL TRANSFER REACTIONS

47

phoryl group of broken DNA strands, but in addition i t catalyzes a specific reaction between the 3’-hydroxyl group of nicked DNA and the adenine dinucleotide derivative at the broken 5’-phosphate terminus to form the phosphodiester bond and AMP (278-285).] It is noteworthy that D P N is the only compound that can serve as an adenylyl donor for the DNA ligase from normal E . coli cells (274, 275, 286, 287), whereas ATP is a specific adenylyl donor for the phageinduced DNA ligase in E . coli (272, 276, 288, 289, 291) as well as for the enzyme from mammalian sources (290).As predicted by the equations in Fig. 4, the isolated adenylylated ligase from phage-infected E. coli yields ATP in the presence of Mg2+and added PPi [reverse of reaction (Ia) ] (27.9, 681); similarly, the isolated adenylylated DNA ligase derivative from uninfected cells forms D P N in the presence of N M P [reverse of reaction (Ib) ] (274, 275, 292). Furthermore, the adenylylated derivative of both kinds of ligase forms AMP in the presence of nicked DNA, reactions (11) and (111) (274, 278-280, 292). Following incubation with their respective adenylyl group donors (DPN or ATP), adenylylated derivatives of both forms of DNA ligase have been isolated by gel filtration (272, 281, 288). Gumport and Lehman (277) showed that proteolytic degradation of both adenylylated forms of ligases leads to the release of an adenosine 5‘-monophosphoramidate derivative of lysine. This indicates that the adenylylation of these enzymes by ATP or D P N involves covalent attachment of an 278. Z. W. Hall and I. R. Lehman, JBC 244, 43 (1969). 279. C. I,. Harvey, T. F. Gabriel, E. M. Wilt, and C. C. Richardson, JBC 248, 4523 (1971). 280. B. M. Olivera, Z. W. Hall, and I. R. Lehman, Proc. Nut. Acud. Sci. U. S. 81, 237 (1968). 281. B. Weiss, A. Thompson, and C. C. Richardson, JBC 243, 4556 (1968). 282. W. Seifert, D. Rabussay, and W. Zillig, FEBS L e t t . 18, 175 (1971). 283. D. Rabussay, R. Mailhammer, and W. Zillig, in “Proceedings of The Second

International Symposium on Metabolic Interconversidn of Enzymes” (0. Wieland, E. Helmreich and H. Holzer, eds.), p. 213. Springer-Verlag, Berlin and New York, 1972. 284. C. G . Goff and K. Weber, personal communication. 285. T. Honjo. Y. Nishizuka, 0. Hayaishi, and I. Kato, JBC 243, 2553 (1968). 286. S. B. Zimmerman, J. W. Little, C. K. Oshinsky, and M. Gellert, Proc. Nut. Acad. Sci. U. S 57, 1841 (1967). 287. B. M. Olivera and I. R. Lehman, Proc. N a t . Acud. Sci. U.S. 57, 1700 (1967). 288. B. Weiss and C. C. Richardson, Proc. Nut. Acad. Sci. U.S. 57, 1021 (1967). . S. 289. A. Becker. G. Lyn, M. Gefter, and J. Hurwitz, Proc. N a t . Acad. S C ~U. 58, 1996 (1967). 290. T. Lindahl and G. M. Edelman, Proc. Nut. Acud. Sci. U.S. 81, 680 (1968). 291. B. W. Weiss and C. C. Richardson, JBC 242, 4270 (1967). 292. J. W. Little, S. B. Zimmerman, C. K. Oshinsky, and M. Gellert, Proc. Nut. Acad. Sn’. U. S. 58, 2004 (1967).

48

E. R. STADTMAN

adenylyl group to an N‘-lysyl residue on the enzyme. [Reactions (Ia) and (Ib) in Fig. 4.1 Whereas these are the only examples to date in which adenylylation of an NC-lysyl residue of an enzyme occurs, there was an earlier preliminary report (293) that cell-free extracts of Mycobacterium avium contain an enzyme catalyzing the formation of free adenosine 5’-monophosphoramidate by reaction of ATP with ammonia Eq. (59). 0 ATP

+ NHI

-+

H,N-

h-0-Ad + PPi d-

(59)

It was reported, further, that the AMP-NH, derivative is an intermediate in the synthesis of amino acids from the corresponding a-ketoacid precursors ( 2 9 4 ) .However, the enzymes used in these investigations were relatively crude preparations and since alternative explanations for the experimental results were not excluded, validity of the conclusions remains in doubt.

D. COVALENT MODIFICATION OF RNA POLYMERASE 1 . DPN-Dependent Covalent Modification of the

(Y

Subunit

Soon after infection of E . coli with bacteriophage T,, the a subunit of RNA polymerase undergoes a chemical modification which was first recognized by its increase in negative charge, as judged by polyacrylamide gel electrophoresis (295, 296). In the meantime it was demonstrated that the chemical modification involves covalent attachment of 5’-adenylic acid or a derivative thereof to the a subunit (282, 283, 297). Treatment of the altered enzyme with snake venom phosphodiesterase leads to the release of 5’-AMP (297). However, the altered enzyme contains one adenine group and two phosphoryl groups (284, 297). Therefore, the alteration may not involve an adenylyl group transfer reaction. Perhaps the modification involves adenosine diphosphate ribosylation of the a subunit as was shown earlier to be involved in the diphtheria toxin-dependent modification N. Katunuma, ABB 78, 547 (1958). N. Ellfolk and N. Katunuma, ABB 81, 521 (1959): G. Walter, W. Seifert, and W. Zillig, BBRC 30, 240 (1968). W. Seifert. P. Qosba, G. Walter, P. Pnlm, M. Schachner, and W. Zillig, Eur. J . Biochem. 9, 319 (1969). 297. C. G. Goff and K. Weber. Cold Spring Hoiboi S y m p . Quont. Biol. 35, 101 293. 294. 295. 296.

(1970).

1.

49

ADENYLYL TRANSFER REACTIONS

of aminoacyl transferase (285). Such a possibility is consistent with the recent discovery that in the in uitro alteration of the subunit DPN rather than ATP is the source of the covalently bound 5'-AMP derivative (284). (Y

2. Possible Artifact

It has been reported that native E. coli RNA polymerase exists in two interconvertible forms: an active, unadenylylated form and an inactive adenylylated form. In a preliminary communication (298) it was shown that in the presence of Mg2+ and relatively impure enzyme preparations adenylyl groups from ATP become attached to RNA polymerase and that this is accompanied by inactivation of the enzyme. Concomitant release of the bound adenylyl groups and reactivation of the enzyme was obtained in the presence of a second enzyme preparation. I n the meantime, workers in several other laboratories have confirmed these same observations but have concluded that the observed inactivation of RNA polymerase is not the result of adenylylation per se but of the nonspecific binding of polyadenylylic acid derivatives that are produced by the combined action of ATPase and RNA phosphorylase, both of which are present in the inactivating enzyme systems. Also, reactivation of the inactivated RNA polymerase is attributed to the hydrolysis of the inhibitory adenylic acid polymer by the action of nucleases that are present in the reactivating enzyme preparation (299).

298. C. A. Chelala, L. Hirschbein, and H. N. Torres, Proc. Nut. Acad. Sci. U . S. 68, 152 (1971). 299. K. Nath and J. Hiirwitz, personal communication.

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Uridine Diphosphory1 Glucose Pyrophos9horylase RICHARD L . TURNQUIST

R . GAURTH HANSEN

I . Introduction . . . . . . . . . A . Measurement of Activity . . . . B . Purification . . . . . . . C . Analytical and Synthetic Applications I1. Metabolic Function . . . . . . . A . Cytology . . . . . . . B . Metabolism . . . . . . . C . Metabolic Regulation . . . . I11. Properties . . . . . . . . . A . Optima . . . . . . . . B . Structure . . . . . . . C . Kinetics . . . . . . . D . Specificity . . . . . . . E . Mechanism . . . . . . .

. . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . . .

. . . . . . . . . . . . . . .

. . . . . . . . . . . . . . .

51 52 53 54 55 55 57 59 62 62 62 65 68 69

.

I Introduction

Uridine diphosphoryl glucose pyrophosphorylase (UTP:a-wglucose-lphosphate uridylyltransferase. EC 2.7.7.9) catalyzes the formation of nucleoside diphosphate sugars from nucleoside triphosphates and sugar1-phosphates [Eq. ( 1 ) ] . NTP

+ sugar-1-phosphate 2 NDP-sugar + PP. 51

(1)

52

RICHARD L. TURNQUIST AND R. GAURTH HANSEK

The enzyme appears to be ubiquitous in nature, and, since its discovery (I, d ) , it has been detected and purified from a variety of sources (3-9). Its universal occurrence is not surprising since a major product of the reaction that it catalyzes, UDPglucose, has a central role as a glucosyl donor in numerous cellular transformations (10). A. MEASUREMENT OF ACTIVITY Several different assay procedures have been developed to detect and quantify UDPglucose pyrophosphorylase. The rate of reaction (1) can be determined in either direction using a variety of substrates. I n the direction of pyrophosphorolysis of the nucleoside diphosphate sugar when the sugar is glucose, the quantity of glucose 1-P produced can be determined using coupling enzymes which convert it to glucose 6-P and 6-phosphogluconate (11). In addition to the appropriate enzymes, phosphoglucomutase and glucose 6-phosphate dehydrogenase, the assay mixture contains NADP', pyrophosphate, and the nucleoside diphosphate glucose. The reduction of NADP' is followed a t 340 nm. If the nucleoside diphosphate sugar contains uridine, guanine, adenine, or inosine, the nucleoside triphosphate formed in the reaction may be used to phosphorylate 3-P-glyceric acid, which can be reduced by NADH (12 ) . Glyceraldehyde-3-P dehydrogenase and 3-P-glycerate kinase are used as coupling enzymes, and the oxidation of NADH is followed spectrophotometrically. 1. H. M. Kalckar and E. Cutolo, Proc. Znt. Congr. Biochem., 2nd, 1962 p. 260 (1953). 2. A. Munch-Petersen, H. M. Kalckar, E. Cutolo, and E. E. B. Smith, Nature (London) 172, 1036 (1953). 3. E. E. B. Smith, G . T. Mills, and E. M. Harper, J. Gen. Microbiol. 16, 426 ( 1957). 4. E. F. Neufeld, V. Ginsburg, E. Putman, E. W. Fanshier, and W. Z. Hassid, ABB 69, 602 (1957). 5. C. Villar-Palasi and J. Larner, BBA 30, 449 (1958). 6. J. H. Pazur and E. W. Shuey, Fed. Proc., Fed. Amer. Soc. Ezp. Biol. 20, 216 (1961). 7. M. Axelos and C. Peaud-Lenoil, Bull. SOC.Chirn. Biol. 51, 261 (1969). 8. P. N. Viswanathan, Zndiun J. Biochem. 6, 124 (1969). 9. I. J. Russell and D. R. Lineback, Carbohyd. Res. 15, 123 (1970). 10. V. Ginsburg, Advan. Enzymol. 26, 35 (1964). 11. A. Munch-Petersen, Acta Chem. Scand. 9, 1523 (1955). 12. H. Verachtert, S. T. Bass, L. L. Seifert, and R. G. Hansen, Anrtl. Biochern. 13, 259 (1965).

2.

URIDISE DIPHOSPHORTL GLUCOSE PTROPHOSPHORYLASE

53

The synthesis of UDPglucose may be quantified with UDPglucose dehydrogenase (IS), which oxidizes the nucleoside sugar to UDPglucuronate and concomitantly reduces the NAD'. Two moles of NAD' are reduced for every mole of UDPglucose formed, increasing the sensitivity of the assay. A more time consuming but also more sensitive assay for UDPglucose formation involves incubating the enzyme with U T P and radioactive sugar-1-phosphates. After the reaction is stopped, the radioactive nucleoside diphosphate sugars are adsorbed on charcoal, then eluted, and the radioactivity measured (14, 16). B. PURIFICATION Uridine diphosphoryl glucose pyrophosphorylase was successfully crystallized from calf (16), human, (17),lamb, goat, and rabbit (18) livers. The procedure utilized an alkaline extraction of homogenized liver followed by protamine sulfate treatment. The enzyme was precipitated from the supernate fraction with ammonium sulfate, and, after dialysis, was treated with calcium phosphate gel and adsorbed on a DEAEcellulose column. After elution, the enzyme was again concentrated with ammonium sulfate, dissolved, and crystallized from ammonium sulfate solution. Various procedures have been used successfully in partially purifying the enzyme from other sources. Ginsburg (19) purified the enzyme from mung bean acetone powder using ammonium sulfate fractionation, alumina C y treatment and cellulose chromatography. Bovine mammary pyrophosphorylase was purified from acetone powder using essentially the same procedure as used for human liver ( 2 0 ) . Tsuboi et al. (21) purified human erythrocyte enzyme using calcium phosphate gel, ammonium sulfate, cold ethanol, DEAE-cellulose and Sephadex G-200 fractionations. Although the specific activity of the enzyme was quite 13. J. 1,. Strominger, H. M. Kalckar, J. Axelrod, and E. S. Maxwell, JACS 76, 6411 (1954). 14. D. M. Carlson and R. G. Hansen, JBC 237, 1260 (1962). 15. A. Munch-Petersen, Acta Chem. Scand. 11, 1079 (1957). 16. G. J. Albrecht, S. T. Bass, L. L. Seifert, and R. G. Hansen, JBC 241, 2968 (1966). 17. J. Knop and R. G. Hansen, JBC 245, 2499 (1970). 18. J. Knop, Master's Thesis, Michigan State University, 1969. 19. V. Ginsburg, JBC 232, 55 (1958). 20. V. S. Steelman and K. E. Ebner, BBA 128, 92 (1966). 21. K. K. Tsuboi, K. Fukunaga, and J. C. Petricciani, JBC 244, 1008 (1969).

54

RICHARD L. TURNQUIST AND R. GAURTH H A N S E N

high, crystallization was not achieved. Franke and Sussman (22) purified the enzyme from the slime mold, Dictyostelium discoideum, to apparent physical and immunochemical homogeneity by ammonium sulfate fractionation followed by elution through Sephadex G-100, Hypatite C, and polyacrylamide gel. Using various combinations of these procedures, the enzyme has been purified from human brain ( 2 3 ) , rat mammary tissue (241, rabbit muscle ( 2 5 ) , maize (26), slime mold (27), bacteria (28-30),and yeast ( 3 1 ) .

C. ANALYTICALAND SYNTHETIC APPLICATIONS Several synthetic and analytical techniques have been designed which utilize UDPglucose pyrophosphorylase and the reaction it catalyzes. The enzyme has been used to effect the synthesis of ["C]UDPglucose, using labeled glucose, glucose 6-P or glucose-fructose mixtures as starting materials ( 3 2 . 3 4 ) .The enzymic synthesis of UDPglucosamine has also been described (35) which probably utilizes UDPglucose pyrophosphorylase. The enzyme was also used to determine PPi with an application to tissue extracts (36).Whenever PPi is produced in a reaction, the potential is there to use the pyrophosphorylase as an analytical coupling reagent; for example, the procedure was used to determine RNA polym22. J. Franke and M. Sussman, JBC 248, 6381 (1971). 23. D.K. Basu and B. K. Buchhawat, J . Neurochem. 7, 174 (1961). 24. R. S. Emery and R. L. Baldwin, BBA 1 3 , 223 (1967). 25. C. Villar-Palasi and J. Larner, ABB 86, 61 (1960). 26. J. D.Vidra and J. D. Loerch, BBA 159, 551 (1968). 27. G. Gustafson and B. E. Wright, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 30, 1069 (1971). 28. A. Kamogawa and K. Kurahashi, J . Biochem. (Tokyo) 57, 758 (1965). 29. T. Chojnacki, T. Sawicka, and T. Korzybski, Acta Bbchim. Pol. 15, 293 (1968). 30. H. Nikaido and T. Nakae, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 29, A598 (1970). 31. A. Munch-Petersen and H. M. Kalckar, "Methods in Enzymology," Vol. 2, p. 675, 1955. 32. E. R. Trucco, Nature (London) 174, 1103 (1954). 33. L. Glaser, JBC 232, 627 (1958). 34. A. Wright and P. W. Robbins, BBA 104, 594 (1965). 35. F. Malay, G. F. Malay, and H. A. Lardy, JACS 78, 5303 (1956). 36. J. C. Johnson, M. Shanoff, 8. T. Bass, J. A. Boeai, and R. C. Hansen, Anal. Biochem. 26, 137 (1968).

2.

URIDIKE DIPHOSPHORTL GLUCOSE PYROPHOSPHORYLASE

55

erase activity in biological materials. With radioactive tracers the method can be made sensitive to as little as 0.2 pmoles of PPi (37). II. Metabolic Function

A. CYTOLOGY While UDPglucose pyrophosphorylase is present in nearly all tissues, it is usually most abundant in those that display active polysaccharide synthesis. The enzyme may account for 0.2-0.3% of the extractable protein of calf liver (16), and as much as 1% of the protein in slime mold cells ( 3 8 ) .The amount of enzyme in tissue may also vary greatly with the age and physiological state of the organism. These factors probably determine the ultimate success of enzyme purification procedures. In higher animals, liver most often has the highest concentration of UDPglucose pyrophosphorylase. Skeletal muscle, heart, and kidney have intermediate enzyme levels, while spleen, lung, brain, testis, and fatty tissues have relatively low activity levels (39, 40). In most animal tissues, the pyrophosphorylase activity generally corresponds to the glycogenic activity or glycogen content of the cells. In sheep (41), chickens (.do), rats (@), pigeons, and humans ( 4 3 ) , both pyrophosphorylase activity and glycogenolysis rise during fetal life, reaching a peak near birth or hatching. Activity then falls as much as 60% as the animal ages. Silkworm ovary tissue shows high pyrophosphorylase activity near the middle of the pupal stage, which corresponds to rapid enzymic conversion of blood trehalose to glycogen (44). In tumor tissue, which characteristically has high glycolytic activity and decreased glycogen content, the pyrophosphorylase activity values are as much as 50-60% less than those of normal tissues (46). Brain tissue is the exception to the rule in that it has a low specific activity of UDPglucose pyrophosphorylase but a high glycogen content (39). 37. H. Flodgaard, Eur. J . Biochem. 15, 273 (1970). 38. P. Newel1 and M. Sussman, J M B 49, 627 (1970). 39. C. Villar-Palasi and J. Lamer, ABB 86, 270 (1960). 40. M. T. Rinaudo, C. Giunta, M. L. Boazi, and R. Bruno, Enzymologia 36, 321 (1969). 41. F. J. Ballard and I. T. Oliver, BJ 95, 191 (1965). 42. F. J. Ballard and I. T. Oliver, BBA 71, 578 (1963). 43. K. J. Isselbacher, Science 126, 652 (1957). 44. 0. Yamashita, J . Sericult. SOC.Jup. 38, 329 (1969). 45. V. N. Nigam, H. L. MacDonald, and A. Cantero, Cancer Res. 22, 131 (1962).

56

RICHARD L. TURNQUIST AND R. GAURTH HANSEN

While most of the UDPglucose pyrophosphorylase in animal cells is found in solution in the cytoplasm (25, 44, @), about 5-10% is bound to the microsomal fraction. It has therefore been concluded that glucose 1-P is constantly being recycled into glycogen and that this mechanism may help regulate glycogen storage ( 4 7 ) . In higher plants, enzyme level variations are usually associated with a change in capacity for starch and sucrose synthesis ( 4 8 4 2 ) . In lower plants, the enzymic activity may vary with cellulose or trehalose formation (bS, 6 4 ) . In plant cells, too, UDPglucose pyrophosphorylase has been found fully (65) or mostly (56, 57) dissolved in the cytoplasm. The enzyme may be especially concentrated in chloroplasts and starch granules (8, 68, 5 9 ) . Pyrophosphorylase activity may be markedly changed by physiological manipulation. Sugar beets synthesize increased amounts of the enzyme when pyrocatechol or vanadyl sulfate is applied to the foliage (60, 61). Tri-iodothyronine will significantly increase the pyrophosphorylase activity in hypothyroid rat muscle (62) while 1311 will lower enzyme levels in the same tissue. Riboflavin deficiency will also lower pyrophosphorylase levels in rats (6s).Meal fed rats have higher enzymic activities in muscle (50%) and adipose (300%) tissues than do nibbling rats (64, 6 5 ) . 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59.

E. Reid, BBA 32, 251

(1959).

V. T. Maddaiah and N. B. Madsen, Can. J . Biochem. 46, 521 (1968). J. F. Turner, Awt. J . Biol. Sci. 22, 1321 (1969). J. F. Turner, A w t . J . Biol. Sci. 22, 1145 (1969). R. Pressey, Plant Physiol. 44, 759 (1969). C. Y. Tsai, F. Salamini, and 0. E. Nelson, Plnnt Physiol. 46, 299 (1970). K. C. Tovey and R.M. Roberts, P h n t Soil 48, 406 (1970). B. E. Wright and M. L. Anderson, BBA 31, 310 (1959). K. Zetsche, 2. Naturjorsch. B 23, 369 (1968). A. E. S. Gussin and J. H.McCormnck, Phytochemistry 9, 1915 (1970). M. A. Hall and L. Ordin, Plant Physiol. 40, Suppl. XXXVIII (1965). M. A. Hall and L. Ordin, Physiol. Plant 20, 624 (1967). I. F. Bird, H. K. Porter, and C. R. Stocking, BBA 100, 366 (1965). W. A. Huber, M. A. R. de Fekete, and H. Zieglcr, Planta 87, 360 (1969). 60.B. Singh and D. J. Wort, Plant Physiol. 44, 1321 (1969). 61. B. Singh and D. J. Wort, Physiol. Plnnt 23, 920 (1970). 62. C. Pitra, E. G . Krause, and A. Wollenberger, Endokrinologie 54, 225 (1969). 63. H. B. Burch, 0. H. Lowry, M. E. Bradley, and P. F. Max, Jr., Anzei. J . Phyaiol. 219, 409 (1970). 64. J. H. Wiley and G . A. Leveille. Fed. Proc., F e d . Amer. SOC.Exp. B i d . 28, 625 (1969).

65. J. H. Wiley and G . A. Leveille, J . N u t i . 100, 85 (1970).

2.

DRIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

57

B. METABOLISM A prime function of UDPglucose pyrophosphorylase in most animal cells is to activate glucosyl residues for the synthesis of glycogen. Although glycogen synthesis is obviously not solely dependent upon the activity of pyrophosphorylase, changes in pyrophosphorylase activity may affect glycogen levels ( 6 6 ) . Pathological conditions that foster abnormally high concentrations of pyrophosphorylase may result in increased levels of stored glycogen ( 6 7 ) . Since UDPglucose inhibits glycogen phosphorylase, the increase in glycogen content probably results from a combination of enhanced synthesis and decreased catabolism (68). The role of UDPglucose pyrophosphorylase in the synthesis of cellulose in higher plants has been the subject of considerable controversy. While UDPglucose is the principal sugar nucleotide involved in the production of most plant polysaccharides, GDPglucose has generally been considered to be the glucosyl donor in cellulose production (69). Evidence now indicates, however, that in some plants, cellulose is produced either totally or in part from UDPglucose (57, 70-73). In lower plants and microorganisms cellulose is synthesized with UDPglucose as the glucosyl donor (33, 7 4 ) , but the identity of the ultimate donor in higher plants awaits further clarification. A similar controversy exists in relation to the biosynthesis of starch. Both UDPglucose and ADPglucose have been implicated as the glucosyl donor (8, 75-77). While differences exist between plants and even within the same plant (78) both nucleotides and their respective pyrophosphorylases are probably involved. Sucrose, which is the primary source for starch production, is converted by sucrose synthetase to UDPglucose 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78.

R. Kornfield and D. H. Brown, JBC 238, 1604 (1963). G. Okuno, S. Hisukuri, and M . Nishikawa, Nature (London) 212, 1490 (1966). N. B. Madsen, BBRC 6, 310 (1961). W. Z. Hassid, Science 165, 137 (1969). D. 0. Briimmond and A. P. Gibbons, Biochem. 2. 342, 308 (1965). L. Ordin and M. A. Hall, Plant Physwl. 42, 205 (1967). L. Ordin and M. A. Hall, Plant Physiol. 43, 473 (1968). G. Franz, Phytochemistry 8, 737 (1969). C. Ward and B. E . Wright, Biochemistry 4, 2021 (1965). L. F. Leloir, M. A. R. de Fekete, and C. E. Cardini, JBC 236, 636 (1961). L. F. Leloir, BJ 91, 1 (1964). T. Murata, T. Sugiyama, and T. Akasawa, A B B 107, 92 (1964). Y. Tanaka and T. Akazawa, Plant Cell Physiol. 9, 405 (1968).

58

RICHARD L. TURNQUIST AND R. GAURTH HANSEN

and fructose. The UDPglucose is converted by pyrophosphorylase to glucose 1-P, and then to ADP glucose, which seems to be the immediate glucosyl donor for starch formation (48,49, 79,80). Since in some plants there is evidence that UDPglucose is the only nucleotide involved in starch synthesis, two different mechanisms may be operating. I n maize endosperm (61), sucrose is hydrolyzed by invertase to glucose and fructose. The glucose is converted to glucose 6-P and then to glucose 1-P which in turn is converted by UDPglucose pyrophosphorylase to UDPglucose. In this endosperm, the UDPglucose can be used directly in the synthesis of starch. In other plants, sucrose may be converted by sucrose synthetase to UDPglucose which is then converted to glucose 1-P by pyrophosphorylase as in normal starch production. Here, however, the glucose 1-P is converted directly into starch by starch phosphorylase without first being incorporated into ADPglucose (60, 81). Both of these latter pathways occur in young plants and seem to be temporary until ADPglucose pyrophosphorylase is present. Uridine diphosphoryl glucose is also the glucosyl donor for sucrose production which utilizes essentially the reverse pathway of starch synthesis (82, 83).The glucose 1-P resulting from starch phosphorolysis is coupled with fructose through intermediate formation of UDPglucose via the pyrophosphorylase pathway (84, 86). Uridine diphosphoryl glucose pyrophosphorylase is also necessary for the metabolism of galactose. In the normal galactose metabolism pathway, the enzyme galactose 1-P uridylyltransferase (EC 2.7.7.12) utilizes UDPglucose to convert galactose 1-P to UDPgalactose. However, the (a)

Galactose

(b) Gal-1-P

+ ATP

+ UDP-Glc

kinase

transferase

+ ADP TJDP-Gal + Glr-1-P Gal-1-P

-

(2)

epimersse

(r)

TJDP-Gal

UDP-Glr

pyrophosphorylase pathway has the capacity to synthesize UDPgalactose even in the absence of the transferase (86). Without UDPglucose 79. M. A. R . de Fekete and C. E. Cardini, A B B 104, 173 (1964). 80. T. Murata, T. Sugiyama, T. Minamikawa, and T. Akanawa, ABB 113, 34 ( 1966). 81. M. A. R. de Fekete, Planta 87, 311 (1969). 82. C. E. Cardini, L. F. Leloir, and J. Chiriboga, JBC 214, 149 (1955). 83. M. A. R . de Fekete, Planta 87, 324 (1969). 84. D. P. Burma and D. C. Mortimer, ABB 62, 16 (1956). 85. M. D. Hatch, J. A. Sacher, and K. T. Glasaiou, Plant Physiol. 38, 338 (1963). 86. T. Sawicka m d T. Chojnachi, Clin. Chim. Acta 23, 463 (1969).

2.

URIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

+ ATP kinaae Gal-1-P + ADP pyrophoephorylase Gal-1-P + UTP , ’ UDP-Gal + PPi

59

(a) Galact,ose (b) (c)

UDP-Gal

.

epimerase

’ UDP-Glc

pyrophoephorylase

(d) UDP-Glc

+ PPi ,

(3)

’G l e l - P

+ UTP

pyrophosphorylase, galactose will not be incorporated into microbial cell walls (87-89). Uridine diphosphoryl glucose pyrophosphorylase participates in the synthesis of numerous other compounds including various cell wall po1y.mers in both higher plants (90-92) and microorganisms (88,93), trehalose (94), glycosides (96),glycolipids (96), heparin (97), microbial antigens (98), lactose (20), glucuronides (99, IOO), and rhamnose (87, 101).

C. METABOLIC REGULATION Since UDPglucose participates in numerous metabolic pathways, the enzyme that catalyzes its synthesis may not be subject to extensive metabolic control. Experimental results tend to bear this out. While UDPglucose pyrophosphorylase is subject to product inhibition like many other enzymes, additional controls on either its synthesis or the reaction it catalyzes are minimal. I n plants, the administration of indole acetic acid (auxin) may cause 87. T. A. Sundararajan, A. Rapin, and H. M. Kalckar, Proc. Nut. Acad. Sci. U . S. 48, 2187 (1962). 88. T. Fukasawa, K. Jokura, and K. Kurahashi, BBRC 7, 121 (1962). 89. T. Fukasawa, K. Jokura, and K. Kurahashi, BBA 74, 608 (1963). 90. D. S. Feingold, E. F. Neufeld, and W. Z. Hassid, JBC 233, 783 (1958). 91. S. H. Goldemberg and L. R. Marechal, BBA 71, 743 (1963). 92. 0. A. Pavlinova and M. F. Prasblova, Fiziol Rast. 17, 295 (1970). 93. M. Lieberman, C. Buchanan, and A. Markovitz, Proc. Nut. Acad. Sci. U. S.

65, 625 (1970). 94. R. Roth and M. Sussman, JBC 243, 5081 (1968). 95. G. Franz and H. Meier, Planta Med. 17, 396 (1969). 96. J. A. Curtino, R. 0. Calderon, and R. Caputto, Fed. Proc., Fed. Amer. SOC. E z p Biol. 27, 346 (1968). 97. I. Danishefsky and 0. Heritier-Watkins, BBA 139, 349 (1957). 98. R. L. Bernstein and P. W. Robbins, JBC 240, 391 (1965). 99. M. Shikamnra and K. K. Tsuboi, Amer. J . Dis. Child. 102, 600 (1961). 100. R. M. Roberts and K. M. K.Rao, Fed. Proc., Fed. Amer. SOC.Exp. Bid. 30, 1117 (1971). 101. G. A. Barber, ABB 103, 276 (1963).

60

RICHARD L. TURNQUIST AND R. GAURTH HANSEN

a large increase in the rate of glucose incorporation into cell wall polysaccharides, but the effect seems to be on enzymes other than UDPglucose pyrophosphorylase (102, 103). Stimulation of the autonomic nervous system, either sympathetic or parasympathetic, will affect glycogen metabolism in the mammalian liver, but it is glycogen synthetase which is affected and not UDPglucose pyrophosphorylase (104, 106). Insulin was tested in mammals with contradictory results. The hormone was reported to have no effect on pyrophosphorylase levels (106), but the lack of it, caused by alloxan treatment, decreased UDPglucose pyrophosphorylase activity in rat salivary glands by as much as 70% (107).It was speculated that insulin might exert a regulatory effect on uridylyltransferases. Generally, however, the control on UDPglucose pyrophosphorylase activity is genetic, regulating the amount of enzyme synthesized. Most of the genetic studies were carried out in microorganisms, especially Escherichia coli, and the location of the UDPglucose pyrophosphorylase structural gene was precisely determined (93, 108). The gene is not located in the gal operon (89, 109). Uridine diphosphoryl glucose pyrophosphorylase activity increases dramatically but unequally during cap formation in Acetabularia. Enzymic activity appears to be concentrated in the apical cells and decreases basally. The gradient is not the result of activators or inhibitors but rather of actual variations in the total amount of enzyme produced by differential synthesis. This is true even in anucleate cells, indicating that the synthesis may be regulated in the cytoplasm a t the level of translation of long-lived messenger RNA (641. The unequal distribution of the pyrophosphorylase may be the result of a messenger RNA gradient toward the apex of the stalk, implying a migration of the RNA from the nucleus in the rhizoid (110). Investigations into the metabolic control of UDPglucose pyrophosphorylase in the slime mold, Dictyostelium discoideum, have generated considerable controversy. As the organism differentiates to the plasmodium stage, large amounts of protein disappear, carbohydrate synthesis 102.. A. Abdul-Baki and P. M. Ray, Plant Physiol. 42, Suppl., S-4 (1967). 103. A. Abdul-Baki and P. M. Ray, Plant Physiol. 47, 537 (1971). 104. T. Shimazu and T. Fujimoto, BBA 252, 18 (1971). 105. T. Shimazu, BBA 252, 28 (1971). 106. C. Villar-Palasi and J. Larner, ABB 94, 436 (1961). 107. T. Szymczek, B. Swiatkowska, and M. Jachimowicz, Acta Biochim. Pol. 18, 177 (1971). 108. J. A. Shapiro, J . Bacteriol. 92, 518 (1966). 109. F. Jacob and J. Monod, J M B 3, 318 (1961). 110. K. Zetsche, Planta 89, 244 (1969).

2.

URIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

61

increases, and there is an apparent increase in the specific activity of UDPglucose pyrophosphorylase (53,111). In the 18-hr maturation process, the specific activity of the enzyme was reported to increase as much as tenfold (112, 113). A large increase in enzymic activity resulting in elevated carbohydrate metabolism was reported (74, 114). Serious questions were subsequently raised as to whether increased pyrophosphorylase activity exerts an effect on carbohydrate metabolism or whether the pyrophosphorylase activity increases a t all. Experiments indicated that the specific activity of the enzyme increases only slightly (11 5 ) , and reports of large increases measured in vitro resulted from artifacts such as enzyme instability and the effects of different substrate concentrations (116, 117). Kinetic considerations indicate that even if the pyrophosphorylase levels did increase, this alone could not cause the observed increase in UDPglucose synthesis (118), which may be simply the result of increases in the levels of UTP and glucose 1-P which are substrates for the pyrophosphorylase (119). By contrast it has been argued that the increases in enzyme levels are real and not resulting from differential stability. The kinetic models are somewhat suspect in that substrate molecules do not occur randomly throughout the cytoplasm but are compartmentalized and concentrated (1go). Cycloheximide and actinomycin D prevent the enzyme increase, showing that it is dependent upon new protein synthesis (112). Uridine diphosphorylglucose pyrophosphorylase accumulation, controlled at the level of genetic transcription, is linked with and necessary for the morphological changes that occur during differentiation (38, 121). Enzyme concentrations, however, are rarely the limiting factors in metabolic reactions; substrate availability usually determines reaction rates. Computer kinetic models of slime mold differentiation have indicated that changes in the pyrophosphorylase concentration over a wide range would have little or no effect on UDPglucose synthesis. If enzyme is not limiting, the reaction will 111. J. M. Ashworth, BJ 106, 28p (1968). 112. J. M. Ashworth and M. Sussman, JBC 242, 1696 (1967). 113. P. C.Newell, J. S. Ellingson, and M. Sussman, BBA 177, 610 (1969). 114. R. Roth, J. M. Ashworth, and M. Sussman, Proc. Nut. Acad. Sm’. U. S. 59, 1235 (1968). 115. R. G. Pannbacker, Biochemistry 6, 1287 (1967). 116. B. Wright and D . Dahlberg, J. Bacteriol. 95, 983 (1968). 117. R. Marchall, D.Sargent, and B. E. Wright, Biochemistry 9, 3087 (1970). 118. B. Wright, W. Simon, and B. T. Walsh, Proc. Nut. Acud. Sci. U . S. 80, 644 (1968). 119. B. E. Wright, J . Cell. Physiol. 72, Suppl. 1, 145 (1968). 120. P. C. Newell and M. Sussman, JBC 244, 2990 (1969). 121. P. C. Newell, M. Longlands, and M. Sussman, J M B 58, 541 (1971).

62

RICHARD L. TURNQUIST AND R. GAURTH HANSEN

only go as fast as the rate a t which substrate is made available (122). However, the enzyme increase may be necessary to overcome UDPglucose product inhibition and allow the observed changes in glucose 1-P levels ( 2 7 ) . The value of these models has been questioned since many parameters may not be accurately known and their estimation might lead to serious error in the results. It is now generally agreed that UDPglucose pyrophosphorylase activity does increase in the slime mold during differentiation, but the effect that this increase has on carbohydrate metabolism is still the subject of debate.

111. Properties

A. OPTIMA All of the UDPglucose pyrophosphorylases that have been studied show an absolute requirement for a divalent cation. Magnesium a t 1-3 mM seems to satisfy this requirement best, while Mnz+, Coz+,and NiZ+ have about one-fourth the optimal activity. The cations become inhibitory a t high concentrations. Mercaptoethanol or dithiothreitol is required for human liver ( 1 7 ) , erythrocyte ( 2 1 ) , and mung bean (19) enzymes to protect them from oxidation. While most UDPglucose pyrophosphorylases can be isolated and stored a t 4", the enzyme from E . coli was found to be labile a t 0" (98). The optimal pH range for activity of the various UDPglucose pyrophosphorylases is broad and generally is slightly alkaline, as follows: yeast, 6.5-8.0 (31) ; E . coli, 7.5-9.0 (28); slime mold, 7.8 (22); pea seedlings, 7.5-9.0 (123); mung bean, approximately 8 (19) ; bovine mammary tissue, 8-9 (20); rat mammary tissue, 7.5-9 (124); human erythrocytes, 8-9 (21); human liver, 7.6-9.2 (17); rabbit muscle, 6.5-8 (26); and calf liver, 8.5 (16).

B. STRUCTURE The molecular weight. and multimeric structure of calf liver UDPglucose pyrophosphorylase have been thoroughly studied. A molecular 122. B. E. Wright,

Behav. Sci. 15, 37 (1970). 123. D. H. Turner and J. F. Turner, BJ SO, 448 (1958). 124. D. K. Fitzgerald, S. Chen, and K. E. Ebner, BBA 178, 491 (1969).

2.

URIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

63

weight of 480,000 was reported, with eight probably identical subunits of molecular weight 60,000 (125).Electron micrographs support these figures (Fig. 1 ) . Ultracentrifugation and other studies indicate that multimers of the enzyme exist. These are probably dimers, trimers, and tetramers of the 480,000 species. The crystalline form of the enzyme is diamond-shaped (16). The enzymes isolated from human liver and human erythrocytes appear to be similar, if not identical, and not greatly different from the calf liver enzyme. The human enzymes are of slightly smaller molecular weight, about 440,000 @ I ) , and, like the calf enzyme, appear to dimerize (17).The human liver enzyme crystallizes as a long needle. R a t mammary gland pyrophosphorylase appears to be similar in size, with a reported molecular weight of 450,000 (124). This enzyme increases markedly in activity during incubation of organ explants and tissue extracts. The increase is not dependent upon hormonal activity and is not inhibited by puromycin. The molecular weight does not change, nor does the K , for UTP, although the V,,, increases markedly. The increase in activity is a function of pH, temperature, and enzyme concentration and is inhibited by urea. The protein molecule appears to undergo structural changes that lead to a more active form of the enzyme. The slime mold, D . discoideum, appears to have two forms of UDPglucose pyrophosphorylase. The largest fraction (90%) has a molecular weight of 390,000 with subunits of about 55,000 (22). Another lighter fraction comprising about 10% of the total activity has also been found (190).The two fractions are not interconvertible and the lighter is much more labile. The different forms may provide UDPglucose for different metabolic functions and thus may be separately controlled. In the microorganism SaZmoneZZa typhimurium, UDPglucose pyrophosphorylase exists in different forms, which have been designated 11, IIIa, and IIIb. A fourth form, IV, is found in certain mutants. The four enzymes differ significantly in their reaction kinetics, pH optima, and heat stability, but all have UDPglucose pyrophosphorylase activity (1266). Two genes are involved in the synthesis of the enzyme. When one gene (gal F ) is deleted, the only form that appears is IV. The second gene (gal U ) is a structural gene that codes for a polypeptide found in all forms of the enzyme. The presence of the gal F gene modifies the basic polypeptide into forms 11, IIIa, and IIIb (127).The molecular 125. S. Levine, T. A. Gillett, E. Hageman, and R. G. Hansen, JBC 244, 5729 (1969). 126. T. Nakae and H. Nikaido, JBC 246, 4386 (1971). 127. T. Nakae and H. Nikaido, JBC 248, 4397 (1971).

FIQ.1. Crystalline (top) and molecular (625,OOOX) structure of bovine liver UDPglucose pyrophosphorylase. 04

2.

URIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

65

FIG.2. hterconversion of enzyme monomers and dimer.

weights of isozymes 11, IIIa, IIIb, and I V are 40,000, 40,000, 8O,OOO, and 8O,OOO, respectively. Probable interconversions are shown in Fig. 2. The present thinking is that gal U codes for the basic 80,000molecular weight dimer (IV), which is stable. I n the presence of gal F , however, the enzyme is split into two molecules of monomer IIIa. Form I I I a can be converted to dimer I I I b or into monomer I1 by cell factors that are not associated with either gene and can be reproduced in vitro (128). Since all forms have different UDPglucose inhibition constants, the variety of isozymes may serve some regulatory function.

C. KINETICS The substrate affinities for UDPglucose pyrophosphorylase from various sources are indicated by the Michaelis constants shown in Table I (129-131). Turnover numbers vary from 83,000 for calf liver enzyme (16) to 8,900 for slime mold pyrophosphorylase (22). Table I1 shows inhibition constants for UDPglucose pyrophosphorylase. Uridine diphosphoryl glucose shows a highly selective product inhibition, especially in animal cells, and therefore probably exerts considerable self-regulation in metabolic control. The dissimilarity of inhibition constants between mammalian and plant enzymes suggests a difference of biological significance ( 2 1 ) . Several workers have found competitive inhibition between UDPglucose and UTP, and noncompetitive inhibition between U T P and PPi (17,21, 22) ; PPi shows noncompetitive product inhibition with both substrates in the slime mold ( 2 2 ) and is inhibitory in high concentra128. 129. 130. 131.

T. Nakae, JBC 246, 4404 (1971). I. T. Oliver, BBA 52, 75 (1961). G. L. Gustafson and J. E. Gander, JBC 247, 1387 (1972). A . Munch-Petersen, Acta Chem. Scand. 9, 1523 (1955).

66

RICHARD L. TURNQUIST AND R. GAURTH HANSEN

TABLE I SUBSTRATE AFFINITIES FOR UDPQLUCOSE PYROPHOSPHORYLASE FROM VARIOUS SOURCES K, X 106 Ensymesource Human liver Human erythrocyte Human erythrocyte Calf liver Bovine mammary Rat mammary Rat liver Guinea pig brain Rabbit muscle Dog heart Mung bean Mung bean Pea seeds Wheat Sorghum Slime mold Yeast Yeast E . wli E. w2i S . typhimurium Form I1 Form IIIa Form IIIb Form IV

Ref. K,,+UDPG i7 91 86 16 I0 94 199 i29 2% 91 19 Ii iI.9 62 130 $9 131

0.15 0.26 0.28-0134

G-1-P 9.5 17 6.7 5.5 11 39 18-28 4.5-8.6

UTP

PPi

4.8 33 3.3 20 14

21 48

5 2.3

8.4 100

6

10

4.5 2.8 11 11

36 16 23

8.5

0.14

100 0.25-0.33 1

4.8 26

3.0 11

4.8

2.9

3.4 2.9

7.4 14.4

3.6

15.9

5.4 44

98

98 28 126

UDPG

0.2

310 13

5.6 17 7 9 25 13 84 67 37 27

tions as a substrate in E. coli (98). The product inhibition by PPi of the slime mold enzyme is suggested as a possible alternate regulatory mechanism for the alleged changes in intracellular substrate concentration (28). Calf liver enzyme appears to be sensitive to Pi,which inhibits competitively with PPi at physiological concentrations. This may provide a mechanism whereby an increase in Pi,as the result of energy drain and hydrolysis of ATP,would result in a lowering of UDPglucose production and a redirection of glucose metabolism from storage to glycolysis. The inhibition by UDP, a product of glycogen synthesis, may also be significant in vivo in that it must be converted to UTP, an energy requiring reaction, for glycogen synthesis to proceed. If energy (ATP)reserves are low, glycogen synthesis would be inhibited ( 1 6 ) .

2.

67

URIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

TABLE I1 FOR UDPQLUCOSE PYROPHOSPHORYLASE INHIBITION CONSTANTS Product Ki Enzyme source

Ref.

Human liver Human erythrocyte Calf liver Dog heart Mung bean Sorghum Slime mold S. lyphimurium Form I1 Form IIIa Form IV

17 81

UTP

x

UDPG

81 130 97 186

Pi with PPi

UDP with UDPG 10

10

1.5

8 7

2.3

370

16

81

Ki X 106

106

10.4

15

16 5.0 5 10 6.3 1.3

3000-9000

400

Uridine diphosphoryl glucose is necessary for the metabolism of galactose. Galactose 1-P in high concentrations (50-fold excess over glucose 1-P) will competitively inhibit the pyrophosphorylase (129). Inhibition of the enzyme slows production of UDPglucose, which causes further accumulation of galactose 1-P, which in turn further inhibits the enzyme. Thus a cyclic process is established in which a compound inhibits its own metabolism (132, 133). D-Galactosamine is also a competitive inhibitor with glucose 1-P. It, too, requires UDPglucose for its metabolism; thus, a similar inhibition cycle is established (134, 136). The loss of UDPglucose not only affects the metabolism of the inhibitory compounds but also reduces glycogen, polysaccharide, and glucuronide metabolism, Certain pathological conditions may increase the concentrations of these inhibitors enough for them to limit UDPglucose synthesis (129). In E . coli, both UDPglucose and TDPglucose are needed for the first steps of synthesis of the polysaccharide component of antigens. Uridine diphosphoryl glucose pyrophosphorylase is inhibited by TDPglucose ( K i= 2 x The inhibition may serve a useful purpose in that blockage of later stages of polysaccharide synthesis by phages will result in the accumulation of both UDPglucose and TDPglucose. The cross 132. E. L. Talman, Physiol. Chem. Phys. 1, 131 (1969). 133. E. L. Talman, Physiol. Chem. Phys. 1, 255 (1969). 134. D. Keppler and K. Decker, Eur. J . Bbchem. 10, 219 (1969). 135. D. 0. R. Keppler, J. F. M. Rudigier, E. Bischoff, and K. Decker, Eur. J . Biochem. 17, 246 (1970).

68

RICHARD L. TURNQUIST AND R. GAURTIl HANSEN

inhibition may reinforce normal product inhibition and thus more effectively shut down early synthetic steps. The enzyme is also inhibited by TDPrhamnose (Ki = 5.2 X which is a final product of carbohydrate metabolism in the organisms (98). Chloroazanil, a triazine derivative, has also been found to be an inhibitor of UDPglucose pyrophosphorylase (136).

D. SPECIFICITY Considering its relatively high turnover number, its favorable equilibrium, and its low K,,, values for glucose 1-P and UTP, the prime function of UDPglucose pyrophosphorylase is probably to catalyze the formation of UDPglucose. However, the enzyme shows activity toward other substrates as well. Calf (16) and human (17)liver enzymes have been used to catalyze the phosphorylation of UDPglucose, TDPglucose, CDPglucose, GDPglucose, UDPgalactose, UDPxylose, and UDPmannose. Therefore, the enzyme is not absolutely specific for either the base or the sugar moiety of the substrate. The percent of initial reaction velocities compared to UDPglucose varies from 2.2% for TDPglucose to 0.1% for GDPglucose (17). Reaction rates of (‘abnormal” substrates may be increased if concentrations are raised. While UDPgalactose is pyrophosphorylated by human liver enzyme at about 2% of the rate of UDPglucose at equal substrate concentrations, the rate may be increased to as much as 10% a t higher concentrations (17). This nonspecificity of human UDPglucose pyrophosphorylase has led to postulations about its physiological importance in patients with galactosemia. Galactosemia is a molecular disease in which galactose is not metabolized because of a lack of galactose 1-P uridylyltransferase (EC 2.7.7.12). The result is an accumulation of galactose 1-P. There is, however, some conversion between glucose 1-P and UDPgalactose ( 1 S 7 ) , giving rise to the speculation that a UDPgalactose pyrophosphorylase (EC 2.7.7.10) might be present in human blood (138). Such activity has been described in microbes (139), rats, pigeons, and humans (138, 140). This enzyme might increase as a child ages, thereby lessening the symptoms of the 136. W. Kreutner and N. 0. Goldberg, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 26, 508 (1967). 137. R. Gitzelmann, Pediat. Res. 3, 279 (1969). 138. H. D. Abraham and R. R . Howell, JBC 244, 545 (1969). 139. G. T. Zancan, Can. J . Microbwol. 17, 563 (1971). 140. X. J. Isselbacher, JBC 232, 429 (1958).

2.

URIDINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

69

disease (43). The existence of such an enzyme has been questioned, however, since UDPglucose pyrophosphorylase, which is present in galactosemics, will catalyze the same reaction. Further, in studies in which UDPgalactose pyrophosphorylase activity was reported, no effort was made to separate this from UDPglucose pyrophosphorylase activity (141). Additionally, the ratio of the two activities remained constant throughout purification from human liver. The ratio was also constant during sucrose gradient sedimentation and polyacrylamide gel electrophoresis, indicating that both reactions are catalyzed by the same enzyme (17). Uridine diphosphoryl galactose pyrophosphorylase activities reported for human erythrocytes have been quite low, about 1% of the UDPglucose pyrophosphorylase activity (21, 138), and some workers found no activity a t all (86, 142, 149). This is surprising since, even in the absence of a specific UDPgalactose pyrophosphorylase, UDPglucose pyrophosphorylase should show some activity. The lack of activity in these cases might result from insufficient concentrations of UDPgalactose or galactose l-P in the in vitro assay mixtures. Low concentrations of the substrates would not overcome the high K , values of UDPglucose pyrophosphorylase for these substrates. In a galactosemic victim, however, the galactose l-P concentration might be increased to a point where significant catalysis could occur. Based on the above evidence it would seem likely that UDPgalactose pyrophosphorylase is not present in human red blood cells, and the activity found is probably attributable to UDPglucose pyrophosphorylase.

E. MECHANISM The most detailed study of the mechanism of UDPglucose synthesis utilized calf liver enzyme. I n one set of experiments the enzyme was incubated with substrate or substrate analogs and chromatographed through Sephadex G-25-80 (144). Uridine diphosphoryl glucose and U T P formed stable complexes with the pyrophosphorylase, while UMP, PPi, UMP plus PPi, glucose l-P and UDP did not. Since UDPglucose and U T P were doubly labeled and both labels appeared in the enzymesubstrate complex, it is likely that the entire substrate molecules were 141. 142. (1964). 143. (1967). 144.

W. K. Ting and R. G. Hansen, Proc. SOC.Exp. Biol. Med. 127, 960 (1968). W. G. Ng, W. R . Bergren, and G. N. Donnel, Nature (London) 203, 845 W. G. Ng, W. R. Bergren, and G. N. Donnel, Clin. Chim. Acta 15, 489

T. A . Gillett, S. Levine, and R. G. Hansen, JBC 2 4 , 2551 (1971).

70

RICHARD L. TURNQUIST AND R. GAURTH HANSEN

bound. Only after prior incubation with UDPglucose or UTP could PPi be complexed with the enzyme, indicating an obligatory order of substrate binding. Magnesium was not required for the binding of UDPglucose but was necessary for binding PPi, which replaced glucose 1-P from the enzyme-UDPglucose complex. Thus, it appears an ordered Bi-Bi mechanism is most likely, with the nucleoside phosphate the first substrate to be added and the last product to leave, as follows: G-1-P

IJTP

+E

E(UTP)

11

Jt

E(UDPG)

+ MgPPi

E

+ UDPG

(4)

Mg*+

Human erythrocyte pyrophosphorylase seems to have a similar mechanism ( 2 1 ) . Uridine diphosphoryl glucose and UTP inhibit one another competitively, while the other substrate-product combinations show noncompetitive inhibition. A distinguishing characteristic of the ordered Bi-Bi mechanism is competitive inhibition between the first substrate added and the last product released ( 1 4 6 ) . Calculating the equilibrium constant from experimental kinetic data according to a Haldane relationship was also consistent with an ordered Bi-Bi mechanism. Although there is considerable variation in kinetics and enzyme structure, UDPglucose pyrophosphorylases from yeast (16), mung bean (4, 19, d l ) , dog heart (.%?I), and slime mold (29) also appear to catalyze the formation of UDPglucose by the mechanism described in Eq. (4). Metal activated enzymes such as UDPglucose pyrophosphorylase may bind metal and substrate in one of four possible coordination schemes. These include metal bridge complexes (E-M-S) that may be either simple or cyclic, enzyme bridge complexes (M-E-S) , and substrate bridge complexes (E-S-M) (14.6’). Calf liver UDPglucose pyrophosphorylase appears to form a substrate bridge complex with the substrate serving as the only attachment of the metal to the enzyme. Several lines of evidence indicate this. Complexes of this type are generally limited to enzymes with nucleoside di- and triphosphate substrates that show high affinity for the metal. Since the enzyme will bind substrate equally well with or without the metal, simple and cyclic metal bridge complexes tend to be ruled out. The metals in substrate bridge complexes will not be bound to the enzyme unless the substrate is present. Studies measuring longitudinal proton magnetic relaxation rates (PRR) have demonstrated little or no enhancement unless all three complex components 145. W. W. Cleland, BBA 67, 104 (1963). 146. A. S. Mildvan, “The Enzymes,” 3rd ed., Vol. 2, p. 445, 1970.

2.

71

U R I D I N E D I P H O S P H O R Y L GLUCOSE PYROPHOSPHORYLASE

were present (147’). Calcium will often inhibit enzymes that form metal bridge complexes (E-M-S), and it can serve as an activator of substrate bridge enzymes. Calcium will serve as an alternative activator for UDPglucose pyrophosphorylase (148). Based on the above evidence, the divalent cation seems to activate the phosphorous atom of the substrate that is to be attacked by deshielding the 8- and 7-phosphorous atoms, thus making them more susceptible to substitution. The phosphate ligands probably complex with the metal by replacing its coordinated water molecules (146). Two pieces of PRR evidence, however, do not fit the above mechanism. First, there is no indication that an enzymepyrophosphate-metal complex will form. Second, when the E-S-M complex forms, free metal is released into solution (147).This indicates that more UTP is bound than metal, which makes a substrate bridge complex unlikely. Two possible explanations have been offered (149). First, an enzyme bridge complex (M-E-S) may form in which the binding of UTP weakens the metal binding by causing changes in enzyme conformation allowing the metal to be released. Such a situation, however, would be expected to allow the enzyme to bind the metal whether the substrate was present or not. Evidence indicates this does not occur. A, second, more attractive explanation is that an E-S-M complex forms in which the metal is bound more strongly to the nucleotide before the ternary complex is formed. The metal may be bound to the UTP in a tridentate (Y,p, y coordination. When the metal-UTP binds to the enzyme, the metal shifts to coordination which is much weaker than the tridentate coordination. (Y

.A/

M 0 I

b o\’I

I R-0-P-0-P-0-P-0 I I

0 f

0

f

P

+enzyme

I

-enzyme

0 Y

0

- R-0-P-0I

I

I

-P-O-P-O I

0 f

0

0 f

P

I I 0 Y

The weaker bond may allow some of the metal to be released from the complex. Neither of the above mechanisms has been proved, but the accumulated evidence would make the second more likely. ACKNOWLEDQMENT

This work was supported by National Institutes of Health Grant AM13709.

147. G. H.Reed, H. Diefenbach, and M. Cohn, JBC 247, 3066 (1972). 148. R. G. Hansen, unpublished observations. 149. A. S. Mildvan and M. Cohn, Advan. Enzymol. 33, 1 (1970).

This Page Intentionally Left Blank

A denosine D;Phosphory1 Glucose Pyrophosphorylase JACK PREISS

. .

.

I. Introduction . . . . . . . . . . 11. Classification of ADPglucose Pyrophosphorylasee . . . . 111. Kinetic Properties of the ADPglucose Pyrophosphorylases . . A. General Effects of Activator . . . . . . . B. The ADPglucose Pyrophosphorylases of Microorganisms Degrading Glucose via the Entner-Doudoroff Pathway . C. The ADPglucose Pyrophosphorylase of R hodospirillum rubrum . . . . . . . . D. ADPglucose Pyrophosphorylases of Higher Plants and Green Algae . . . . . . . . E. The ADPglucose Pyrophosphorylases of the Enterobacteriaceae . . . . . . . . . F. The ADPglucose Pyrophosphorylase of Serratia marcescens G. The ADPglucose Pyrophosphorylase of Aeromonas jormicans H. The Kinetic Properties of ADPglucose Pyrophosphorylases Isolated from E . coli B Mutants Altered in Their Ability to Accumulate Glycogen . . . . . . IV. Physical Properties of the ADPglucose Pyrophosphorylases . .

.

.

.

.

. .

73 75 77 77 78

81 86 94 107 108

109 117

1. Introduction

In 1961, Recondo and Leloir showed that adenosine diphosphoryl glucose (ADPglucose) was 10-fold more active than UDPglucose in the transfer of glucose from the sugar nucleotide to the starch granule by the starch synthetase ( 1 ) . This study was made with chemically synthesized 1. E. Recondo and L. F. Leloir,. BBRC 6, 85 (1961). 73

74

JACK PREISS

ADPglucose. Subsequently, the enzyme responsible for the synthesis of ADPglucose from glucose-1-P and ATP, ADPglucose pyrophosphorylase (synthase), was isolated and identified ( 2 ) . The enzyme required Mg2+ for activity. Adenosine diphosphoryl glucose was later shown to be a naturally occurring metabolite in various algal (3, 4 ) and plant systems (5-7) and in Salmonella typhirnurium (8, 9). The main, if not the sole, function for ADPglucose in nature is as a precursor of the glucosyl residues in bacterial glycogen and plant starch. The reactions responsible for the synthesis of a-1,4-glucans are reaction ( l ) ,catalyzed by ADPglucose pyrophosphorylase ; and reaction (2), catalyzed by ADPglucose, (~-1,4glucan-4-glucosyltransferase. ADPglucose pyrophosphorylase is found only in those tissues actively engaged in a-1,4-glucan synthesis.

+ +

ATP a-glucose-1-P ADPglucose a-1,Pglucan

+

ADPglucose PPi a-1,4-glucosyl glucan

+ ADP

(1) (2)

The regulation of a-1,4-glucan synthesis occurs both in plants and in bacteria. Many bacteria have the ability to accumulate glycogen when they reach stationary phase, provided an excess carbon source is available. The biosynthesis of this carbon reserve material in bacteria is regulated; thus, it does not occur in the exponential phase of growth and only occurs in conditions of nongrowth. Likewise, starch levels in green algae and in higher plants are dependent on light conditions. Exposure of the plant leaf to light results in an increase in the number of starch granules in the chloroplasts, while placing the leaf in the dark results in the diminution and eventually in the disappearance of the starch granules. An important feature of the enzyme catalyzing the synthesis of ADPglucose is its activation by glycolytic intermediates and inhibition by 5’-adenylate, ADP, or orthophosphate. Modulation of the activity of the ADPglucose pyrophosphorylasc by the above metabolites has been the main emphasis in studies of thc ADPglucose pyrophosphorylase from many sources. These studies strongly suggest that regulation of a-1,4glucan synthesis occurs at the level of ADPglucose synthesis and that both ADPglucose and a-1,4-glucan synthesis are regulated by the level 2. J. Espada, JBC 237, 3577 (1962). 3. H . Kauss and 0. Kandler, 2. Nnturforsch. B 17, 858 (1962). 4. T. Kanazawa, K. Kanazawa, M. R. Kirk, and J. A. Bassham, BBA 258, 656 (1972). 5. E. Recondo, M. Dankert, and L. F. Leloir, BBRC 12, 204 (1963). 6. T. Murata, T. Minamikawa, and T . Akazawa, BBRC 13, 439 (1963). 7. A. C. Cassells and M. A. Harmey, ABB 128, 486 (1968). 8. V. Ginsburg, JBC 241, 3750 (1966). 9. R. M. Scher and V. Ginsburg, JBC 243, 2385 (1968).

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

75

of glycolytic intermediates and the energy state in the cell. The description of these kinetic studies and discussion of their physiological significance are the major aspects of this chapter.

II. Classification of ADPglucose Pyrophosphorylares

A consistent pattern is usually observed between the source (plant or micoorganism 1 of the ADPglucose pyrophosphorylase, the metabolites that are most effective as activators of the enzyme, and the type of carbon utilization pathway occurring in that tissue or organism (Table I). The Enterobacteria accumulating glycogen (Escherichia coli, Aerobacter aerogenes, Aerobacter cloacae, Salmonella typhimurium, Citrobacter freundii, and Escherichia aurescem) contain an ADPglucose pyrophosphorylase activated by fructose-dip, TPNH, and pyridoxal-5’-P and inhibited by 5’-adenylate (1&13). These organisms are known to use glycolysis as their main route for glucose catabolism. Another class of ADPglucose pyrophosphorylase is found in Aeromonas formicans which is activated by either fructose 6-P or fructose 1,6-diP (M. Paule, C. Lammel, and J. Preiss, unpublished results). In contrast, Serratia marcescens, an enteric, has an ADPglucose pyrophosphorylase not activated by any metabolite tested ( I d ) . Both Aeromonas and Serratia also degrade glucose via the Embden-Myerhof pathway. A group of organisms that cataboliae glucose via the Entner-Doudoroff pathway contains an ADPglucose pyrophosphorylase activated both by fructose 6-P and pyruvate (14-17).Rhodospirillum rubrum, a photosynthetic organism that cannot metabolize glucose but can grow either as a heterotroph in the light or dark on various tricarboxylic acid intermediates and associated metabolites, or as an autotroph on CO, and HB, contains an ADPglucose pyrophosphorylase that is activated specifically by pyruvate (18-20). The ADPglucose pyrophosphorylase found in leaves 10. J. Preiss, L. Shen, and M. Partridge, BBRC 18, 180 (1965). 11. J. Preiss, L. Shen, E. Greenberg, and N. Gentner, Biochemistry 5, 1833 (1965). 12. G. Ribereau-Gayon, A. Sabraw, C. Lammel, and J . Preiss, ABB 142,675 (1971). 13. N. Gentner, E. Greenberg, and J . Preiss, BBRC 36, 373 (1969). 14. L. Shen and J. Preiss, BBRC 17, 424 (1964). 15. L. Shen and J . Preiss, ABB 116, 374 (1966). 16. L. Eidels, P. L. Edelmann, and J . Preiss, ABB 140, 60’(1970). 17. I,. Eidels and J . Preiss, ABB 140, 75 (1970). 18. C. E. Furlong and J . Preiss, Progr. Photosyn. Res. 3, 1604 (1969). 19. C. E. Furlong and J . Preiss, JBC 244, 2539 (1969). 20. M . R. Paule, Biochemistry 10, 4509 (1971).

4

m

ACTIVATORS.4ND Source Enterobacteria

Amomonas formicans

INHIBITORS ' O F

Primary activators Fructose 1,6--diP TPNH Pyridoxal P Fnictose 6-P Fructose I,6-diP None Fructose 6-P Pyruvate

Serratia marmscem Arthrobacter viscosus Agrobaderium tumefaciens Rhodopseudomonas capsulaln Rhodospirillum m b u m Pyruvate Plant leaves Green algae

3-Phosphoglycerate

TABLE I ADPGLUCOSE PYROPHOSPHORYLASES FROM VARIOUS

SOURCES

Secondary activatorsa

Inhibitors

ZPhosphogly cerate 3-Phosphogly ceraldehyde Phosphoenolpyruvate -

5'-AMP

Glycolysis

ADP

Glycolysis

5'-AMP Pi AMP ADP None

Glycolysis Entner-Doudoroff pathway

Ribose 5-P ZDeoxyribose 5-P None Fructose 6-P Fructose-diP

P,

Carbon utilization pathways

Glucose-not met,abolized, anaerobic TCA cycle, reductive dicarboxylic acid cycle, Calvin-Bassham cycle Calvin-Bassham cycle, Hatchslack pathway

0 Secondary activators are those compounds that activate to lesser extents than the primary activators. Considerably higher concentrations of the secondary activators are required to elicit the activation response.

4

r

0

w

'd

8

E x

7m

3.

ADENOSINE DIPHOSPHORTL GLUCOSE PYROPHOSPHORYLASE

77

of plants (21-25) and in green algae ( 2 6 ) is activated by 3-phosphoglycerate. The plant and algal ADPglucose pyrophosphorylases are very sensitive to inhibition by inorganic phosphate. The activators for the various groups of ADPglucose pyrophosphorylase are significant metabolites in the metabolic pathways utilized by the plant tissues or microorganisms. The significance of these metabolites in their respective pathways and their activation of ADPglucose and a-1,$-glucan synthesis has been discussed previously in reviews or research articles (12, 18, 19, 2 7 ) . Table I shows that the activator site is relatively nonspecific for most ADPglucose pyrophosphorylases. A number of glycolytic intermediates are capable of activating all classes of the enzyme albeit a t varying effectiveness. This overlapping of specificity for the activators in the various ADPglucose pyrophosphorylase groups suggests that the activator sites for the different classes arc very similar to each other. Thus, one may hypothesize that mutation of part of the gene specifying the activator site of the ADPglucose pyrophosphorylase has occurred via evolutionary processes to enable the specificity of the activator site to be compatible or coordinated with the metabolic activities going on in the organism. If true, this would suggest that the genetic information specifying the activator site for the ADPglucose pyrophosphorylase of S. marcescens has been lost during evolution.

111. Kinetic Properties of the ADPglucose Pyrophosphorylases

A. GENERAL EFFECTS OF ACTIVATOR

Several effects of the activators on the pyrophosphorylase have been observed. The concentrations of the substrates, ATP, glucose 1-P, PPi and ADPglucose, required to give 50% of maximal velocity (So.6or K , in those cases where the saturation curve follows Michaelis-Menten kinetics) is significantly lowered in the presence of activators. For most 21. H. P. Ghosh and J. Preiss, JBC 240, 960 (1965). 22. H. P . Ghosh and J. Preiss, JBC 241, 4491 (1966). 23. J. Preiss, H. P. Ghosh, and J. Wittkop, i n “The Biochemistry of Chloroplasts” (T. W. Goodwin, ed.), Vol. 2, p. 131. Academic Press, New York, 1967. 24. G. G. Sanwal, E. Greenberg, J. Hardie, E. C. Cameron, and J. Preiss, Plant Physiol. 43, 417 (1968). 25. P. W. MacDonald and G. A. Strobel, Plant Physwl. 46, 126 (1970). 26. G. G. Sanwal and J. Preiss, ABB 119, 454 (1967). 27. J . Preiss, Curr. T o p . Cell. Regul. 1, 125 (1969).

78

JACK PREISS

ADPglucose pyrophosphorylases the So.5value of ATP and ADPglucose is lowered about 5-15-fold in the presence of activator. The activator may also increase the maximal velocity for ADPglucose synthesis anywhere from 2.5-60-fold. The magnitude of activation is dependent on the source of enzyme and pH. Thc activators also modulate the sensitivity of the various ADPglucose pyrophosphorylases by the various inhibitors, and this is discussed in detail for a number of the enzymes below.

B. THEADPGLUCOSE PYROPHOSPHORYLASES OF MICROORGANISMS DEGRADING GLUCOSE VIA THE ENTNER-DOUDOROFF PATHWAY As seen in Table I those organisms that catabolize glucose via the Entner-Doudoroff pathway contain an ADPglucose pyrophosphorylase that is activated by fructose 6-P and pyruvate. Other compounds capable of activating but to lesser extents or at higher concentrations are deoxyribose 5-P, ribose 5-P and 2-keto-3-deoxy-phosphogluconate.These latter activators may be considered as analogs of fructose 6-P in that they are furanoside sugar phosphates. 2-Keto-3-deoxy-phosphogluconatealso is similar to pyruvate in structure. Other pyruvate analogs that activate slightly and at higher concentrations are a-ketobutyrate and hydroxypyruvate. However, compounds capable of activating other classes of ADPglucose pyrophosphorylases such as fructose-dip and 3-phosphoglycerate do not activate the enzymes present in Rhodopseudomonas capsulata (17),Arthrobacter viscosus ( l b ) , and Agrobacterium tumejaciem ( 1 7 ) . Table I1 lists some of the kinetic constants for the activators fructose 6-P and pyruvate. Both activators increase the maximal velocity of ADPTABLE I1 KINETIC PARAMETERS OF ADPGLUCOSE PYROPHOSPHORYLASES FROM OROANISMS CATABOLIZINO GLUCOSE VIA THE ENTNER-DOUDOROFF PATHWAY Stimulationof VmaX of ADPglucose synthesis

Ao.6

Hill fi Fructose Fructose Source of ADPglucose f3-P Pyruvate &P Pyruvate Fructose pyrophosphorylase (pM) (pM) (-fold) (-fold) f3-P Pyruvate

A . viscoaua R. capaulata A . lumefaciens

110

77 220

310 55 80

7.0 6.5 5.5

6.0 3.5 4.0

1.7 1.3 1.1

1.0 1.3 1.1

3.

79

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

glucose synthesis about 3.5-7-fold. The concentrations required for halfmaximal stimulation of V,,,,, A,.,, range from 55 to 310 p M for pyruvate and 77 to 220 pM for fructose 6-P. The fructose 6-P saturation curve for the A . viscosus enzyme is sigmoidal giving a Hill constant, f i value ($8, 29) of 1.7 while the pyruvate curve is hyperbolic. For the other pyrophosphorylases in this class the fructose 6-P and pyruvate saturation curves are slightly sigmoidal or hyperbolic. Fructose 6-P also increases the apparent affinity of the various substrates for the R. capsulata, A . viscosus, and A . tumefaciens enzymes. The values (concentration of substrate needed for 50% of maximal velocity) of ATP, glucose-l-P, pyrophosphate, and ADPglucose are decreased about 2-5-fold in the presence of activator with the R . capsulata enzyme, about 2-fold with the A . tumefaciens pyrophosphorylase, and 3-5-fold with the A . viscosus enzyme. Although pyruvate increases the maximal velocity it appears to have little effect on the So., values of the substrates for three ADPglucose pyrophosphorylases in this group. There are some variations in the shape of the ATP and ADPglucose saturation curves among the three ADPglucose pyrophosphorylases. The saturation curves for these substrates are hyperbolic with the A . tumefaciens and A . viscosus enzymes but are slightly sigmoidal for the R. capsulata enzyme. The presence of activator does not change the shape of the ATP saturation curves and saturation curves for PPi and glucose l-P in the presence or absence of activator are hyperbolic. However, the Mg2+saturation curves for all three enzymes are highly sigmoidal giving Hill constant A values of 3.8-5.0; this value is relatively unaffected by the activator. With this class of ADPglucose pyrophosphorylases there appears to be little interaction among activator sites or among substrate sites. It was found that inorganic phosphate, sulfate, 5’-AMP, ADP, phosphoenolpyruvate (PEP), GMP, and G D P are effective inhibitors of the A . viscosus ADPglucose pyrophosphorylase. I n all cases the activators fructose 6-P and pyruvate can reverse or antagonize the inhibition caused by the above inhibitors. Increasing concentrations of fructose 6-P or pyruvate can completely reverse the inhibition caused by sulfate, phosphate, 5’-AMP, or ADP concentrations in the 1-2.0 mM range. The inhibitors do not change the shape of the fructose 6-P and pyruvate saturation curves with the Hill A values remaining 1.7 to 1.8 and 1.0, respectively. I n the absence of activator, the 1 0 . 5 values (concentration of inhibitor required for 50% inhibition) for phosphate, sulfate, 5’-adenylate, ADP, PEP, GDP, and GMP are 0.58, 0.45, 0.7, 0.6, 0.060, 0.080, and J. Hill, BJ 7, 471 (1913). 29. J. P. Changeux, Cold Spring Harbor S y m p . Quant.

28. A.

Bwl.28,

497 (1963).

80

JACK PREISS

0.034 mM, respectively. The 5’-AMP and ADP saturation curves are sigmoidal while the other inhibitor curves are hyperbolic in shape. Rhodopseudomonas capsulata enzyme is inhibited by the same compounds as the A . viscosus pyrophosphorylase. However, the interactions between inhibitors and activators are slightly different for the R. capsu2ata enzyme. Fructose 6-P cannot completely overcome the inhibition caused by phosphate or ADP. I n addition, the inhibitors increase the sigmoidicity of the fructose 6-P saturation curve. For example, a t 0.25 M Pi the for fructose 6-P is increased from 76 to 112 pM and the Hill a is increased from 1.4 to 1.8. At a higher concentration of phosphate, 1.25 mM, the fi for fructose 6-P is 2.5 and A0.5is 258 tJM. Table I11 shows that the inhibitor saturation curves for R. capsulata are hyperbolic (Pi, PEP, and AMP) or slightly sigmoidal (ADP). However in the presence of activator the inhibitor curves become sigmoidal. Thus, there are some slight variations between the enzymes in this group with respect to activator and inhibitor responses. It should be noted that in the R. capsulata system PEP is the most effective inhibitor. Along with G M P and GDP, PEP is also the most effective inhibitor for the A . tumefaciens and A . viscosus ADPglucose pyrophosphorylases. The physiological significance of the PEP inhibition is not clear. It is difficult to reconcile the in vitro inhibition of the ADPglucose pyrophosphorylase by PEP with the notion that increased PEP levels in the cell would be expected to occur under conditions of “high energy charge.” Nevertheless, it should be pointed out that the PEP in-

TABLE I11 INHIBITION

OF

IN

Inhibitor Phosphate

5’-AMP ADP PEP

R. capsdata ADPGLUCOSE PYROPHOSPHORYL.4SE

PRESENCE AND ABSENCE OF ACTIVATOR Activator concn. (mM 1

(mM)

Hill ii for inhibitor

None F6P, 0 . 2 F6P, 1 . 0 Pyruvate, 1 . 1 None F6P, 1 . 0 None F6P, 0 . 3 F6P, 1 . 0 None F6P, 0 . 2 FBP, 1 . 0

0.26 1.13 5.8 3.2 0.64 7.3 0.41 1.3 1.8 0.047 0.17 1.6

0.9 2.0 2.8 1.4 1.1 2.3 1.3 1.7 1.8 0.9 1.3 1.7

10.6

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PPROPHOSPHORYLASE

81

hibition is minimal in the presence of fructose 6-P. It would be of interest to know what the relative concentrations of these effector molecules would be in these bacterial cells under conditions where glycogen is being synthesized. Another point of interest with the A . viscosus, R. capsulata, and A . tumefaciens pyrophosphorylases is that these enzymes are relatively nonspecific with respect to activity seen with nucleoside triphosphates other than ATP, and sugar nucleotides other than ADPglucose when compared to the ADPglucose pyrophosphorylases of the other classes. I n the presence of the activator, fructose 6-P, C T P has about 8% the activity observed for ATP with the A . viscosus enzyme, and 2.5% of the A T P activity with the R. capsulata and A . tumefaciens ADPglucose pyrophosphorylases. Other nucleoside triphosphates giving greater than 1% of the activity observed with ATP with the above enzymes are dATP, UTP, and XTP. An effect of fructose 6-P and pyruvate on the A . viscosus ADPglucose pyrophosphorylase is the shift of the pH optimum from 10.0 to about 8.0. I n the absence of activator there is very little activity in the range of pH 6.5-7.0. However, in the presence of the activators the activity in the neutral pH range is greatly increased. The p H optimum shift by the activators seen with the A . viscosus enzyme is not observed for the ADPglucose pyrophosphorylases of A . tumefaciens and R. capsulata.

C. THEADPGLUCOSE PYROPHOSPHORYLASE OF Rhodospirillum rubrum Pyruvate is the only glycolytic intermediate capable of activating the R. rubrum ADPglucose pyrophosphorylase (18, 19). The only other metabolite found to activate the enzyme is a-ketobutyrate. Another distinct property of the R . rubrum enzyme is that it is not inhibited by either Pi, 5’-AMP, or ADP. No inhibitor of physiological importance has been found for this enzyme. Pyruvate increases the maximal velocity of pyrophosphorolysis and synthesis of ADPglucose about 2-fold. It also decreases the So.5values for ATP (from 3.4 to 0.36 mM with 5 mM pyruvate) and for ADPglucose (from 2.0 to 0.38 m M with 25 m M pyruvate). The decreases in the K , values for pyrophosphate and a-glucose 1-P, however, are only about 1.5-2-fold. Pyruvate also decreases the So.5value for MgC1, about 1.5-2.0-fold and shifts the p H optimum of ADPglucose synthesis from 8.5 to 7.5. Rhodospirillum rubrum is capable of growth under a number of heterotrophic conditions as well as under autotrophic conditions. Adenosine diphosphoryl glucose pyrophosphorylase activity is seen whether the cells

82

JACK PREISS

are grown aerobically in the dark with malate or anaerobically in the light with either malate, acetate, acetate + CO,, or CO, + H,. The activator specificity of the pyrophosphorylase does not change with cells grown under different conditions, Thus, the metabolite pyruvate alone is important in the regulation of glycogen synthesis in R. rubrum. This is consistent with the observations made by Stanier et al. in 1959 (SO). These investigators showed that incubation of starved cells of R. rubrum in the light with either succinate, malate, or pyruvate caused accumulation of glycogen. If the cells were incubated with acetate or butyrate, the reserve polymer that accumulated was poly-P-hydroxybutyrate. Only small amounts of glycogen accumulated under these conditions. The accumulated poly-/?-hydroxybutyrate was utilized if CO, was made available to cells and under these conditions glycogen accumulated. Glycogen would also be formed if R . rubrum was incubated with CO, + acetate or CO, + H,. The pattern of labeling of glycogen by [1-14C]succinate and [ 2-I4C]succinate suggested that the hexose units of the polysaccharide were formed by conversion of the succinate to pyruvate, and subsequent hexose synthesis through a reversal of the glycolytic sequence. Thus, Stanier et aZ. (SO) concluded that compounds (such as succinate, malate, or glutamate) which led to formation of pyruvate resulted in glycogen formation. Data by Kikuchi et aZ. (31) also suggest that in R. rubrum grown in the light under anaerobic conditions, dicarboxylic acids liberate CO, mainly a t the levels of malate and oxaloacetate to yield pyruvate. Since incubation of R . rubrum with acetate gave little glycogen, but incubation of the cells with acetate + CO, did give rise to significant amounts of glycogen, Stanier et al. (SO)suggested that CO, may play an essential role in the formation of C, compounds from acetate by R. rubrum. I n this respect Cutinelli et al. (32) have shown that CO, is an important carbon source during photosynthetic growth of R. rubrum with acetate. The incorporation of CO, specifically into the carboxyl group of alanine and the incorporation of the carboxyl and methyl groups of acetate into the a- and /?-carbon atoms of alanine, respectively, suggested to these investigators the formation of pyruvate by addition of CO, t o an 30. R. Y. Stanier, M. Doudoroff, R. Kunisawa, and R. Contopoulou, Proc. Nut. Acad. Sci. U . S. 45, 1246 (1959). 31. G. Kikuchi, S. Tsuiki, A. Muto, and H. Yamada, in “Microalgae and Photosynthetic Bacteria” (Japanese Society of Plant Physiologists, eds.), p. 547. Univ. of Tokyo Press, Tokyo, 1963. 32. C. Cutinelli, G. Ehrensvard, L. Reio, E . Saluste, and R. Stjernholm, Ark. ZCemi 3, 315 (1951).

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

83

acetyl derivative. Buchanan et al. (33) have recently demonstrated the formation of pyruvate from GO, and acetyl-CoA in cell-free extracts of R. rubrum that had been grown on CO, and H,. This enzymic reaction requires reduced ferredoxin (FDH,) . Acetyl-CoA

+ COz + ferredoxin.Hs+ ferredoxin + CoA + pyruvate

Thus mechanisms for the synthesis of pyruvate, the allosteric activator of ADPglucose synthesis in R. rubrum, are available in this photosynthetic organism grown under various nutritional conditions that give rise to accumulation of glycogen. In this respect the demonstration of the following reaction in R . m b r u m by Buchanan and Evans (34) is pertinent to glycogen synthesis in this organism. Pyruvate

+ ATP + PEP + AMP + Pi

This unique reaction is catalyzed by phosphoenolpyruvate synthase and is distinct from pyruvate kinase. Two energy rich bonds of A T P are cleaved to give rise to PEP AMP + Pi and to allow the equilibrium of the reaction to lie in favor of PEP formation. Because of this reaction pyruvate may be considered the first glycolytic intermediate in gluconeogenesis in R. rubrum. The central position that pyruvate plays in carbon metabolism is thus reflected in its function as the sole activator for ADPglucose synthesis in that organism. R h o d o p s e u d o m o w capsulata, another photosynthetic anaerobe, as indicated previously has an ADPglucose pyrophosphorylase activated both by pyruvate and fructose 6-P. In contrast to R . rubrum, R. capsulata is able to grow on glucose as well as on various TCA cycle intermediates. The glucose is catabolized via the Entner-Doudoroff pathway ( 1 7 ) . The presence of the two activators for the R. capsulata pyrophosphorylase may be rationalized in that R. capsuluta utilizes a pathway for glucose degradation as well as having the ability to grow as a photosynthetic heterotroph.

+

1. The Effect of Temperature o n the Kinetics of the R. rubrum ADPglucose Pyrophosphorylase

A detailed study of the effect of temperature on the rate of ADPglucose synthesis catalyzed by the R. rubrum enzyme has been done (20).Figure 1 shows that in the absence of pyruvate a plot of In V,,, vs. 1/T deviates from linearity at temperatures above 26". The maximal velocity increases 33. B. B. Buchanan, M. C. W. Evans, and D. I. Arnon, Arch. Mikrobiol. 59, 32 (1967). 34. B. B. Buchanan and M. C. W. Evans, BBRC 2 5 484 (1985).

84

JACK PREIBS 4.01

FIG.1. Plot of the natural log of the rate of ADPglucose synthesis vs. the reciprocal of absolute temperature in the presence and absence of 20 mM pyruvate. (Adapted from Fig. 11 of reference 20; copyright (1971) by the American Chemical Society. Reprinted by permission of the copyright owner.)

to a maximum a t 35" but then decreases. Only 10% of the maximal velocity seen a t 35" is observed a t 50". I n the presence of pyruvate the curve remains linear up to 32" and the initial velocity increases with increasing temperature up to 55". The inactivation observed a t elevated temperatures is freely reversible. However, above 60" the enzyme undergoes irreversible denaturation. These results have been interpreted by Paule (20) in terms of a simple model in which the enzyme is capablc of existing in three conformational forms, one high (H) and two low temperature forms (La and Lu) with only the low temperature forms exhibiting catalytic activity; La is the enzyme form which in the prcsencc of La

(Active

forms)

(Inactive form)

pyruvate is distinguished by its p H optimum of 7.4 and an enthalpy of activation of 11.2 kcal/mole (calculated from Fig. 1) ; Lu diffcrs from La in that its pH optimum is 8.6 and its enthalpy of activation is 14.2 kcal/mole. According to the model, in the presence of pyruvate the L a form predominates over the Lu form. Pyruvate also stabilizes the activc forms of thc enzyme over the inactive (H) form. By assuming that thc H form has no catalytic activity the equilibrium constant between the high and the low temperature form of the enzyme was calculated by Paule (ZU) from Fig. 1. The plot of In keq between the high and the low

3.

ADENOSINE DIPHOSPHORTL GLUCOSE PTROPHOSPHORTLASE

85

temperature forms vs. 1/T gave linear curves with AH" and AS" values in the presence of pyruvate of 45,800 cal/mole and 147.6 eu, respectively. In the absence of pyruvate AH" was 28,500 cal/mole and AS", 86.4 eu. Thus, the data suggest that the temperature phenomenon seen in Fig. 1 results from a temperature-dependent eonformational change of the enzyme which is effected by pyruvate. The thermodynamic quantities determined by Paulc arc of thc cxpcctcd magnitude for thc suggcsted conformational change. Similar tcmpcrature-dcpendeiit eonformational changes have been observed in other systems (35-41). Studies of thc effect of temperature on the concentrations of substrate and divalent metal ion needed for maximal velocity indicate that changes in the concentration of substrate needed to achieve maximal velocity cannot account for the anomalous Arrhenius plots seen in Fig. 1. The data presented in Fig. 1 were obtained in the presence of saturating concentrations of substrate and metal ion in the presence or absence of pyruvate. The pH optimum is also constant over the temperature range; therefore, the phenomenon is probably not resulting from changes in the degree of ionization of groups on the enzyme. Paule also studied the effect of temperature on the kinetic parameters of the substrates ( 2 0 ) . A decrease in the temperature of the reaction mixture decreased the concentration of substrates and divalent metal ion needed for half-maximal velocity (So.,) as well as the Hill constant, W, for ATP. At 20" in the presence of pyruvate the So.5value for ATP was 0.12 mM and the Hill w was 1.1. However, at 50", So.6 increased to 0.78 mM and f i increased to 1.8. Thus, the ATP saturation curve was changed from a hyperbolic form to a sigmoidal form. I n the absence of pyruvate the ATP saturation curve was still sigmoidal a t 19" giving an So.5value of 1.25 m M and a Hill w of 2.0. At 46" the So., and A values increased to 4.13 m M and 4.0, respectively. Similar effects were noted for a-glucose l-P and MgC12, but the So.5values only changed about 1.5to 2.0-fold. The sigmoidicity of the MgCl, saturation curve was essentially unaffected, but in thc absence of pyruvate the a-glucose l-P curve became slightly sigmoidal at 46". Thus, higher temperature caused grcatcr coopcrativity bctween substrate sites and between activator sites. V. Massry, B. Cnrti, nnd H. Gnnther, JBC 241, 2347 (1966). G. Kdnitsky nntl H. Resnik, JBC 234, 1714 (1959). C. H. Sucltcr and W. Melandrr, JBC 238, 4108 (1963). F. J. Knynr and C. H. Siicltrr, Biochentidry 7, 1678 (1968). F. J. Kaync and C. H. Wd trr. JACS 87, 897 (1965). C. H. Suclter, R . Singlrton, F. J. Kayne, 8. Arrington, J. Glass, and A. S. Mildvan, BiochentistTy 5, 131 (1966). 41. C. H. Sucltcr, Biochemistry 6, 418 (1967). 35. 36. 37. 38. 39. 40.

86

J A C K PREISS

There appear to be similarities between the allosteric effect of pyruvate and the effects of lowered temperatures of incubation. As indicated above pyruvate decreases the concentration of the substrates ATP, MgCl,, and glucose 1-P needed for half-maximal velocity, as well as decreasing the sigmoidicity of the saturation curves of ATP, ADPglucose, and glucose 1-P. The same effects are also seen when the incubation temperature is decreased. However, in contrast to the addition of pyruvate lower incubation temperatures do not shift the pH optimum from pH 8.6 to 7.4. Pyruvate also lowers the enthalpy of activation by 3 kcal/mole (Fig. 1). Thus pyruvate must have either additional or slightly different conformational effects on the ADPglucose pyrophosphorylase of R. rubrum than those effected by lower temperature. 2. Reaction Mechanism of

R. rubruin ADPglucose Pyrophosphorylase

Since the substrate saturation curves of the R . rubrum enzyme are hyperbolic at low temperatures and in the presence of activator, initial velocity kinetic studies were done to gain insight in the reaction mechanism (42). The initial velocity studies yielded intersecting reciprocal plots indicating that the kinetic mechanism of this enzyme is sequential. Product inhibition patterns eliminated all known sequential mechanisms except the ordered Bi-Bi or Theorell-Chance mechanisms. However, some small intercept effects suggest the existence of significant concentrations of central transitory complexes. The kinetic constants obtained in the analysis also favored the ordered Bi-Bi mechanism. Adenosine triphosphate [szP]pyrophosphate isotope exchange a t equilibrium studies also supported a sequential nonrandom mechanism and, in addition, indicated that ATP is the first substrate to add and that ADPglucose is the last product to dissociate from the enzyme.

D. ADPGLUCOSE PYROPHOSPHORYLASES OF HIGHER PLANTS AND GREENALGAE 1. Spinach Leaf ADPgluwse Pyrophosphorylase All ADPglucose pyrophosphorylases of higher plants and green algae are activated by 3-phosphoglycerate (3PGA) and are inhibited by orthophosphate (61-26).Other glycolytic intermediates such as PEP, fructose1,6-diP (FDP), and fructose 6-P activate to lesser extents and a t much 42.

M.R. Paule and J. Preiss, JBC

246, 4602 (1971).

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

87

higher concentrations. The enzyme of this class studied in the greatest detail is that obtained from spinach leaf (21-23). 3-Phosphoglycerate decreases the K, of ADPglucose from 0.93 to 0.15 mM, the K , of pyrophosphate from 0.50 to 0.04 mM, and the K,,, of ATP from 0.45 to 0.04 mM. The K , value of glucose 1-P is 0.07 mM in the absence of 3PGA and is decreased to 0.04 mM in its presence. All substrate saturation curves are hyperbolic either in the presence or absence of 3PGA. The MgCl, saturation curve is sigmoidal in the presence or absence of the activator and its So.,, 1.6 mM, is not affected by 3PGA. The stimulation by 3PGA is dependent on pH and results from the different pH optima of the activated and unactivated reaction. The stimulation is also dependent on the buffer used. In the presence of 3PGA the activity of the spinach leaf pyrophosphorylase in tris-Cl, triethanolamine, glycylglycine, and imidaaole buffers is about equal with a broad optimum between p H 7 and 8. In the absence of the activator the activity varies with the different buffers. Highest activity is observed with glycylglycine buffer at pH 7.0. In imidaaole buffer the optimum

FIG.2. Effect of phosphate concentration on ADPglucose synthesis catalyzed by the spinach leaf enzyme at pH 7.5. The bottom figure is a Hill plot (28, 29) of the data.

88

JACK PREISS

is'between pH 7.5 and 8.0 with the maximal activity being only 67% of the maximal activity observed with glycylglycine buffer at pH 7.0. The pH optimum in triethanolamine and tris-Cl buffers is pH 7.0 but the maximal activity is only 50 and 33% of that observed in glycylglycine buffer, respectively. In glycylglycine buffer the synthesis of ADPglucose is activated by 3PGA 80-fold a t pH 8.5, 23-fold a t pH 8.0, ll-fold a t pH 7.5, and 9-fold a t pH 7.0 (9s). Pyrophosphorolysis of ADPglucose is stimulated by 3PGA 2.5-fold at pH 7.0, 3.8-fold at 7.5, 5-fold a t pH 8.0, and 15-fold at pH 8.5 in glycylglycine buffer (99). In the synthesis reaction the AO.Sof 3PGA is 2.0 X M a t p H 7.5. While in the pyrophosphorolysis reaction Ao., is 4 pM a t pH 7.5. It is of interest that the 3PGA curve is hyperbolic in shape a t pH 7.0 and 7.5 but becomes progressively sigmoidal as the pH increases. At pH 8.5 the Hill constant fi is 1.8. The spinach leaf ADPglucose pyrophosphorylase is very sensitive to inhibition by orthophosphate (29, W ). Adenosine diphosphoryl glucose synthesis is inhibited 50% by 22 pM Pi in the absence of activator a t pH 7.5. However, as seen with other ADPglucose pyrophosphorylases, the activator can reverse or antagonize the allosteric inhibition. In the presence of 1 mM 3PGA, 50% inhibition of ADPglucose synthesis requires 1.3 mM phosphate (Fig. 2). Thus, the activator increases the concentration of inhibitor required for 50% inhibition about 450-fold. The 3PGA saturation curve which is hyperbolic becomes sigmoidal in the presence of the inhibitor, phosphate. As shown in Fig. 3, Pi a t 0.5 mM increases Ao.5 of 3PGA from 20 to 230 rJJM and increases fi from 1.0 to 1.9. At 0.75 mM Pi, the 6 and Ao.bvalues of 3PGA are increased to 2.5 and 300 pM, respectively. Conversely, 3PGA increases the sigmoidicity of the Pi inhibition curve. Figure 2 shows that the Hill interaction coefficient A for PI in the absence of 3PGA is 1.2 and is increased to 2.9 in its presence. Thus, the inhibitor, phosphate, causes an increase in the interaction among activator sites and the presence of the activator, 3PGA, increases the interaction among the inhibitor sites. Phosphate is a noncompetitive or mixed inhibitor with respect to the substrates, ADPglucose, PP+,ATP, and glucose l-P. Under certain conditions Pi changes the hyperbolic PPi and ADPglucose saturation curves to sigmoidal shapes suggesting that there are multiple substrate sites on the enzyme which normally do not interact. Adenosine diphosphate is a poor inhibitor of the spinach leaf enzymc which in the absence of 3PGA has an value 1.2 mM. I n the presence of 1 mM 3PGA its is 2.0 mM. The ADP inhibition curve is sigmoidal with a Hill constant fi of 1.8 which is not changed by 3PGA. Conversely, the 3PGA saturation curve remains hyperbolic in the presence of ADP.

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

3-Phosphoplyceric acid (mM)

89

-

lop IJPG4

FIG.3. Reversal of PCinhibition of the spinach leaf ADPglucose pyrophosphorylase activity by 3-phosphoglycerate at pH 7.5. The bottom figure is a Hill plot (28, ,9929) of the data.

Adenosine diphosphate is a mixed type of inhibitor with ATP and a competitive inhibitor with ADPglucose. 2. Other Leaf ADPglucose Pyrophosphorylases

A number of studies indicate that the ADPglucose pyrophosphorylase of a number of leaves (butter lettuce, tobacco, tomato, barley, sorghum, maize, sugar beet, rice, peanut, and kidney bean) are activated by 3PGA and inhibited by Pi (25, 24). Table IV summarizes data indicating the concentration required for 50% of maximal activation (A0.5)and the concentrations of Pi required for 50% inhibition in the absence of activator and in the presence of 3PGA a t the concentrations indicated. As shown for the spinach leaf enzyme the inhibition by Pi is reversed by 3PGA. In addition, the hyperbolic activation curve of 3 PGA is converted to a sigmoidal form by phosphate while the hyperbolic phosphate curve is changed to a sigmoidal shape by the presence of 3PGA (24) for the enzymes listed in Table IV. The stimulation by 3PGA of ADPglucose

90

JACK PREISS

synthesis a t pH 7.5 ranges from 5.5-fold for the tomato leaf enzyme to 16-fold for the rice leaf enzyme (24). TABLE IV ACTIVATION AND INHIBITION OF LEAF ADPGLUCOSE PYROPHOSPHORYLASES

Pi I0.s

-3PGA

+3PGA

(PM)

(PM)

Concn. of 3PGA (mM)

30 80 20 190 50 60 22

1,010 880 2,300 410 430 270 1,200

1.0 2.5 1.4 2.2 0.87 1.0 1.0

3PGA Ao.6

Plant leaf

(PM ) ~~

Turkish tobacco Red cherry tomato Barley Sorghum Sugar beet Rice Spinach

45 90 7 370 190 180 20

3. Chlorella pyrenoidosa ADPglucose Pyrophosphorylase The ADPglucose pyrophosphorylase from this green alga is very similar in properties to the leaf enzymes (26).The V,,, of synthesis and of pyrophosphorolysis are stimulated 18-fold and 7-fold, respectively, by 3PGA a t pH 8.5, the optimum for both the activated and unactivated reaction. The algal enzyme is also inhibited by phosphate. The Io.6 value is 0.18 mM and the Hill f i is 1.3 in the absence of 3PGA. In the presence of 2 mM 3PGA, 10.5 and fi are increased to 1.0 mM and 1.6, respectively. Conversely, the A 0 . 5 of 3PGA was 0.4 mM and the Hill interaction coefficient fi is 1.0 in the absence of Pi. In the presence of 0.1 mM Pi, fi is increased to 1.3 and Ao.5to 0.5 mM, while in the presence of 0.5 mM Pi, A0.6 is 0.72 mM and fi is 1.6. These results are similar to that observed for the leaf enzymes in that greater interaction is seen between the inhibitor sites when activator is present and between the activator sites when inhibitor is present. The ATP and ADPglucose saturation curves of the Chlorella pyrenoidosa enzyme are sigmoidal in the presence or absence of the activator 3PGA. This result is different from that observed for the spinach leaf enzyme: 3PGA only decreases the So.6values about 1.6-fold; ATP from 0.8 to 0.5 mM, and ADPglucose from 2.8 to 1.8 mM. The ADPglucose pyrophosphorylases of Chlorella vulgaris, Scenedesmus obliquus, and Chlamydomonas reinhardii are also activated by 3PGA and inhibited by orthophosphate (26).

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

91

4. Physiological Significance of 3-Phosphoglycerate Activation

and Phosphate Inhibition of the Leaf and Algal ADPglucose Pyrophosphorylases Because of the great sensitivity of the leaf ADPglucose pyrophosphorylases to 3PGA, the primary CO, fixation product of photosynthesis, and Pi it is suggested that they play a significant role in the regulation of starch biosynthesis. The level of Pi has been shown to decrease in leaves during photosynthesis because of photophosphorylation, and glycolytic intermediates are known to increase in the chloroplast in the light. This situation would therefore contribute to conditions necessary for optimal starch synthesis via the increased rate of formation of ADPglucose. In the light the levels of ATP and reduced pyridine nucleotides are also increased leading t o the formation of sugar phosphates from 3PGA. In the dark there is an increase in phosphate concentration with concomitant decreases in the levels of glycolytic intermediates, ATP, and reduced pyridine nucleotides. This would lead to inhibition of ADPglucose synthesis and therefore starch synthesis. I n order to confirm this hypothesis, knowledge is needed of the concentrations of the various effector molecules a t the actual site of the ADPglucose pyrophosphorylase. However, no information of this sort is known, and a t present it is difficult to obtain. At best, the results obtained by workers on the concentrations of the glycolytic intermediates ( 4 3 ) , phosphate (44, &), and ATP (&) in the chloroplast qualitatively support the hypothesis of regulation of starch synthesis by 3PGA, other glycolytic intermediates, and phosphate levels. Heber and Santarius (44, 45) have shown that the concentration of Pi in the chloroplasts of spinach leaf in the dark is about 5-10 m M and decreases about 30-50% in the light. Figure 4 shows that a t these concentrations 3-phosphoglycerate can partially reverse the inhibition by phosphate. At 5 mM 3PGA there is an increase of 5-fold in the rate of ADPglucose synthesis when the phosphate concentration is decreased to 7.5 mM, and 23-fold increase is observed when the phosphate is decreased to 5 mM. Thus, under these conditions a decrease of phosphate concentration of only 30-50% in the chloroplast may cause a significant acceleration of ADPglucose synthesis and therefore of starch synthesis. 43. U. W. Heber, in “The Biochemistry of Chloroplasts” (T. W. Goodwin, ed.), Vol. 2, p. 71. Academic Press, New York, 1967. 44. U. W. Heber and K. A. Santarius, BBA 109, 380 (1965). 45. K . A. Santarius and U.W. Heber, BBA 102, 39 (1965).

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$

8

5 mM Pi

3-Phosphaglycerate, mM

FIQ.4. Reversal of PCinhibition of the spinach leaf ADPglucose pyrophosphorylase activity by 3-phosphoglycerate concentration.

Recently, Kanaeawa et al. (4) have shown in C . pyrenoidosa cells that both starch and ADPglucose synthesis occurs in the light. Starch synthesis abruptly ceases and the ADPglucose level drops to below detectable limits when the light is turned off. Uridine diphosphoryl glucose levels do not change perceptibly in the light to dark transition ( 4 ) . Adenosine diphosphoryl glucose is not detectable a t any time later in the dark despite the high steady state level of ATP and hexose phosphates. Kanaeawa et al. indicate that this observation provides strong support for the importance of the regulatory role of ADPglucose pyrophosphorylase in starch synthesis in vivo. Thus, the allosteric effects exerted by 3PGA and Pi appear to be physiologically important. Since the level of 3PGA does not appreciably change in the dark to light transition (4, 43) while the phosphate levels appear to increase in the dark and decrease in the light (44, 45) it is possible that the variation of this effector molecule is the most important control element. MacDonald and Strobe1 reported that wheat leaves infected with the fungus, Puccinia striiformis, accumulated more starch than noninfected leaves (26). They could correlate the starch accumulation with the inverse of the variation observed in Pi levels in diseased leaves during the infection process. They indicated that their data suggested that in diseased leaves the variations in the level of Pi and, to a lesser extent, variations in the level of activators of the wheat leaf ADPglucose pyrophosphorylase (3PGA, FDP, etc.) regulated the rate of starch synthesis via control of the activity of ADPglucose pyrophosphorylase.

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PTROPHOSPHORYLASE

93

5. ADPgliicose Pyrophosphoqjlases of hTonchloroph.ylloits

Plant Tissue The ADPglucose pyrophosphorylases occurring in nonphotosynthetic plant tissues [maize endosperm and embryo (46, 4 7 ) ; wheat germ, etiolated peas, and mung bean seedlings; potato tuber, carrot roots, and avocado mesocarp (23)J are qualitatively similar to the leaf cnzymes in that they are activated by 3-phosphoglycerate. The stimulation by 3PGA is 1.5-10-fold for these enzymes. The enzyme representative of this group and studied in most detail is the one isolated from maize endosperm (46). Activation of ADPglucose synthesis by 3PGA is 1.5-2-fold a t pH 7.9 and 3-4-fold a t p H 6.7. However, the A,,.5 value for 3PGA is very high (2.2 mM) compared to that of the leaf enzymes. Fructose 6-P also stimulates about 3-fold with an Ao.5 of 4.0 mM. Phosphate a t 3 mM causes 50% inhibition in the absence of activator. However, in the presence of 10 mlM 3PGA, 10 mM Pi was required for 50% inhibition; Pi also changed the 3PGA saturation curve from a hyperbolic form to a slightly sigmoidal curve. As with other ADPglucose pyrophosphorylases the activator 3PGA increases the apparent affinity of the enzyme for the substrates. Thus, the So., values for MgCl?, ATP, and glucose 1-P in the absence of 3PGA are 5.2, 0.17, and 0.10 mM, respectively. I n the presence of 10 mM 3PGA, these values are 3.2, 0.10, and 0.05 mM, respectively. Both the MgClr and ATP saturation curves are sigmoidal in the absence of 3PGA, but in the presence of 3PGA the ATP saturation curve becomes hyperbolic. The saturation curve for glucose 1-P is hyperbolic in the absence of 3PGA and also in its presence. Maize tissue also has an ADPglucose pyrophosphorylase in the embryo which is distinct from the endosperm enzyme (47,48). The embryo enzyme is more heat stable a t 60" and is more sensitive to inhibition = 0.32 mM ( 4 8 ) ] .Furthermore, 3PGA has no effect on the by Pi phosphate inhibition changing neither Io.5 nor the hyperbolic shape of the curve. The 3PGA saturation curve is sigmoidal with Ao.5being 4.2 mM and fi = 1.9. Stimulation of V,,,, is 3-fold. In contrast to all other ADPglucose pyrophosphorylases studied the activator increases SO.&of the substrates, ATP and glucose 1-P. The variation in properties of the maize enzymes as compared to the leaf enzymes may reflect differences between leaf and endosperm cells 46. D . B. Dickinson and J. Preiss, ABB 130, 119 (1969). 47. D. B. Dickinson and J. Preiss, Plant Physiol. 44, 1058 (1969). 48. J. Preiss, C. Lammel, and A. Sabmw, Plant Physiol. 47, 104 (1971).

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JACK PREISS

with respect to intracellular levels of metabolites. However, the most important phenomena in regulating starch biosynthesis in endosperm may be regulation of synthesis of the starch biosynthetic enzymes, ADPglucose pyrophosphorylase and ADPglucose :~-glucan-4-glucosyltransferase (49, 60). Starch-deficient maize mutants, shrunken-2 and britt.le-2, which have only about 10-1276 of the ADPglucose pyrophosphorylase activity observed in the normal maize endosperm, synthesize only 2530% as much starch as normal maize (47, 51). These data would suggest that the major portion, if not all, of the starch synthesized in the normal endosperm is via the ADPglucose pathway.

E. THEADPGLUCOSE PYROPHOSPHORYLASBS OF THE ENTEROBACTERIACEAE The ADPglucose pyrophosphorylases of Escherichia coli, Citrobacter freundii, Salmonella typhimurium, Escherichk aurescens, Aerobacter aerogenes, and Aerobacter cloacae are activated by the glycolytic intermediates FDP, 3PGA, PEP, 2-phosphoglyceric acid (PPGA) and 3-phosphoglyceraldehyde as well as by TPNH and pyridoxal 5-P (PLP) (1013). Of these, FDP, TPNH, and pyridoxal 5-P are the most effective activators. However, one organism of the Enterobacteria, Serratia marcescens, contains an ADPglucose pyrophosphorylase that is not appreciably activated by any metabolite tested (12). The activator sites for the enteric ADPglucose pyrophosphorylase appears to be quite nonspecific. A detailed study of the specificity of the E . coli B activator site has been undertaken and results are summarized in Table V (11, 13, 6 2 ) . Pyridoxal 5-P is the most effective activator in terms of concentration required for 50% of maximal stimulation and extent of activation. Fructose-dip stimulates to the same extent but has a 4-fold higher Ao., at pH 7.0 and ll-fold higher Ao.5at pH 8.5; TPNH has approximately the same Ao., as does fructosedip at pH 7.0 or 8.5 but stimulates ADPglucose synthesis about 75% as much. Other compounds that are effective as fructose-dip are sedoheptulose-1,7-diP, ~-arabinitol-l,5-diP,and 4-pyridoxic acid 5-P. Physiological metabolites that stimulate to lesser extents than fructose-dip and 49. C.-Y. Tsai and 0. E. Nelson, Plant Physiol. 46, 299 (1970). 50. J. L. Osbun, J. S. Hawker, E. Greenberg, C. Lammel, E. Y. C. Lee, and J. Preiss, Plant Physiql. (1973) (in press). 51. C.-Y. Tsai and 0. E. Nelson, Science 151, 341 (1986). 52. S. Govons, N . Gentner, E. Greenberg, and J. Preias, JBC (1973) (in presa) .

3.

95

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

TABLE V ADPGLUCOSE PYROPHOSPHORYLASE FROM E . coli Ba

ACTIVATORSPECIFICITY OF

THE

pH 7.0

pH 8.5

A0.6

Activator

Ir,,

None Noneb FDP PLP TPNH TPN PEP 2PGA 3PGald Sedoheptulose dip 1,5-Arabinitol dip 1,3-Glycerol dip Erythrose 4-P 4-Pyridoxic acid 5-P

2.9 20 100 110 78 42 37 47 43

(PM)

fi

-

-

68 16 64 230 260 310 380

2.0 2.7 2.0 2.3 1.8

2.0 2.0

-

-

A0.6

vmx

GM)

2.5 16 100 110 72 10 35 52 62 96 88 52 43 110

-

-

120 11 130 550 800 450 700 70 120 700 1,ooo 70

1.8 2.8 2.3 1.9 1.6 1.9 1.9 1.9 1.9 2.3 1.6 2.4

fi

The reaction mixture for measuring ADPglucose synthesis a t pH 7.0 or 8.5 contained 1.5 mM ATP, 0.5 mM glucose 1-P, 5.0 mM MgCl,, plus activator. To achieve maximal rates in the absence of activator the concentrations of ATP, glucose 1-P, and MgC12 were raised to 7.5, 1.0, and 25 mM, respectively. VmSxvalues are relative to the value obtained with FDP which is arbitrarily set a t 100. A0.6 and ii values were obtained from Hill plots. b Substrate concentrations were raised to give maximal velocity in absence of activator (see footnote a). 0

only a t considerably higher concentrations are PEP, BPGA, 3-phosphoglyceraldehyde (3PGald), 2 keto-3-deoxy-6-phosphogluconate,and TPN. Other analogs isosteric with fructose 1,6-diP which activate as effectively are 1,5-pentanediol dip, D-glucitol 1,6-diP, and xylitol 1,5-diP. Compounds similar to T P N or T P N H which give significant activation at higher concentrations than required for FDP are PRPP, 2’-PADPR, PG2’-P, and PG2’-PP. Compounds that gave no stimulation of ADPglucose synthesis or pyrophosphorolysis a t concentrations of 0.05 and 1.5 mM were 2-deoxy-~-glucose 6-P, D-fructose 1-P, D-fructose 6-P, D-glucosamine 6-P, D-glucose 6-P, P-glycerol P, D-galactose 6-P, D-ribose 5-P, 2-deoxy-~-nbose 5-P, D-sedoheptulose 7-P, dihydroxyacetone P, glycolaldehyde P, 2,3-diphosphoglycerate, 3’5’-cyclic adenylate, glyceric acid, lactic acid, 0-serine P, pyruvate, fructose, glucose, fumarate, succinate, malate, a-ketoglutarate, NaHC03, DPN, DPNH, pyridoxal, pyridoxamine 5-P, pyridoxine 5-P, and deoxypyridoxine 5‘-P. Slight

96

JACK PREISS

stimulation (10% or less of that of fructose-dip) was seen for 3-phosphoglycerate, acetyl-P, acetyl-CoA, PG3'-PP, glycerol 1,2-diP, and PPGPP [MSI (63)1. The saturation curves of the activators are sigmoidal and when plotted according to the Hill equation A values of 1.6-2.8 are obtained (Table V) . Table V also shows that the values for the activators obtained at pH 7.0 are usually about 2-fold lower than those obtained a t pH 8.5. The only exception to this is the Ao., for PLP which is 16 p M at p H 7.0 and 11 pJ4 a t pH 8.5. Although the activator specificities of the ADPglucose pyrophosphorylases from other enterics have not been studied as extensively as the E. coli B enzyme, results obtained with these enzymes indicate that the activator site specificity is very similar to the E. coli enzyme. Noteworthy is that the C . freundii enzyme is most sensitive to activation. The A0.5 values of FDP, TPNH, PLP, and glyceraldehyde 3-P are 32, 78, 7.4, and 117 p M , respectively ( 1 2 ) . The structural requirements for an activator of the E. coli enzyme may be visualized in Fig. 5. Those compounds having a structure similar 'HO

E-4-P

COO-

PLP

Pyridoxic acid-5'-P

CH,OPO,HI

c=o I

HO-C-H

H-C-OH

C&OPO,H I (CHOH),,= 1to 4 I CH,OPO,H-

I

H-C-OH I C&OPO,H-

Alditol-DIP

COOI

H-C-OH I

CH,OPO,H3 PGA

FDP

COOC-O-PO,Hti

c HE

COOI H-C-O-PO$iI CH,OH

PEP

2 PGA

FIG.5. Structures of some activators of E . coli B ADPglucose pyrophosphorylase 63. M. Cashel and B. Kahlbacher, JBC 245, 2309 (1970).

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

97

to fructose-diP (isosteric analogs) are capable of activation (e.g., sedoheptulose 1,7-diP, glycerol 1,3-diP). Some activators (e.g., PLP, erythrose 4-P, 3-phosphoglyceraldehyde, 4-pyridoxic acid 5-P, BPGA, and PEP) contain instead of two phosphate residues either a carboxyl or aldehyde residue plus one phosphate group. Thus, if basic amino acid residues, particularly lysine residues, are responsible for binding the activators it may be that the requirement for binding may be satisfied by one phosphate group plus an additional anionic or aldehydic component. An important aspect of activation may be neutralization of basic groups on the protein by the activator, and this may be achieved by phosphate and either carboxyl or aldehyde groups. Triphosphopyridine nucleotide may be considered an analog of F D P since a portion of the molecule contains a ribose 2,5-diP residue. Kinetic experiments suggest that the activators are bound to the same site (11, IS). Previous experiments indicate that 2-phosphoglycerate shares common sites with F D P (11).At subssturating concentrations of PLP, both TPNH and F D P can stimulate ADPglucose synthesis ( I S ) , but at saturating concentrations of PLP, addition of TPNH or FDP does not stimulate and may inhibit. Furthermore, the addition of FDP or TPNH decreases the sigmoidal nature of the P L P activation curve ( I S ) . These results are consistent with the view that FDP, TPNH, and PLP all bind to the same sites. 1. Effect of Activators on Substrate Kinetic Parameters

The effects of FDP, TPNH, and PLP on the kinetic parameters of the enzymes from C. freundii (12),E. coli (11, 5 2 ) , and S. typhimurium (12) have been studied. As with other ADPglucose pyrophosphorylases the activators stimulate the V,,, of synthesis and of pyrophosphorolysis and increase the apparent affinity (decrease So.5) for the substrates and for the metal ion. Table VI shows results obtained with the E . coZi B enzyme (11, 5 2 ) . The activators increase the apparent affinity of the enzyme for the substrates ATP, ADPglucose, pyrophosphate, and glucose l-P anywhere from 3- to 20-fold (Table VI). Of interest is that in the presence of the activators FDP and TPNH or in the absence of activator both ADPglucose and ATP saturation curves show cooperative effects while with PLP as the activator, the ATP curve is hyperbolic. The pyrophosphate saturation curve is hyperbolic in the absence or presence of activator, while glucose l-P saturation curves are hyperbolic in the presence of activator and exhibit negative cooperativity (54, 56) in the absence of activator. (Hill plots of the data indicate an f i value of 1.0 for the glucose l-P curve in the presence of activator and 0.6-0.8 in the absence of

98

JACK PREISS

TABLE V I EFFECTOF ACTIVATORS ON KINETICPARAMETERS OF THE E. coli B ADPULUCOSE PYROPHOSPHORYLASE~ pH 7.0

pH 8.5

Activator (mM)

(mM)

Ti

(mM)

Ti

a-Glucose-1-P

None FDP, 1.5 PLP, 0.05 TPNH, 1.5

0.16 0.033 0.040 0.033

0.8 1.0 1.0 1.0

0.12 0.036 0.037 0.030

0.6 1.0 1.o 1.0

ATP

None FDP, 1.5 PLP, 0.05 TPNH, 1.5

2.9 0.39 0.14 0.29

2.0 1.8 1.0 1.5

1.3 0.38 0.13 0.27

1.8 1.8 1.0 1.5

MgCll (synthesis)

None FDP, 1.5 PLP, 0.05 TPNH, 1.5

ADPglucose

None FDP, 1.5 PLP, 0.05 TPNH, 1.5

0.9 0.11 0.06 0.14

None FDP, 1.5 PLP, 0.05 TPNH, 1.5

0.61 0.11 0.10 0.12

Substrate

Pyrophosphate

S0.S

SO.6

11.5 2.0 1.3 2.1

3.9 4.6 4.9 4.7

12.3 2.3 1.6 1.8

2.2 1.9 1.4 1.3

-

-

-

-

-

-

1.0

-

-

1.1

-

1.0 0.94

-

-

4.5 4.8 7.3 5.5

4 T% is the Hill constant and 50.6 represents the concentration of substrate or metal ion required for half of maximal activity. Both were obtained from Hill plots.

activator.) The reciprocal plot obtained in the absence of activator is that expected for a saturation curve exhibiting negative cooperativity (64, 66). The So.Band r?, values obtained a t pH 7.0 and 8.5 for all substrates are essentially the same except for &.5 of ATP in the absence of activator. At pH 8.5 8 0 . 5 for ATP is 2.9 mM and is decreased to 1.3 mM a t pH 7.0.Thus, these kinetic studies reveal very complex interactions between activator sites and the catalytic sites; the nature of the satura54. A. Levitzki and D. E. Koshland, Jr., in “Metabolic Regulation and Enzyme Action” (A. Sols and S. Grisolia, eds.), p. 271. Academic Press, New York, 1970. 55. A. Levitzki and D. E. Koshland, Jr., Proc. Nut. Acud. Sci. U . S. 62, 1121 (1969).

3. ADENOSINE

99

DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

tion curves for substrate not only changes on addition of activator but also depends on the particular activator used. 2. Znteraction of Activator and Inhibitor The E . coli B, C . freundii, and S. typhimurium ADPglucose pyrophosphorylases are inhibited by 5'-adenylate and to lesser extents by ADP and Pi (11-13, 52, 56, 57). The sensitivity to inhibition is modulated by the concentration of activator (12, 13, 52, 56, 57). Increasing the concentration of activator makes the enzyme more resistant to 5'-adenylate. This is shown for the E. coli B enzyme in Table VII. At 34 pM FDP, the enzyme is most sensitive to 5'-adenylate; 3.2 CJM 5'-AMP gives 50% inhibition If the FDP concentration is progressively increased the concentration of 5'-AMP required for 50% inhibition also increases. At 1.7 mM FDP the Io.6 for $-AMP is 70 pM. Table VII also shows that if the fructose-dip concentration is decreased below 34 pM, the E . coli B enzyme becomes more resistant to inhibition in two ways. Higher TABLE VII AMP INHIBITION OF ADPGLUCOSE SYNTHESIS CATALYZED BY THE E . coli ADPGLUCOSE PYROPHOSPHORYLASE AT DIFFERENT LEVELS OF TEE ACTIVATOR, FDP

V Oresistant to AMP FDP (mM)

Relative

vo

(mM)

1.70 0.56 0.17 0.056 0.034 0.011 0

100 98 65 35 18.5 5 3.8 14

0.07

1.75

0.03 0.011 0.0034 0.0032 0.0073 0.03 0.35

1.65 1.48 1.37 1.25 0.93 0.68 0.50

00

inhibition

10.6

?i

(%I

2.0 3.8 7.0 24 30 55

a Reaction mixtures (pH 8.5) containing no FDP and where the concentration of ATP, glucose 1-P, and MgCla have been raised to 7.5,1.0,and 25 mM, respectively, to provide for conditions of maximal activity in the absence of activator. The concentration of ATP, glucose 1-P, and MgClt in the other reaction mixtures are 1.5, 0.5,and 5 mM, respectively. V Ois the reaction velocity in the absence of inhibitor and is relative to the velocity obtained a t saturating FDP concentration which is arbitrarily set as 100. l 0 . 6 is the concentration of inhibitor giving 50% inhibition, and It is the Hill constant.

56. N. Gentner and J. Preiss, BBRC 27, 417 (1967). 57. N. Gentner and 3. Preiss, JBC 243, 5882 (1968).

100

JACK PREISS

I,,.5 values for 5'-AMP are observed, and part of the enzymic activity becomes resistant to inhibition. In the absence of FDP about 30% of the activity is not inhibited even by 5.0 mM AMP. If the concentrations of ATP, glucose 1-P, and MgCl, arc raised to givc maximal activity in the absence of activator then 55% of the activity is resistant to inhibition. At high concentrations of fructose-dip the 5'-adenylate inhibition curve is sigmoidal, but as the FDP concentration decreases the inhibition curve becomes less sigmoidal, as indicated by a decrease in A. Finally, a t low concentrations of FDP or in its absence, the 5'-AMP curve Hill A value becomes less than one, suggesting that the 5'-adenylate sites arc interacting in a noncooperative manner. The binding of inhibitor under these conditions would hinder the binding of another molecule of inhibitor to the enzyme. This negative cooperativity of AMP sites would explain the A values of the inhibition curves being less than one and the percentage of activity not subject to inhibition. The response of the E. coli B enzyme to 5'-AMP inhibition in the presence of the activators T P N H and P L P is the same as ohserved for fructose-dip (62).Increasing the concentrations of TPNH or PLP makes the enzyme less sensitive to inhibition by 5'-AMP. This interaction between the inhibitor and activators is also seen for the C . freundii and S. typhimurium ADPglucose pyrophosphorylases (12). As indicated above, both ADP and Pi are also inhibitors of the ADPglucose pyrophosphorylase of the Enterobacteriaceae. Table VIII shows the values of ADP and Pi compared to 5'-AMP for three enteric ADPglucose pyrophosphorylases. The concentration of activator, in this case, FDP, modulates the sensitivities of the E . coli B and S. typhimurium enzymes to inhibition by ADP or Pi. TABLE VIII KINETIC CONSTANTS FOR INHIBITORS OF THE ENTERIC ADPQLUCOSE PYROPHOSPHORYLASES 10.6

Source of enzyme

E . wli B S. typhimurium

C. jreundii a

Activator (mM)

5'-AMP

ADP (mM)

Pi

VO"

FDP, 1.7 FDP, 0.15 FDP, 1 . 0 FDP, 0.25 FDP, 0.10

0.07 0.008 0.11 0.028 0.022

1.2 0.21 1.2 0.42 0.33

1.7 0.54 1.7 0.93 1.8

100 55 100 80 100

As defined in Table VII.

Relative

3.

101

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

3. Effect of Inhibitors on the Substrate and Activator Kinetic Constants

The effect of AMP on the substrate and activator saturation curves has been studied with the E . coli B ADPglucose pyrophosphorylase and with F D P as the activator (57). Figure 6 shows the effect of the presence of B'-AMP on the FDP saturation curve. Adenosine monophosphate increases the concentration of FDP required for half-maximal activation. The percent inhibition caused by a given concentration of AMP is greater a t subsaturating levels of F D P than a t saturating concentrations which is consistent with the data shown in Table VII. The rate with saturating F D P is decreased about 60% by 0.092 mM AMP; but in the range of 0-0.5 mM FDP, this amount of AMP causes nearly complete inhibition of the rate. FDP only partially reverses the inhibition caused by AMP. In addition, the f i value for F D P increases in the presence of AMP. Figure 7 shows that AMP also increases the concentration of ATP required to yield half-maximal velocity. A greater percentage inhibition is seen a t nonsaturating ATP concentrations than a t saturating concentrations. Adenosine monophosphate inhibition also increases the concentration of Mg2+required for 50% of maximal stimulation, but maximal velocity either in the presence or absence of inhibitor was always reached in the range of 5-8 mM Mg2+.$-Adenosine monophosphate as well as ADP and Pi are noncompetitive inhibitors of the substrate glucose-1-P (57) ;

A

[AMP]. mM Ao,5, mM - .MC

0.046

0.76

-n 2.8

FDP concentration, mM

F I ~ 6. . The effect of fructose-dip concentration of the inhibition of E . coli B ADPglucose pyrophosphorylase by 5'-adenylate.

102

JACK PREISS 20

-

-

No AMP

O.WmM AMP

0.092mMAMP

I\

AMP. mM

s,aATP

mM --

~

0.25 0.62 0.86

0 0.046

0.092 I

0

1.0

I

I

* . . ....

I

.

2.0

.

.. .

,...

I

.

?i 2.2 2.2 2.5

.

30

ATP, concn., m M

FIG.7. The effect of 5’-adenylate on the ATP saturation curve of E . coli B ADPglucose pyrophosphorylase. The experiment was done in the presence of 1.5 mM fructosedip. (Adapted from Fig. 11 of reference 67; Biosynthesis of Bacterial Glycogen VI. By Norman Gentner and Jack Preiss.)

ADP and Pi also increase the Ao., value for FDP and the So.5value for ATP when present in the reaction mixture (N. Gentner and J. Preiss, unpublished results). An interesting effect of 5’-AMP is on the shape of the ATP saturation curve in the presence of PLP. The ATP saturation curve is hyperbolic with P L P as the activator but changes to a sigmoidal curve in the presence of AMP. Thus, the Hill fi is increased from 1.1 to 2.0 in the presence of 15 5’-AMP; A!%.~of ATP is increased from 0.12 to 0.43 mM. These types of studies have not been done with the C. freundii and S. typhimurium enzymes. However, the data available indicate the same effects will be found. 4. Effect of Mn2+on the ADPglucose Pyrophosphorylase Activity of

E . coli B A divalent cation is required for the synthesis or pyrophosphorolysis of ADPglucose; Co2+and Mn2+were found to be able to replace Mg2+as the divalent cation in these reactions (11 ) . I n pyrophosphorolysis Co2+ and Mn2+were not as effective as Mg2+with the E . coli enzyme ( l l ) ,but the maximal rates of ADPglucose synthesis in the presence of Mgz+ or

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

103

Mn2+were about the same. Activity in the presence of Co2+was one-third that observed with Mg2+.At lower concentration Mn2+was more effective than Mg2+.Furthermore, maximal activity with Mgz+was observed when the concentration of Mg2+was about 2-3-fold in excess of the ATP concentration ( 5 4 ) .For Mn2+,maximal activity was always reached at a 1 :1 ratio between ATP concentration and Mn2+concentration. Other studies have revealed that the kinetics of ADPglucose synthesis in the presence of Mn2+were different frcjm the kinetics of the reaction carried out in the presence of Mn2+. At pH 8.5 and with 1.5 mM [MnATP]2- the FDP and T P N H saturation curves are hyperbolic, each giving Hill fi values of 1.05 and giving A,,.5 values of 29 and 36 p i l l , respectively. This is in contrast to the results observed in Table V where in the presence of MgZ+,the activation curves of FDP and T P N are sigmoidal and the Ao.5values obtained were 3- to 4-fold higher. I n the presence of MnZ+the PLP saturation curve was also less sigmoidal than that obtained in the presence of Mgz+ (Table V ) . With Mn2+the Ao.5value is 7 p M and the Hill fi value is 1.6. Thus, the activator sites interact to a much lesser extent in the presence of Mn2+.I n the presence of the inhibitor 5'-AMP however, the hyperbolic fructose-dip activation curve is changed to a sigmoidal form (67). Regardless of whether Mgz+or MnZ+serves as the cation, interactions between activators and the inhibitors AMP, Pi, and ADP were found to be such that the sensitivity of the enzyme to inhibition was modulated by the activator concentration ( 6 7 ) . However, as indicated previously, with Mg2+,the I$ values of the Hill plots for the inhibitors progressively decreased with decreasing concentrations of F D P indicating a lessening degree of cooperative interaction between sites binding inhibitor. The enzyme is also relatively insensitive to these inhibitors in the absence of activator. In contrast, when MnZ+serves as the divalent cation, the TI values for the inhibitors appear to remain constant with decreasing FDP concentration and the rate in the absence of activator is quite sensitive to these inhibitors. Thus, the inhibitor and activators sites respond differently in the presence of Mn2+or Mg2+.Table IX summarizes some differences between the Mg2+form of the ADPglucose pyrophosphorylase and the MnZ+form of the enzyme. At subsaturating concentrations of Mg2+,MnZ+enhances the activity of the enzyme; a t subsaturating concentrations of Mn2+,Mg2+can enhance the activity. Presumably, the concentration of free Mg2+ is greatly in excess of that of Mn2+in the E. coli cell. It is likely that Mg2+ is the physiologically important divalent cation of the ADPglucose pyrophosphorylase.

104

J A C K PREISS

TABLE IX SUMMARY OF PROPERTIES BETWEEN THE M G ~ AND + THE MN3+ ACTIVATED FORMS OF ADPGLUCOSE PYROPHOSPHORYLA~E FROM E. coli Divalent cation Kinetic study

Mg*+

Metal ion:ATP ratio at maximal activity

Variable

Rate vs. FDP concentration curve

Sigmoidal (ii = 2)

Mn'+ 1:1, in presence or absence

of FDP

Sensitivity to inhibitors in Relatively insensitive absence of FDP

Hyperbolic in absence of inhibitors fi > 2 in presence of inhibitors Relatively sensitive

Sensitivity to inhibitors in More sensitive at subsatu- More sensitive a t subsatupresence of FDP rating FDP concentrarating FDP concentrations; ii value for inhibition; ii value for inhibitor largely unaffected by tor decreases with concentration of FDP decreasing FDP concentrations Interaction between metal Presence of Mn*+increases Presence of Mg*+ increases the apparent affinity of ions the apparent affinity of enzyme for Mg*+,i.e., enzyme for Mn*+,i.e., Mn*+enhances activity Mg*+enhances activity a t at low Mg*+ concentralow Mns+ concentrations tions

5. Response of the E. coli ADPglucose Pyrophosphorylase to Energy Charge The allosteric activators of E. coli ADPglucose pyrophosphorylase have the following effects: ( 1 ) they increase V,,,, of ADPglucose synthesis (4-8-fold), (2) they decrease the values for all of the substrates and metal ion (2-15-fold), (3) they modify the shape of the saturation curve for glucose 1-P, (4) they modulate the sensitivity of the enzyme to inhibition, and (5) their presence allows the enzyme to become fully inhibited by 5'-adenylate. This resistance of the enzyme to inhibition, however, is only seen a t very low and nonphysiological concentrations of FDY and is probably not indicative of the physiologica'l function of the activator. Thus, the presence of activator makes the enzyme more active via changes 1 through 4, resulting in the increase of activity a t low concentrations of substrates and metal ions. I n the presence of activator the rate of ADPglucose synthesis can be increased by 20- to 40-fold under certain

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

105

conditions. In the presence of inhibitor the variation in activity with increasing activator concentration can even be greater. This complex interaction of substrates of the ADPglucose pyrophosphorylase and various effector molecules has been studied as a function of energy charge by Shen and Atkinson (58).The energy charge is defined by Atkinson and Walton (59) as [ATP] + Yz[ADP]/[ATP] + [ADP] +[AMP]. The concept of energy charge has been reviewed by Atkinson in a number of papers and reviews (6‘0-62). Suffice it to say here that regulatory enzymes from pathways involved in the synthesis of ATP are highly active at low levels of energy charge but decrease sharply in activity a t high energy charge levels. In contrast, regulatory enzymes from biosynthetic pathways that utilize ATP are relatively inactive at low energy charge levels and their activities increase dramatically at high energy charge levels. In Fig. 8 the response of the ADPglucose pyrophosphorylase to energy charge can be seen. At an energy charge of 0.7 the E . coli B enzyme has little activity with either FDP, TPNH, or P L P as activator. This is essentially the same result as that reported by Shen and Atkinson (58). At an energy charge of 0.8, which is considered to be the physiological

r c ..->

100

-

80

--

60

--

-

0.5

0.6

0.7

0.8

0.9

1.0

Energy charge

FIG. 8. The response of E . coli B ADPglucose pyrophosphorylase activity to energy charge in the presence of activators at pH 7.0: FDP, 1.5 mM, TPNH, 1.5 mM, and PLP 50 p M . 58. 59. 60. 61. 62.

L. C. Shen and D. E. Atkinson, JBC 245, 3996 (1970). D. E. Atkinson and G. M. Walton, JBC 242, 3239 (1967). D. E. Atkinson, Biochemistry 7, 4030 (1968). D. E. Atkinson, Annu. Rev. Microbiol. 23, 47 (1969). D. E. Atkinson, “The Enzymes,” 3rd ed., Vol. 1, p. 461, 1970.

106

JACK PRFJSS

value for E . coli (63),little activity is seen with either T P N H or PLP as activator, but the activity seen with F D P has increased sharply and is about 18% of the activity observed a t an energy charge of 1.0. Further increases in the energy charge result in dramatic increases in the activity of the ADPglucose pyrophosphorylase. Thus, the E . coli B ADPglucose pyrophosphorylase activity is very sensitive to energy charge and its response is consistent with its function in the biosynthesis of glycogen, an energy storage compound. This sharp response to energy charge has been shown by Shen and Atkinson to be modulated by the concentration of activator (68). Shen and Atkinson have also studied the effect of activator concentration on the activity of the ADPglucose pyrophosphorylase a t an energy charge of 0.85. In this case only FDP gave significant activation of the E. coli B enzyme. Very little activation was noted with T P N H and other glycolytic intermediates, suggesting that FDP is the important physiological activator. Unpublished experiments from our laboratory indicate that PLP also activates at an energy charge of 0.85, but the concentrations required are much greater than those observed under physiological conditions. Shen and Atkinson (58) noted that the E. coli B ADPglucose pyrophosphorylase exhibits the sharpest energy charge response of any known enzyme in an ATP utilizing (biosynthetic) sequence. They suggested that this response is consistent with the participation of ADPglucose pyrophosphorylase in the synthesis of an energy storage compound, glycogen. Enzymes functioning in the synthesis of energy storage compounds would perhaps only function a t higher energy charges than those enzymes participating in the synthesis of macromolecules needed for growth. During conditions where growth occurs, macromolecule biosynthesis would proceed, while the ADPglucose pyrophosphorylase would be inactive. Where growth ceases as a result of limitation of nutrients other than a carbon source, the ADPglucose pyrophosphorylase would now be able to compete because feedback inhibition of the biosynthetic sequences would decrease the biosynthetic enzymic activities. Moreover, the accumulation of glycolytic intermediates would result in an increase in the activity of the ADPglucose pyrophosphorylase. I n the presence of excess carbon nutrient or energy source during conditions of nongrowth the energy charge would also be expected to increase slightly, thus resulting in a significant increase in the ADPglucose pyrophosphorylase activity. The activator and inhibitor interactions observed in the kinetic studies, and the response of the ADPglucose pyrophosphorylases to energy charge and modulation of its response to energy charge by activator, strongly 63. A. G. Chapman, L. Fall, and D. E. Atkinson, J . Bactem'ol. 108, 1072 (1971).

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

107

suggest that ADPglucose and glycogen syntheses are controlled by the concentrations of activators and inhibitors in the bacterial cell.

F. THEADPGLUCOSE PYROPHOSPHORYLASE OF Serratia marcescens Unlike other enteric bacteria, S. marcescens has an ADPglucose pyrophosphorylase which does not appear to be activated by a known metabolite. Compounds such as FDP, TPNH, PLP, 3PGA, fructose 6-P., pyruvate, or ribose 5-P do not activate more than 20%. They also do not reverse to any extent the inhibition caused by 5’-adenylate or ADP. Phosphoenolpyruvate stimulates ADPglucose synthesis about 60% with an Ao.5of 0.75 mM. However, unlike the activators seen for other ADPglucose pyrophosphorylases, PEP does not lower the for any of the substrates. The 8 0 . 5 for glucose l-P is 0.049 m M in the absence of PEP and of ATP in the 0.059 mM in the presence of 2.3 mM PEP, while the presence or absence of PEP is 0.3 mM. Also, PEP does not reverse or antagonize the inhibition caused by 5’-AMP or ADP. The low activation by PEP occurs only a t moderately high concentrations compared t o the activators of other ADPglucose pyrophosphorylases. Whether PEP activation is significant in vivo is therefore doubtful. It is noteworthy that the 8 0 . 5 values of ATP and glucose l-P are the same as those observed for the other enteric bacterial ADPglucose pyrophosphorylases in the presence of their activators. The ATP and ADPglucose saturation curves are sigmoidal in shape giving Hill ri. values of 1.6 and 1.8, respectively, indicating interaction of these substrates sites. The glucose l-P saturation curve is hyperbolic. 5’-Adenosine monophosphate is a potent inhibitor of the S. marctscens enzyme with an 1 0 . 5 of about 15 &, while ADP is less effective with Io.6 being 230 Both inhibition curves are hyperbolic, in contrast to what is observed with the ADPglucose pyrophosphorylases of E . coli B, S . typhimurium, and C. freundii. Phosphate is a poor inhibitor of ADPglucose synthesis with an value of 8.7 mM. Although no interaction between PEP and the inhibitor 5‘-AMP is observed, kinetic studies indicate some interaction between the substrate ATP and 5’-AMP. I n the presence of 5 pM 5’-AMP the s 0 . 5 for ATP increases from 0.30 to 0.75 mM. The sigmoidicity of the ATP curve is also increased (Hill A from 1.7 to 2.0). Results shown in Table X are in agreement with this. A decrease in the concentration of ATP increases the sensitivity of the enzyme 5’-AMP inhibition as indicated by a decrease in the Io.6 value. Thus, control of ADPglucose synthesis in S. marcescens appears to be regulated solely by the energy charge of the cell or by the ATP-AMP ratio.

a.

108

JACK PREISS

TABLE X OF 5'-hENYLATE INHIBITION OF THE s. WUWCeSCenS MODULATION ADPQLUCOSE PYROPHOSPHORYLASE BY ATP CONCENTRATIONS ATP concn. (mM) 2.5 1.0

0.25

1 0 . 6 AMP

VO"

(PM)

100 76 31

15 8.8 3.3

V Ois the velocity obtained in the absence of 5'denylate and is relative to the velocity obtained with 2.5 m M ATP, which is arbitrarily set at 100. (I

G. THEADPGLUCOSE PYROPHOSPHORTLASE OF Aeromonas formicans The aeromonads are generally classified taxonomically as intermediate between the pseudomonads and enteric organisms since they possess some of the characteristics of each of these two groups (64, 6 5 ) . Morphologically they resemble many of the members of the genus Pseudomonas. They ferment various carbohydrates to yield end products similar to those found in carbohydrate fermentation by enteric organisms (66, 67). Aeromonads give a strong positive test for cytochrome oxidase, a property found in most pseudomonads but absent in enteric bacteria. The guanosine-cytosine content of aeromonad DNA ranges from 58 to 62% (68))higher than most enteric bacteria (69) and lower than most of the pseudomonads (70).Recent studies by Crawford and his associates have shown that Aerornonas formicans contains a P-galactosidase similar to E. coli (71).Pseudomonads do not produce P-galactosidase. Moreover, the genetic regulation and the nature of the enzymes involved in tryptophan biosynthesis in A . formicans are very similar to that found in E . coli and S . typhimurium and unlike those found in the pseudomonads (72). Thus, Crawford et al. (72)have speculated that the arrangement of the chromosome of the aeromonads would be more homologous to the enteric bacteria than to the pseudomonads. Glycogen and the enzymes involved in glycogen biosynthesis, glycogen synthetase, and ADPglucose pyro64. R. R. Colwell and J. Liston, J. Bacteriol. 8.2, 1 (1961). 65. 0. Lysenko, J . Gen. Microbiol. 25, 379 (1961). 66. I. P. Crawford, J . Bacteriol. 68, 374 (1954). 67. R. Y. Stanier and G . A. Adams, BJ 38, 168 (1944). 68. M. Sebald and M. Veron, Ann. Znst. Pasteur, Park 105, 897 (1963). 69. R. R. Colwell and M. Mandel, J. Bacleriol. 87, 1412 (1964). 70. M. Mandel, J. Gen. Microbiol. 43, 273 (1963). 71. S. R. Rohlfing and I. P. Crawford, J. Bacteriol. 91, 1085 (1966). 72. I. P. Crawford, S. Sikes, and D. K . Melhorn, Arch. Mikrobiol. 59, 72 (1967).

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ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

109

phosphorylase have been found in A. formicans. Glycogen is a storage compound found in the enterics but not in the pseudomonads. The A . formicans ADPglucose pyrophosphorylase resembles the enteric enzyme faintly in that it is activated by FDP. However, PLP and T P N H are not activators of the Aeromonas enzyme. I n addition, fructose 6-P is a slightly better activator than FDP for the Aeromonas pyrophosphorylase while fructose 6-P is completely inactive as an activator for the enteric ADPglucose pyrophosphorylases. Fructose 6-P and FDP increase maximal velocity 2.2- and 1.6-fold, respectively, and the A0.5of fructose 6-P is about 13 pM. Thus, the enzyme is quite sensitive to activation by fructose 6-P. Fructose 6-P also lowers the So.5values of the substrates 2-4-fold. As with many other ADPglucose pyrophosphorylases the ATP and ADPglucose binding sites show cooperative interaction. Both the ATP and ADPglucose saturation curves are sigmoidal with Hill f i values of 1.6 and 1.5, respectively. Adenosine diphosphate is the most effective inhibitor of the Aerornonas enzyme, but it is much less effective than inhibitors of other classes of ADPglucose pyrophosphorylases; Io.5 for ADP is 3.8 mM in the absence of fructose 6-P and 5.0 mM in the presence of 0.5 mM fructose 6-P. Interaction between the fructose 6-P sites and ADP sites is also seen when the effect of ADP on the fructose 6-P activation curve is studied. I n the presence of 5 mM ADP, the Ao.5value of F6P is increased from 13 p i l l to 1.0 mM. However, F6P never completely reverses the ADP inhibition. The interaction between the inhibitor and activator of the Aerornonas enzyme has not been studied in great detail, and further studies are required to determine the nature of the ADP inhibition. The possibility that ADP inhibits in part as a result of its binding of the Mg2+ion necessary for the catalytic reaction has not been ruled out. Nevertheless, the available data clearly indicate that the Aeromonas enzyme is distinct from the other classes of ADPglucose pyrophosphorylases.

H. THEKINETICPROPERTIES OF ADPGLUCOSE PYROPHOSPHORYLASES ISOLATED FROM E. coli B MUTANTS ALTERED IN THEIR ABILITYTO ACCUMULATE GLYCOGEN The most significant property of the various classes of ADPglucose pyrophosphorylases is the kinetic interaction between inhibitor and activator effector molecules. This suggests that ADPglucose synthesis (and therefore glycogen synthesis) is regulated in vivo by the concentrations of inhibitor and activator in the cell. Strong support for this suggestion has been obtained by the study of a number of E. coli mutants that are

110

JACK PREISS

altered in their ability to accumulate glycogen as compared to the parent strain and which are found to contain ADPglucose pyrophosphorylases altered in their regulatory properties (27, 62, 73-78). The kinetic properties of these mutants are described in the following sections. 1. E. coli B Mutants SG6 and CL1136’

Table XI compares the titers of the glycogen biosynthetic enzymes, ADPglucose pyrophosphorylase and ADPglucose :(Y-1,kglucan-4-a-glucosyltransferase (glycogen synthetase) , the rates of accumulation of glycogen, and the maximum amount of glycogen accumulated by the ,

TABLE XI GLYCOGEN ACCUMULATION IN E . COzi B AND MUTANTSSG5

.4ND

CL1136.1

Glycogen E. wli strain

Medium

Rate (pmoles/ Accumulation g-hr) (mg/g)

ADPglucose pyrophosphorylase (ccmoles/g-hr)

Glycogen synthetase (rmoles/ g-hr)

B

Enriched minimal

23 32

20 20

292 326

442 517

SG5

Enriched minimal

48 59

53 35

267 313

348 408

CL1136

Enriched minimal

77 114

57 74

320 279

444 658

The enriched and minimal media with glucose as a carbon source and the assays for glycogen accumulation are described in reference (75). Accumulation of glycogen is expressed aa milligram of anhydroglucose per gram (wet weight) of cells, and the value given is the maximal amount accumulated in stationary phase. The rate of glycogen accumulation is expressed as the change of micromoles of anhydroglucose per gram of cells (wet weight) per hour. ADPglucose pyrophosphorylase activity was measured in crude extracts of cells prepared as described previously (11) in the presence of 1.5 mM FDP. This concentration of activator gave optimal rates of enzymic activity in extracts from all organisms. 73. S. Govons, R. Vinopal, J. Ingraham, and J. Preiss, J . Bacterial. 97, 970 (1969). 74. J . Cattaneo, M. Damotte, N . Sigal, F. Sanchez-Medina, and J. Puig, BBRC 34, 694 (1969). 75. J. Preiss, 5. Govons, L. Eidels, C. Lammel, E. Greenberg, P. Edelmann, and A. Sabraw, in “Miami Winter Symposia” (W. J. Whelan, ed.), Vol. 1, p. 122. NorthHolland Publ., Amsterdam, 1970. 76. J. Preiss, in “The Biochemistry of the Glycosidic Linkage” (R. Piras and H. G. Pontis, eds.), p. 517. Academic Press, New York, 1972. 77. J. Preiss, Zntru-Sn’. Chem. Rep. (1973) (in press). 78. J. Preiss, A. Sabraw, and E. Greenberg, BBRC 42, 180 (1971).

3.

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ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

E . co2i B mutants SG5 and CL1136 with the parent strain. The maximum amount of glycogen accumulated in SG5 and CL1136 is about 3-fold more than that accumulated by the parent strain in enriched media containing 1% glucose. In minimal media containing 0.6% glucose the amount of glycogen accumulated in SG5 and CL1136 is 2- and 4-fold greater, respectively, than that found in E . coli B. The rates of glycogen synthesis in both media are about 2- and 3.5-fold greater for the mutants SG5 and CL1136, respectively. The levels of activities present in the three organisms are about equal and therefore cannot account for the increased rate of accumulation of glycogen present in the mutants. Figure 9 shows that the apparent affinities of the mutant ADPglucose

A :

0

025

05

015

10

"I 5

Fructose-DIP, mM

IAl

CL1136

FDP, p M

A , . 0025

0075

00 5

01 '02505

Fructose-DIP, mM

(BI

FIG.9. FructosediP activation curves for E . coli B (A), SG5 (A), and CL1136 (B) ADPglucose pyrophosphorylases at pH 7.0. The reaction mixtures contained 0.5 mM glucose 1-P, 1.5 mM ATP, 5.0 mM MgCL and FDP as indicated in the figure.

112

JACK PREISS

TABLE XI1 KINETIC CONSTANTS OF FDP, TPNH, AND PLP FOR THE E . wla B MUTANT ADPQLUCOSE PYROPHOSPHORYLASES AT PH 7.0

AND

Ao.s Hill constant FDP Enzymesource

E . wli B SG5 CL1136

ii

-

TPNH

PLP

(pM)

(pM)

(pM)

FDP

TPNH

PLP

68 22 5.2

64 31 5.0

16 7 0.9

2.0 1.8 1.0

2.0 2.1 1.0

2.7 1.7 1.15

pyrophosphorylases for the activator FDP are greater than that of the parent strain enzyme. The concentration of FDP required for 50% of maximal activation (AO.B)is about 3-fold less for the SG5 ADPglucose pyrophosphorylase and 12-fold less for the CL1136 enzyme. At 1.5 mM F D P there is a 34-fold stimulation of ADPglucose synthesis catalyzed by the E. coli enzyme but only 14-fold with the SG5 enzyme and 1.5-fold for the mutant CL1136 enzyme. Thus, ADPglucose synthesis catalyzed by the parent strain enzyme is more dependent on FDP than the ADPglucose synthesis catalyzed by the mutant enzymes. The CL1136 ADPglucose pyrophosphorylase is almost fully active in the absence of FDP. Table XI1 shows that the apparent affinities for the activators T P N H and PLP are also greater with the mutant ADPglucose pyrophosphorylases than with the E . coli B enzyme. The shapes of the activation curves for SG5 are slightly less sigmoidal than those obtained with the parent enzyme as indicated by the lower A values. The shapes of the activation curves for the CL1136 enzyme however are hyperbolic (6 = 1) and considerably different from those observed with the E. coli B enzyme. Thus, the cooperativity of the activator binding sites appear to be reduced with the SG5 enzyme and completely lost with the CL1136 ADPglucose pyrophosphorylase. 2. Inhibition of the SGB and CL1136‘ ADPglucose Pyrophosphorylases

The ADPglucose pyrophosphorylases of SG5 and CL1136 are less sensitive to 5‘-adenylate inhibition than the parent enzyme since higher concentrations of 5’-adenylate are required to give 50% inhibition (Table X I I I ) . All enzymes become more sensitive to 5’-AMP when the fructosedip concentration is decreased. Between 0.15 and 1.5 mM FDP, the SG5 enzyme requires about 1.7-fold more 5’-AMP than does the E. coli B enzyme for 50% inhibition while 6-10-fold higher 5’-adenylate

3.

113

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

TABLE XI11 KINETIC CONSTANTS OF BI-ADENYLATE WITH THE E . W l i B AND MUTANT ADPGLUCOSE PYROPHOSPHORYLASES AT P H 7.0

E . wli B

1.5 0.5

0.15 0

100 100 82 2.9

105 41 16

-

SG5

1.7 1.6 1.7

-

100 100 100 7.3

170 74 29 -

CL1136

1.5 1.9 1.4 -

100 100 100 67

680 380 142 7.8

2.25

1.8 1.7 0.97

a V Ois the velocity obtained in the absence of inhibitor sad is relative to the velocity obtained at 1.5 mM FDP which is arbitrarily set at 100.

concentrations are required to give 50% inhibition of the CL1136 ADPglucose pyrophosphorylase. Fructose diphosphate can partially reverse the 5‘-AMP inhibition of the mutant ADPglucose pyrophosphorylases in the same manner as noted previously for the parent strain enzyme (66, 56,67). A very interesting difference has been noted between the CL1136 ADPglucose pyrophosphorylase and the SG5 and parent strain enzymes with respect to inhibition by 5’-adenylate in the absence of activator. As indicated previously, a portion of the parent strain ADPglucose pyrophosphorylase activity is resistant to inhibition. Depending on conditions, 30-5574 of the E . coli B enzyme is not inhibited by high concentrations of 5’-adenylate in the absence of activator (Table VII). The SG5 enzyme is similarly resistant to inhibition in the absence of fructose-dip, and depending on the concentration of ATP and Mg2+the activity resistant to inhibition ranges from 40 to 60%. In contrast, 87% of the CL1136 ADPglucose pyrophosphorylase activity can be inhibited in the absence of FDP by 5’-AMP. The inhibition curve in the absence of FDP is hyperbolic in shape with the Hill constant f i , being close to one. Thus, in the absence of activator, the inhibitor binding sites of the CL1136 ADPglucose pyrophosphorylase do not interact. Since CLll36 ADPglucose pyrophosphorylase is almost fully active in the absence of activator and is also almost fully inhibited in the absence of activator, it is possible that mutation in CL1136 has converted the ADPglucose pyrophosphorylase to a form where its conformation is identical or very similar to the conformation of the E . coli B enzyme in the presence of activator. The effect of energy charge on the mutant enzymes is shown in Table

114

JACK PREISS

TABLE XIV RESPONSEOF THE E . coli B AND MUTANTADPQLUCOSE PYROPHOSPHORYLASES TO ENERQY CHARQEIN THE PRESENCE OF 1.5 mM FRUCTOBE-DIP Relative velocity. Energy charge 1.0 0.95 0.90 0.85 0.80 0.75 0.70 0.6 0.5

E . wli B

SG5

CL1136

100 91 72 46 26 10 4.5 1.0 1 .o

100 98 87 74 64 42 22 8.5 2.0

100 98 94 88 81 74 61 33 14

The velocity obtained at each energy charge value is expressed relative to the velocity obtained at an energy charge of 1.0, which is arbitrarily assigned a value of 100.

XIV and is compared to the response of the E . coli B enzyme to energy charge. At an energy charge value of 0.7 the SG5 and CL1136 ADPglucose pyrophosphorylases have 5- and 13.5-fold greater activity than the E. coli B enzyme. At what is considered to be the physiological energy charge value for E. coli, 0.8 ( 6 3 ) ,the SG5 and CL1136 enzymes have 2%-3 times the activity of the parent enzyme. The CL1136 is highly active even a t an energy charge of 0.6, while the E. coli B and SG5 enzymes are virtually inactive. The CL1136 ADPglucose pyrophosphorylase is more active than the SG5 enzyme under the same conditions of energy charge and concentration of activator. The above data indicate that the increased accumulation of glycogen in the mutants SG5 and CL1136 results from a modification of their ADPglucose pyrophosphorylases, causing a greater affinity for the activators and lower affinity for the inhibitor. These changes in affinity of the effector molecules cause the mutant enzymes to have greater activity than the E. coli B enzyme under the same conditions. The regulatory effects observed in vitro are thus important for the in vivo regulation of glycogen synthesis in E . coli B. Correlation of the relative insensititrity of the SG5 and CL1136 ADPglucose pyrophosphorylases to AMP inhibition and their responses to energy charge with the increased rates of accumulation in the cells are in agreement with the concept that glycogen synthesis is controlled by energy charge (11, 56-58). The important factors involved in the allosteric regulation of ADPglucose and glycogen synthesis in E. coli would be the activation by fructose-dip and inhibition by 5’-AMP.

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

115

3. E . coli Mutant SGl4 ADPglucose Pyrophosphorylase

Mutant SG14 accumulates glycogen a t about 55-65% the rate of E. coli B and contains about 16% of the ADPglucose synthesizing activity as E . coli B ( 7 8 ) .Yet the activity present is still 3-5-fold greater than that required for the observed rate of glycogen accumulation in SG14. The concentrations of ATP and Mg2+required for 50% of maximal activity ( 8 d are 4-5-fold higher for the SG14 enzyme than the E.coli B enzyme. Whereas the So.5values for ATP and Mg2+are 0.39 and 2.38 mM, respectively, for E. wli B enzyme in the presence of 1.5 mM FDP, the So.b values for ATP and Mg2+are 1.6 and 10.2 mM, respectively, for the SG14 ADPglucose pyrophosphorylase in the presence of saturating FDP (4.0 mM). Reports in the literature indicate that the A T P level in growing E. coli ranges from 2 to 6 mM (79, 80) and the MgZ+level is about 1830 mM (81-84). Therefore, the SG14 ADPglucose pyrophosphorylase would essentially be saturated with respect to these substrates. The apparent affinities (&,.=,) for glucose l-P for the E. coli B and SG14 enzymes are about the same (0.036 mM) ( 7 8 ) . The major differences between the SG14 and E. coli B ADPglucose pyrophosphorylases appear to be their sensitivities toward activation and inhibition ( 7 8 ) . About 12-fold more F D P is needed for 50% maximal stimulation of the SG14 ADPglucose pyrophosphorylase (A0.5= 0.82 mM) than for half-maximal stimulation of the E. coli B enzyme, while the A0.5 for PLP for the SG14 enzyme (0.44 mM) is about 25-fold higher than for the E. coli B enzyme. To about the same extent PLP and FDP stimulate ADPglucose synthesis catalyzed by the E. coli B enzyme. However, the stimulation of the SG14 ADPglucose pyrophosphorylase seen with PLP is only one-half that elicited by fructose-dip. A notable difference is that TPNH does not stimulate the SG14 enzyme. Compounds similar to T P N H in structure, such as P R P P and 2'-PADPR, that are capable of activating the E. coli B ADPglucose pyrophosphorylase do not activate the SG14 enzyme. Since the apparent affinity of the SG14 enzyme for its activators is considerably lower than that observed for the E. coli B ADPglucose pyrophosphorylase it was an unexpected finding that SG14 is capable of accumulating glycogen at one-half the rate observed for the parent strain. This rate is accounted for by the relative insensitivity of the SG14 en79. H. A. Cole, J. W. T. Wimpenny, and D. E. Hughes, BBA 143, 445 (1967). 80. A. S. Bognara and L. R. Finch, BBRC 33, 15 (1968). 81. M. Lubin and H. L. Ennis, BBA 80, 614 (1964). 82. S. Silver, Proc. Nat. Acad. Sci. U.S. 82, 764 (1969). 83. J. G. Lusk, R. J. P. Williams, and E. P. Kennedy, JBC 243, 2618 (1968). 84. M. Webb, J. Gen. Microbial. 43, 401 (1966).

116

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zyme to inhibition by 5‘-adenylate. The SG14 ADPglucose pyrophosphorylase is much less sensitive to 5’-AMP inhibition in the concentration range of 0-0.2 mM than is the parent strain enzyme (Table XV). At a saturating concentration of FDP for the SG14 enzyme (4.0 mM) only 7% inhibition of the SG14 enzyme is observed a t 0.2 mM 5’-AMP; the same concentration of 5’-AMP gives 40% inhibition of the E . coli B enzyme. At a concentration of FDP which gives 80% of maximal velocity (1.5 mM) for the SG14 enzyme, 0.2 mM 5’-AMP causes inhibitions of 84 and 33% with the E. coli B and SG14 enzymes, respectively. A decrease of FDP concentration to 1.0 or 0.5 mM increases the sensitivity of the E . coli B ADPglucose pyrophosphorylase to inhibition. However, a t these concentrations of FDP, the sensitivity of the SG14 ADPglucose pyrophosphorylase to $-AMP remains the same or becomes less than that seen a t 1.5 mM FDP. At concentrations of 0.5-1.0 mM of FDP the E . coli B enzyme is inhibited 90% or more by 0.2 mM AMP while the inhibition of the SG14 enzyme ranges from 12 to 30%. TABLE XV E . COli B AND SG14 ADPOLUCOSE PYROPHOSPHORYLASES IN THE PRESENCE OF VARYING

5‘-ADENYLATE INHIBITION OF

CONCENTRATIONS OF FRUCTOSE-DIP AT P H ~

7.0

~

P e r c e n t inhibition

FDP (mM)

AMP (mM)

4.0

0.1 0.2

1 .o

0.5

15

SG14 3 7 39 49 80

1.0 1.5

40 68 90 96

0.1 0.2 0.5 1.0

44 84 98 99

0.1 0.2 0.5 1.0 1.5

72 90 97 98 >98

14 30

0.1 0.2

92 97 >98 >98 >98

4 12 40

0.5

1.5

E . wli B

0.5 1.0 1.5

19 33

54 72

55 68 73

55 60

3.

ADENOSINE DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

117

Although the modification of the SG14 enzyme causes it to have a lower apparent affinity for its activators, it also renders the enzyme less sensitive to 5’-AMP inhibition. These two effects appear t o compensate for each other and allow SG14 to accumulate glycogen a t about one-half the rate of the parent strain. It is of interest to note that the concentration of F D P in E. coli has been found to be in the range of 0.6-2.5 mM (86, 86). The 5’-AMP concentration in E. coli B is estimated to be 0.14 f 0.05 mM (86). The data obtained from the kinetic studies of the SG14 ADPglucose pyrophosphorylase further suggest that fructose-dip is the most important physiological activator of the E . coli ADPglucose pyrophosphorylase. This is based on the observation that T P N H is not an activator of the SG14 ADPglucose pyrophosphorylase and that the concentration of P L P needed for activation of the enzyme (&5 = 0.44 mM) is considerably higher than that reported to be present in E . coli B (87). The concentration of PLP is 6-8 p M , and most of this metabolite is probably protein bound in the cell and unavailable for activation of the ADPglucose pyrophosphorylase. The concentration of FDP in E. coli is about 2.5 mM (86), and the A0.5 of SG14 ADPglucose pyrophosphorylase is 0.82 mM (78). The concentration of FDP in the E. coli cell required for activation of the SG14 enzyme is sufficient to account for the activation of the SG14 ADPglucose pyrophosphorylase required for synthesis of ADPglucose a t the rates required for the glycogen accumulation observed in SG14 (78).

IV. Physical Properties of the ADPglucose Pyrophosphorylases

The kinetic studies reviewed in the previous sections indicate that the ADPglucose pyrophosphorylases of all classes exhibit many interesting and unique properties in their activator and inhibitor specificities and in the interaction of the activators with the substrates and inhibitors. An obvious task is to attempt to correlate these complex kinetic properties of the enzymes with their physical and chemical properties. However, endeavors in this area have been limited by the difficulty in obtaining pure enzymes in quantities sufficient to do the necessary analyses. With the isolation of a mutant of E . coli B derepressed in the syntheses of 85. W. P. Hempfling, M. Hofer, E. J. Harris, and B. C. Pressman, BBA 141, 391 (1987). 86. 0. H. Lowry, J. Carter, J. B. Ward, and L. Glaser, JBC 246, 6511 (1971). 87. W. B. Dempsey, J . Bacterial. 90, 431 (1965).

118

JACK PREISS

ADPglucose pyrophosphorylase and ADPglucose ~-1,4-glucan-4-o-glucosyltransferase (7S, 76) and with the availability of more efficient purification techniques such as affinity chromatography, this difficulty may be overcome in the near future. Presently, three ADPglucose pyrophosphorylases have been purified to apparent homogeneity (19, 88, 89). The R. rubrum enzyme has been purified to homogeneity by preparative gel electrophoresis (19). It shows only one band on analytical disc gel electrophoresis at three different pH values. Molecular weight determinations by sedimentation equilibrium according to Yphantis (90)also indicated homogeneity of the protein, as well as an average MW of 195,000 for an assumed Z, of 0.70 and 245,000 for an assumed Z, of 0.75. The disc gel technique of Hedrick and Smith (91) indicated a MW of about 225,000, in agreement with the sedimentation equilibrium experiments (19). The specific activity of the R . rubrum enzyme is 54.5 pmoles of ATP formed per minute per milligram of protein at 37” in the presence of 25 mM pyruvate (activator) and a t pH 8.25. In the absence of activator the specific activity is 27.2 pmoles of ATP formed per minute per milligram of protein (pH 8.25). The spinach leaf enzyme has also been purified to homogeneity by preparative gel electrophoresis (88). Its specific activity is 93 pmoles of ATP formed per minute permilligram of protein a t 37” and pH 7.5 in the presence of 1 mM 3PGA. I n the absence of activator the specific activity is 35.5 pmoles of ATP formed per minute per milligram a t pH 7.5 and 37”. The disc gel technique of Hedrick and Smith (91) indicated that the MW of the spinach leaf ADPglucose pyrophosphorylase was 210,000. The molecular weight found by sedimentation equilibrium (90) ranged between 195,000 for an assumed 7i of 0.70 and 240,000 for an assumed Z, of 0.75 (88). The E . coli B ADPglucose pyrophosphorylase was purified to homogeneity from a mutant, SG3, derepressed in the levels of the enzyme (89). Its specific activity is 103 pmoles of ATP (or 104 pmoles of ADPglucose) formed per minute per milligram of protein a t 37”, pH 8.0, and in the presence of 1.5 mM FDP. In the absence of activator and under the same conditions of temperature and pH 17 pmoles of ADPglucose (or 59 pmoles of ATP) were formed per minute per milligram. Analysis of the E . coli enzyme by the Hedrick and Smith disc gel electrophoresis procedure (91) indicated a MW of 211,000 5 6,000. Sedi88. G. Ribereau-Gayon and J. Preiss, “Methods in Enzymology,” Vol. 23, p. 618, 1971. 89. H . Ozaki and J. Preiss, “Methods in Enzymology,” Vol. 28, p. 409, 1972. 90. D. A. Yphantis, Biochemistry 3, 297 (1964). 91. J. L. Hedrick and A. J. Smith, ABB 126, 155 (1968).

3. ADENOSINE

DIPHOSPHORYL GLUCOSE PYROPHOSPHORYLASE

119

mentation velocity experiments a t varying protein concentrations (0.753.0 mg/ml) in 0.1 M tris-HC1 buffer, pH 7.2 gave a symmetrical peak a t each concentration. By extrapolation of the s20,wvalues to zero protein concentration the sz0,, a t infinite dilution was determined to be 10.9 S. The molecular weight by sedimentation equilibrium was determined by a meniscus depletion run as described by Yphantis (90) and indicated a MW of 207,000 2 7,000 assuming a partial specific volume for the enzyme of 0.74. Sucrose density gradient centrifugation according to Martin and Ames (92)gave a MW value of 204,000 f 6,000. The standards used were beef liver cat,alase (hlW, 240,000) and pig heart lactate dehydrogenase (MW, 135,000). Electrophoresis in 0.1% sodium dodecyl sulfate by the procedure of Weber and Osborn (93) revealed only one protein component of the pure enzyme with a MW of 53,000& 2,000 using bovine serum albumin, catalase, ovalbumin, yeast alcohol dehydrogenase, and pig heart lactate dehydrogenase as standards. Meniscus depletion sedimentation equilibrium studies (90)of the enzyme in 6 M guanidine-HC1 solution containing 0.102 M P-mercaptoethanol and 0.02 M NaCl (94) gave a MW of the dissociated protein of 48,000 with an assumed partial specific volume of 0.74. Based on these observations it is concluded that the ADPglucose pyrophosphorylase from E. coli B consists of four subunits with identical molecular weights (89). The three ADPglucose pyrophosphorylases purified thus far appear to have very similar molecular weights.

92. R. G. Martin and B. N. Ames, JBC 236, 1372 (1961). 93. K. Weber and M. Osborn, JBC 244, 4406 (1969). 94. K. Kawahara and C. Tanford, JBC 241, 3228 (1966).

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The Adenosvltransferases S. HARVEY MUDD I. Introduction . . . . . . . 11. Methionine Adenosyltransferase . . . A. Significance and Distribution . . B. The Net Reaction . . . . C. Purification and Physical Properties D. Catalytic Properties . . . . E. Regulation and Genetics . . . 111. B,?, Adenosyltransferase . . . . . A. Significance and Distribution . . B. The Net Reaction . . . . C. Purification and Physical Properties D. Catalytic Properties . . . . IV. Conclusion . . . . . . . .

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1. Introduction

The adenosyltransferases comprise a group of enzymes which transfer the adenosyl moiety, i.e., the structure which would result from removal of the 5’-OH group of adenosine (Fig. 1). At present, there are only two enzymes known which catalyze the transfer of this group. The first to be discovered was the enzyme which catalyzes the formation of S-adenosylmethionine, the chief biological methyl donor. This enzyme is now given the systematic name ATP :L-methionine S-adenosyltransferase, EC 2.5.1.6, and the trivial name methionine adenosyltransferase. The second known adenosyltransferase catalyzes the formation of one of the 121

122

S. HARVEY MUDD

FIG.1. The adenosyl group.

coenzymically active derivatives of vitamin B,, ( 1 ) . The latter enzyme might appropriately be designated ATP :B,,, Go-adenosyltransferase (BIZB adenosyltransferase). In the first example the adenosyl group is transferred from ATP to the sulfur atom of methionine. I n the second, the adenosyl group of ATP is transferred to the reduced cobalt atom of the cobalamin molecule. Each of these reactions may occur by a nucleophilic attack upon carbon-5 of the ribose portion of ATP (the -CH,on the right in Fig. 1 ) . The bond between this carbon and the oxygen of the proximal phosphate of ATP is broken, to be replaced by a linkage to either the sulfur of methionine or the cobalt of BIZ. Unfortunately, in the biochemical literature dealing with these two 1. For a complete explanation of the nomenclature applicable to compounds related to BIZsee IUPC, Commission on the Nomenclature of Biological Chemistry [JACS 82, 5581 (1960)l or L. Ljungdahl, E. Irion, and H. G . Woods [Fed. Proc., Fed. Amer. SOC.Exp. B i d . 25, 1642 (1966)l.The compounds discussed in this chapter are, in general, cobamide derivatives. Cobamide consists of a fundamental corrin macro ring system of four nitrogen-containing five-membered rings joined through three bridge carbon atoms and surrounding a central cobalt atom coordinated to the four nitrogens. Designated side chains are present on the ring system. One of these side chains consists of a sequence of propionic acid linked by an amide bond to 2-hydroxypropylamine. This structure is cobinamide. If a phosphate residue is esterified to the 2-hydroxypropylamine of cobinamide and linked also to the 3-OH of a ribose moiety, the resulting compound is termed cobamide. The cobamide derivatives of concern in this chapter contain a heterocyclic base which is attached in an N-glycosidic linkage to the ribose moiety, thus completing a nucleotide sequence. The heterocyclic base is also coordinated to the-. cobalt atom (conventionally in the position “below” the ring). Cobamide derivatives in which the heterocyclic base is 5,6-dimethylbenrimidazole are called cobalamin and will be abbreviated here as BIZ.The substituent a t the sixth coordination position of the cobalt atom (i.e., “above” the ring) will then be designated by prefixes (e.g., adenosyl-BIz or HO-BIZ). In compounds such as HO-BIZ or CN-BlZ, the cobalt atom is assigned a 3’ valence state. Succeeaive one electron reductions yield BIZ,, then BIS., with the cobalt in the 2+ and 1+states, respectively.

4.

THE ADENOSYLTRANSFERASES

123

reactions and their products, it has become customary to use the term adenosyl to designate the group shown in Fig. 1 when discussing S-adenosylmethionine and compounds of similar structure (e.g., S-adenosylhomocysteine), whereas the same group has come to be referred to by the term 6‘-deoxyademsyl when it is present in compounds related to BIZ (e.g., 5’-deoxyadenosylcobalamin) [see, for example, several recent reviews ( % 4 ) ] . The term adenosyl appears to be adequate, and has the advantages of priority (5-7) , simplicity, and lack of potential confusion with other deoxyadenosyl compounds (such as the 2’-deoxyadenosyl series found in DNA and its metabolites). This term will therefore be used for both the methionine and the B,, derivatives in this presentation. I n this chapter each of the two known adenosyltransferases will be discussed. In the summary, points of similarity and difference between the enzymes will be emphasized.

II. Methionine Adenoryltranrferase

A. SIGNIFICANCE AND DISTRIBUTION During early in vitro investigations of methyl transfer reactions in which methionine was the ultimate source of the methyl group, it became clear that a requirement in crude systems for oxygen (8) could be replaced by ATP (9, 10). During further investigation of these reactions, Cantoni demonstrated that ATP was required for an enzyme-catalyzed conversion of methionine to a new derivative (11). This compound he characterized as S-adenosylmethionine (5, 6). The role of S-adenosylmethionine as the major biological methyl donor has by now been amply confirmed. A review of enzyme-catalyzed methyl transfers revealed that, by 1962-63, more than 30 such reactions had been studied and, in each 2. H. A . Barker, BJ IW,1 (1967). 3. H. Weissbach and H. Dickerman, Physiol. Rev. 45, 80 (1965). 4. T. C. Stadtman, Science 171, 859 (1971). 5. G. L. Cantoni, JACS 74, 2942 (1952). 6. G. L. Cantoni, JBC 204, 403 (1953). 7. P. G. Lenhert and D. C. Hodgkin, Nature (London) 192, 937 (1961). 8. W. A, Perlzweig, M. L. C. Bernheim, and F. Bernheim, JBC 150, 401 (1943). 9. H. Borsook and J. W. Dubnoff, JBC 171, 363 (1947). 10. G. L. Cantoni, JBC 189, 203 (1951). 11. G. L. Cantoni, JBC 189, 745 (1951).

124

S. HARVEY MUDD

case, S-adenosylmethionine had been implicated as the immediate methyl donor. The only exceptions were methyl transfers to homocysteine to form methionine itself (12).The general methyl transfer reaction may then be formulated according to Eq. (1) A-R-S@-CHzCH~CH(NHz)COOH

I

CH s

+ XH

--j

CHsX

+ A4-R-S--CH&HzCH(NH~)COOH + H @

(1) where A and R are the adenine and ribose portions of the adenosyl moiety and X symbolizes the methyl acceptor group. No evidence has emerged subsequent to 1963 to challenge the general importance of S-adenosylmethionine as methyl donor or S-adenosylhomocysteine as the thioether formed from this sulfonium compound during the methyl transfer reactions. The widespread role of S-adenosylmethionine is suggested not only by the fact that enzymes capable of transferring the methyl group of this compound have been found in bacteria, yeast, higher plants, amphibians, and many mammalian tissues (12) but also by the fact that methionine adenosyltransferase, the enzyme catalyzing the synthesis of this compound, has also been demonstrated in an extensive array of biological sources, including bacteria (IS), yeast ( I 4 ) , higher plants (I6),and a variety of mammalian tissues (11, 16). Furthermore, recent investigations have revealed several physiological roles for S-adenosylmethionine in addition to that of biological methyl donor. As was first shown by the Tabors and their collaborators, S-adenosylmethionine may be enzymically decarboxylated, forming a compound which may now donate its propylamine group to putrescine to yield spermidine (IS). The physiology and biosynthesis of spermidine and other polyamines, which are now attracting increasing attention because of their possible involvement in regulation of growth, have recently been the subject of several reviews (17-19). Finally, a number of enzyme reactions are now known in which S-adenosylmethionine is involved, not as a substrate, but rather as an activator, a feedback inhibitor, or a 12. S. H. Mudd and G . L. Cantoni, Compr. Bwchem. 15, 1 (1964). 13, H. Tabor, S. M. Rosenthal, and C. W. Tabor, JACS 79, 2978 (1957). 14. S. H. Mudd and G. L. Cantoni, JBC 231, 481 (1958). 15. S. H. Mudd, BBA 34 354 (1960). 16. S. H. Mudd, J. D. Finkelstein, F. Irreverre, and L. Laster, JBC 240, 4382 (1965). 17. H. Tabor, C. W. Tabor, and S. M. Rosenthal, Annu. Rev. Biochem. 30, 579 (1981). 18. H. Tabor and C. W. Tabor, Pharmacol. Rev. 16, 245 (1964). 19. H. G. Williams-Ashman, A. E. Pegg, and D. H. Lockwood, Advan. Enzyme Regul. 7, 291 (1969).

4. THE ADENOSYLTRANSFERASES

125

repressor. A discussion and compilation of these reactions will be found elsewhere (2U,2 1 ) .

B. THE NET REACTION Adenosine triphosphate is the adenosyl donor in the methionine adenosyltransferase reaction. Assignment of this role is based upon the absolute requirement for ATP in the reaction (IS, 1.6, 22) and the observed incorporation of radioactivity from ATP-l'C into ["C] -8adenosylmethionine (22, 23) . For each mole of 8-adenosylmethionine formed, one mole of ATP disappears ( 2 2 ) . The fate of the three phosphate groups of ATP during the adenosyl transfer reaction has been studied extensively. With relatively crude enzyme preparations, there is a stoichiometric release of 3 moles of inorganic phosphate (Pi) from ATP for each mole of S-adenosylmethionine formed ( 2 2 ) . As was first shown by Cantoni and Durell using an enzyme purified from rabbit liver, if contaminating pyrophosphatase activity is removed, the products formed are 1 mole of Pi and 1 mole of inorganic pyrophosphate (PPi) for each mole of S-adenosylmethionine ( 2 2 ) . The same stoichiometry is observed with methionine adenosyltransferase purified from baker's yeast (14). Further experiments with ["PIATP demonstrated that the Pi originates from the terminal or y phosphate group of ATP, whereas the PPi originates from the two proximal (a and p ) phosphates. For example, ATP labeled in the p and y phosphate residues (i.e., [P,y-"P]ATP) yielded szPi and 32PPi.The specific radioactivities of these products were those expected from a cleavage between the p and y phosphate residues of ATP. The alternate cleavage between the a and p residues would have yielded Pi free of radioactivity. These results were extended by observation of the expected labeling pattern when [ Q - ~ ~ P I A T was P used as substrate ( 2 2 ) . Subsequently, patterns of labeling consistent with the formation of PPi from the a and p phosphates, and of Pi from the y phosphate, of A T P were observed with the adenosyltransferases of both yeast (14, 24) and Escherichia coli (13,17). Several studies have been concerned with the stereochemistry of the 20. J. B. Lombardini and P. Talalay, Advan. Enzyme Regul. 9, 349 (1971). 21. S. H. Mudd, in "Metabolic Hydrolysis and Metabolic Conjugation" (W. H. Fishman, ed.), Vol. 3, p. 297. Academic Press, New York, 1973. 22. G. L. Cantoni and J. Durell, JBC 225, 1033 (1957). 23. S. H. Mudd, G . A. Jamieson, and G . L. Cantoni, BBA 34 164 (1960). 24. S. H. Mudd, JBC 238, 2156 (1963).

126

S. HARVEY MUDD

amino acid product formed by methionine adenosyltransferase. S-Adenosylmethionine [and its decarboxylated derivative (13)] is unique among known biological sulfonium compounds in that three different substituents are attached to the sulfur atom, resulting in an optically active center. To determine whether methionine adenosyltransferase synthesizes only one, or both, of the two compounds differing in configuration a t this optical center, D e La Haba and co-workers (26) compared the properties of enzymically formed S-adenosylmethionine with those of S-adenosylmethionine synthesized by the chemical methylation of S-adenosylhomocysteine with CHJ. The enzyme which catalyzes the transfer of the methyl group from S-adenosylmethionine to guanidinoacetate was able to utilize almost all the enzymically formed sulfonium compound, but only half the chemically synthesized material. The sulfonium compound remaining after exhaustive treatment with guanidinoacetate methyltransferase was reisolated and shown to be identical with enzymically synthesized S-adenosylmethionine save for an altered optical rotation. Whereas S-adenosylmethionine formed by the action of methionine adenosyltransferase of either rabbit liver or baker's yeast has a specific rotation ( [ a ] of +47-4B0, the chemically methylated (i.e., racemic) compound has a specific rotation of +52", and the sulfonium compound inactive as a methyl donor has a specific rotation of + 5 7 O . These results indicate that the methionine adenosyltransferases of rabbit liver and baker's yeast form predominantly, or exclusively, only one of the possible sulfonium compounds which differ with respect to the configuration about the trivalent sulfur atom. The material formed by these enzymes may be designated as ( - ) -S-adenosylmethionine, the ( - ) indicating the contribution of the sulfonium center to the overall optical activity of the molecule. The inactive compound is then ( ) -8-adenosylmethionine. During these initial studies it was shown also that two additional enzymes metabolizing S-adenosylmethionine utilize preferentially the ( - ) compound ( 2 6 ) . Subsequently, the list of enzymes which demonstrate specificity for ( - ) -S-adenosylmethionine has been greatly extended [see summary in reference (21)1. Since the methionine adenosyltransferases of human liver (16) and of barley seedlings (16) also form the ( - ) compound, it appears likely that this diastereoisomer is the dominant one in biological materials. Taken together, these studies of methionine adenosyltransferase allow the following formulation of the net reaction catalyzed by the enzyme

+

A-R-P*-P*-P

+ methionine -+

(-)-S-adenosylmethionine

+ P*P*,+ Pi

(2)

25. G . De La Haba, G. A. Jamieson, S. H. Mudd, and H. H. Richards, JACS 81, 3975 (1959).

4. THE

ADENOSYLTRANSFERASES

127

where A-R-P-P-P indicates ATP and the fates of the phosphate groups are shown by the asterisks.

C. PURIFICATION AND PHYSICAL PROPERTIES Methionine adenosyltransferase activity extracted from baker’s yeast was purified about 200-fold by acetone fractionation, adsorption on and elution from calcium phosphate gel, and negative adsorption with bentonite. More uniform preparations of slightly higher specific activity were obtained when the last step was modified to include first a negative adsorption with bentonite, then adsorption on and elution from this material. The adenosyltransferase resulting from the latter procedure had a specific activity of 8-11 pmoles S-adenosylmethionine formed per milligram of protein per 30 min, and represented a 300400-fold purification (24 1 . It was essentially free of pyrophosphatase activity, which had been removed by bentonite adsorption (14) and free also of ATPase activity (2.4). In subsequent purifications of the adenosyltransferase the same basic scheme has been followed, although the bentonite steps have been slightly modified (26, 27). After elution from bentonite the adenosyltransferase activity moves with the chief protein peak during sedimentation through a sucrose gradient (26, 28) and the preparation shows a single symmetrical peak on sedimentation in the analytical ultracentrifuge (27). However, the enzyme is not yet pure, as shown by several lines of evidence: 1. After disc gel electrophoresis several bands of protein are present ( 2 6 , 2 7 ) .Adenosyltransferase activity is associated with the major band (27). 2. Some specific contaminating enzymic activities have been identified in the preparation a t this stage, for example, a peptidase capable of cleaving methionylmethionine. [This peptidase can be removed by chromatography upon DEAE-Sephadex A-50 (26).] Another contaminating activity is a n enzyme catalyzing ADPATP exchange. [This activity can be removed by sedimentation in a sucrose gradient (29).] 3. Chou and Talalay added an alcohol fractionation step to the usual purification procedure to obtain a methionine adenosyl26. H. Hagenmaier, Ph.D. Thesis, Cornell University, Ithaca, New York, 1965. 27. R. C. Greene, Biochemistry 8, 2255 (1969). 28. S. H. Mudd, JBC 237, PC1372 (1962). 29. S. H. Mudd, in “Transmethylation and Methionine Biosynthesis” (S.K. Shapiro and F. Schlenk, eds.), p. 33. Univ. of Chicago Press, Chicago, Illinois, 1965.

128

S. HARVEY MUDD

transferase preparation with a specific activity of 25-32 (30). This preparation would appear to be two to three times as active as the best obtained previously. On the basis of the rate of sedimentation of yeast methionine adenosyltransferase activity in a sucrose gradient and assumptions as to the partial specific volume and shape of the enzyme, a crude molecular weight of 157,000 was calculated ( 2 4 ) . Greene used measurements of sedimentation velocity in a double sector cell to calculate sedimentation and diffusion constants. I n conjunction with an estimate of the partial specific volume, based upon the amino acid composition of the enzyme, the results were used to calculate a molecular weight of 100,OOO ( 3 1 ) . Using enzyme prepared by a somewhat different method, Chou and Talalay calculated a molecular weight of approximately 44,000 as a result of experiments involving gel filtration through Sephadex G-200 (SO). The reasons for these sizable discrepancies in estimates of molecular weight are not known. The activity of yeast methionine adenosyltransferase is quite stable during the purification procedures used. I t is stabilized during heat treatment by the presence of glutathione ( 1 4 ) , or of S-adenosylmethionine, or certain monovalent cations ( 2 4 ) . Methionine adenosyltransferase extracted from rabbit liver has been purified by ammonium sulfate fractionation, followed by isoelectric precipitation, repeated precipitation with ammonium sulfate, and heat treatment. The result was a 90-fold purification with a 20% yield of activity. The specific activity at the final stage was 2.3-3.5 pmoles S-adenosylmethionine formed per milligram of protein per 30 min. Contaminating pyrophosphatase was removed by isoelectric precipitation and ATPase by the heat treatment ( 2 2 ) . Subsequent modifications of this procedure have utilized column chromatography upon DEAEcellulose following the initial precipitation with ammonium sulfate to yield preparations with specific activities of 4.9 (32) or 6.0 (33). The liver adenosyltransferase has been stabilized during purification by GSH ( W ) ,or 2-mercaptoethanol combined with EDTA ( 3 2 ) ,or 2-mercaptoethanol with EDTA and glycerol (33). Stabilization during frozen storage is achieved by the addition of 20% glycerol ( 3 2 ) : The E . co2i methionine adenosyltransferase has been purified approxi30. T. C. Chou and P. Talalay, Biochemistry 11, 1065 (1972). . 6th, 1964 Abstract, Sect. 4, p. 310 31. R. C. Greene, Proc. Znt. C o n g ~ Biochem., (1964). 32. F. Pan and H. Tarver, ABB 119, 429 (1967). 33. J. B. Lombardini, A. W. Coulter, and P. Talalay, Mol. Pharmacol. 6, 481 ( 1970).

4.

THE ADENOSYLTRANSFERASES

129

mately 1500-fold and freed of contaminating pyrophosphatase activity (34), but details of the methods used have not been published.

D. CATALYTIC PROPERTIES 1. Assay

Several methods are available for assay of methionine adenosyltransferase activity. In earlier experiments the methionine-dependent formation of Pi from ATP was determined (11). This assay is useful only if the enzyme is not contaminated by a large excess of ATPase but has been successfully applied to relatively crude rabbit liver preparations (11 ) . If the adenosyltransferase is freed of pyrophosphstase, the formation of each mole of S-adenosylmethionine is accompanied by formation of one mole of Pi rather than three. Excess pyrophosphatase may then be added back to the purified enzyme to restore the 3: 1 stoichiometry and to avoid inhibition by the reaction product, PPi ( 2 2 ) . An alternative method of assay takes advantage of the fact that S-adenosylmethionine has a net positive charge at neutral pH, whereas ATP does not. Treatment of an aliquot of the reaction mixture with Dowex-1 (Cl-) adsorbs unreacted ATP and leaves the S-adenosylmethionine in the supernatant solution, where it is readily quantitated by its absorbance a t 259 nm ( 2 2 ) .This method has the advantage of increased sensitivity and relatively less interference from contaminating ATPase and has been used extensively as originally described (22) or with minor changes (14,24,26,27,3 5 ) . A modification of this method uses [14C]ATP as substrate (27, 3 3 ) . Since it is then possible to measure activity with a relatively low concentration of ATP, this modification appears t o be potentially useful in determination of inhibition by compounds competing with this substrate. A recently developed assay depends upon conversion of [ '"C]methionine to [ 14C]-8-adenosylmethionine. The latter is adsorbed on a small column of Dowex-50 (NH,') at neutral pH, whereas unreacted [14C]methionine passes through the column. [ '"C] -8-Adenosylmethionine is then eluted with aqueous ammonium hydroxide and determined by measurement of radioactivity (16). This assay is simple to perform and, because of its increased sensitivity and freedom from interference, has proven useful for measurements with crude extracts of low activity (16). I t is most advantageous for inhibition studies since measurable rates of 34. H. Tabor and C . W. Tabor, Fed. Proc., Fed. Amer. Sac. Exp. B i d . 19, 6 (1960). 35. R. L. Hancock, Cancer Res. 26, 2425 (1966).

130

S. HARVEY MUDD

reaction are achieved a t extremely low concentrations of the substrate, methionine, permitting detection of inhibition by competing compounds (29, 33). 2. Reversibility, Partial Reactions, and Mechanistic Considerations

It is apparent from the stoichiometry of the overall conversion catalyzed by methionine adenosyltransferase that the reaction must be u complicated one. At least two steps would be expected since during the course of the reaction two bonds originally present in ATP are broken: the C5’-0 bond and the 0-P7 bond. Nevertheless, extensive purification of methionine adenosyltransferase from three sources has given no indication that the native enzyme can be physically separated into more than one component (14, 22, 3 4 ) . No free intermediate has been found to accumulate. Unbound ADP (17,22, 2 3 ) , adenosine, and 3,5’-cycloadenosine (23) have been specifically excluded as intermediates. The origin of Pi exclusively from the y-phosphate of ATP precludes the possibility that free tripolyphosphate (PPPi) is an intermediate since cleavage of this symmetrical compound to Pi and PPi would mean that both the a and y phosphates of ATP would be precursors of Pi. On the other hand, a good deal of evidence is now available to support the hypothesis that bound PPPi is an intermediate in this reaction. Thus, in studies with the purified yeast enzyme it was shown that after denaturation of the enzyme, a small amount of PPPi could be detected in the reaction mixture. The PPP; was somewhat less than stoichiometric with the amount of enzyme, and its formation was dependent upon methionine ( 2 3 ) .It was postulated that this PPPi remains bound to the adenosyltransferase in an orientation determined by its .origin in ATP and is cleaved to Pi and PPi in such a way that only the moiety which was formerly the y phosphate of ATP gives rise to Pi. I n accord with this hypothesis, it was shown that highly purified preparations of yeast (24, 27, 28, SO) and E. coli (18) methionine adenosyltransferase contain tripolyphosphatase activity. The two activities remain together during centrifugation through a sucrose gradient (28) and display the same order of activation by monovalent cations ( 2 4 ) . A functional relationship between the tripolyphosphatase activity and 8-adenosylmethionine may be inferred from the fact that the tripolyphosphatase activity is stimulated 10-12-fold by the presence of 8-adenosylmethionine (27, 28, SO). I n this portion of the reaction 8-adenosylmethionine is acting as an effector since there is no indication that this compound is chemically altered by virtue of its participation in the events of tripolyphosphate hydrolysis. The high specificity of the stimulation is shown by the fact that a num-

4.

THE ADENOSYLTRANSFERASES

131

ber of compounds structurally related to S-adenosylmethionine are ineffective in stimulating the tripolyphosphatase activity (28). In further support of the postulated ability of the methionine adenosyltransferase to bind a condensed phosphate compound, it was shown that the purified yeast enzyme binds PPi strongly enough to form a complex detectable after gel filtration under nonequilibrium conditions, whereas, under the same conditions, Pi did not form a detectable complex with the enzyme. The capacity to bind PPi sedimented with adenosyltransferase activity during centrifugation through a sucrose gradient (28). Binding of PPP, was not examined directly in these experiments because of the tripolyphosphatase activity of the enzyme. However, kinetic studies have indicated that PPPi is a more potent inhibitor of the adenosyltransferase activity than is PPi ($7, 30, S6),and that under certain experimental conditions PPPi is bound in a manner (30) suggesting a very tight, pseudo-irreversible combination with the enzyme (37). All these facts are in agreement with the possibility that the bulk of the PPPi formed during the adenosyl transfer reaction remains bound to the enzyme surface until it has been cleaved (38). Further indication of a role for PPPi in the methionine adenosyltransferase reaction comes from studies of the reversibility of the reaction. It was found that when purified yeast enzyme was incubated with buffer, [ 8J4C] -8-adenosylmethionine, PPPi, MgCl,, and KC1 there was formation of radioactive material which passed through Dowex-50 (H') . Similar results were obtained with S-adenosylmethionine- [ 3H]adenosine, and the radioactive product was shown to migrate with authentic ATP during chromatography on Dowex-1. Replacement of PPPi by a combination of PPi and Pi led to a complete loss of this back reaction. Despite the fact that during these studies the enzyme was saturated with S-adeno36. S. H. Mudd and J. D. Mann, JBC 238, 2164 (1963). 37. W. W. Ackermann and V. R. Potter, Proc. SOC.Exp. Bid. M e d . 72, 1 (1949). 38. There is evidence that a very minor portion of the PPPc formed by the yeast enzyme does in fact dissociate from the enzyme surface prior to hydrolysis. To obtain this evidence, i t was necessary to use ATP-y-"P as substrate. With this compound, any radioactivity found in the product, PP,, is presumed to be the result of dissociation of "PPPI from the enzyme with consequent randomization of the radioactively labeled phosphate group. During such experiments =PPc was formed to an extent suggesting dissociation of .about 2% of the "PPPi. If an appreciable concentration of S-adenosylmethionine were present, less than 1% dissociation occurred ( 2 4 ) . These results are not in disagreement with those described above because previous experiments would not have been sufficiently sensitive to detect this degree of randomization of label. For example, if ATP-a-''P were the substrate, radioactivity would end up chiefly in PP,, but traces could also enter Pc as a result of cleavage of "PP, by residual pyrophosphatase, precluding detection of a small amount of '*P, formation as a result of dissociation of "PPPi.

132

S. HARVEY MUDD

sylmethionine and PPPi, the initial rate of the back reaction was only times the maximal initial rate of the forward reaction (36). Taken together, these findings allow a somewhat more detailed formulation of the reaction catalyzed by methionine adenosyltransferases

5X

+ +

ATP E methionine ATP..E..methionine ATP-E-methionine s ( -)AMe-E-PPPi (-)AMe-E-PPP, (-)AMe E PP, ATP

+ methionine 2 (-)AMe

+ + + P, + PP, + P,

(24 (2b) (24 (2)

where E indicates enzyme, ( - ) AMe indicates ( - ) -8-adenosylmethionine, and the dots represent noncovalent binding. This formulation is in agreement with the evidence (to be reviewed below) which suggests that the immediate act of adenosyl transfer involves transition from a ternary complex, ATP- .E. Smethionine, to the alternative complex, (-)AMe. E * * PPPi. S-Adenosylmethionine is represented here as bound to the enzyme.-PPP+ complex. Further evidence for the binding of ( - ) -8-adenosylmethionine to the adenosyltransferase has been provided by experiments involving gel filtration under nonequilibrium conditions (24, 28), by kinetic studies of inhibition of the overall reaction by this sulfonium compound, and by demonstration of the stimulation of the tripolyphosphatase activity of the adenosyltransferase by S-adenosylmethionine (24, 27,30).The presence of S-adenosylmethionine in association with the enzyme. .PPPi complex exerts an important effect as a result of the ability of S-adenosylmethionine to stimulate the tripolyphosphatase activity. The rates of tripolyphosphate cleavage are such that, in the absence of S-adenosylmethionine, the tripolyphosphatase step would proceed a t a rate of 20% or less of the rate a t which the enzyme is potentially capable of forming S-adenosylmethionine and PPPi (27,28, 30). Tripolyphosphate cleavage under such conditions would be severely rate limiting in the overall reaction. Conversely, in the presence of optimal S-adenosylmethionine, tripolyphosphate is split about twice as fast as the potential rate of the overall reaction (27, 28, 30).It can be calculated that under these conditions the tripolyphosphate cleaving step occupies only about half the time spent by an enzyme molecule during a complete catalytic cycle (36); that is, the tripolyphosphatase step is no longer severely rate limiting when the enzyme is optimally activated by S-adenosylmethionine. A lag in the rate of S-adenosylmethionine formation, first reported in 1963 ( 3 6 ) ,has recently been studied extensively by Chou and Talalay (30).This lag is presumed to result from transition from a situation in which tripolyphosphatase activity is limiting a t early times of incubation (before S-adenosylmethio-

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nine has accumulated) to a situation in which, a t later times, sufficient S-adenosylmethionine has accumulated in the reaction medium to optimally stimulate tripolyphosphatase activity and thus, to a great extent, relieve the rate-limiting effect of tripolyphosphate hydrolysis. In support of this hypothesis, it has been observed that the lag may be largely overcome by addition of S-adenosylmethionine to the initial reaction mixture (30, 36, 39). The adenosyltransfer reaction itself is visualized in the model outlined here as a transition on the enzyme surface in which the adenosyl moiety passes from ATP directly to methionine without the intermediary formation of adenosyl-enzyme. Several lines of evidence suggest that in thp case of methionine adenosyltransferase there is no adenosyl-enzyme complex. To investigate the possible occurrence of such a complex, a search for 32PPPiexchange into ATP is not likely to be helpful since the fact that the major portion of PPPi does not ever leave the enzyme surface prior to hydrolysis would be expected to preclude detection of 32PPPi-ATP exchange. However, formation of an adenosyl-enzyme complex should lead to an exchange of methionine into S-adenosylmethionine, and such an exchange was not detected with the yeast enzyme ( 1 4 ) . A direct search for an adenosyl-enzyme complex by experiments involving gel filtration produced negative results ( 2 4 ) . Finally, Greene (27) and Chou and Talalay (SO),observed convergent lines when reciprocal reaction velocity was plotted against reciprocal concentration of either methionine or ATP with the other substrate being maintained a t several fixed concentrations. Each interpreted these results as suggesting a mechanism in which both substrates must bind to the enzyme prior to formation of any product (27, 30). Were adenosyl-enzyme an intermediate, some formation of PP and P, might be expected prior to binding of methionine.

3. Activators and the Effect of p H Methionine adenosyltransferase requires both a divalent (10, 14) and a monovalent (14) cation for activity. With the enzyme purified from yeast the effects of these agents are rather complicated functions of the 39. This explanation of the lag in synthesis of S-adenosylmethionine and the ability of added S-adenosylmethionine to largely overcome this lag is valid if one or more of the following conditions pertains : (a) S-Adenosylmethionine dissociates from, and reassociates with, the enzyme. -PPPc complex rapidly relative to the rate of tripolyphosphate cleavage ; (b) more than one molecule of S-adenosylmethionine must be bound to each enzyme.-PPP+ complex to maximally stimulate tripolyphosphate hydrolysis ; or (c) in order to stimulate tripolyphosphatase activity S-adenosylmethionine must be bound to the enzyme at a site other than the one at which this compound is generated as a result of the adenosyl transfer step.

134

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pH of the reaction mixture, and the concentrations of ATP and other cations present. At pH 7.6, and an ATP concentration of 0.02M, halfmaximal reaction rates are attained a t concentrations of 6 mM Mg2+and 20-30 mM K+. Inhibition by excess cations is not observed, and reaction mixtures containing concentrations of 0.2-0.3 M , or higher, of MgCl, and KCl have often been used (14, SO, 4U). If the ATP concentration is lowered, or the pH of the reaction mixture is raised to 9, excess Mg2+becomes inhibitory. This inhibition is partially overcome by K+ (27).At pH 9,use of a medium containing 0.1 M KCl, and sufficient Mg2+to form a 1 : l complex with ATP and to leave a 5 mM excess has been recommended (27'). At pH 7.6, Mn2+can almost completely replace Mg2+ (14,27).A number of other divalent cations are less effective or inhibitory (14). A t pH 9, Mn2+is less effective than Mg2+,but addition of small amounts of Mn2+ to solutions containing an optimal concentration of Mg2+synergizes the activity by approximately 50% (27).The monovalent cation requirement a t pH 7.6 can be satisfied by NH,+ or Rb+ as well as K+.Considerably less satisfactory are Na+,Lit, or Cst (14). The tripolyphosphatase activity associated with the yeast adenosyltransferase has a Mg2+requirement similar to that of the overall reaction (27).At pH 7.6 the hydrolytic activity displays a 5-6-fold stimulation resulting from added K+ or NH,+ (@), whereas a t pH 9 only a 2-3 fold stimulation is observed (27).Greene noted, also, that, in contrast to the adenosyltransferase activity, the tripolyphosphatase activity was little affected by changes of pH between 6 and 8.5 and failed to show a maximum a t pH 9 (97). Methionine adenosyltransferase of liver appears to be more specific in its requirements for Mg2+than is the corresponding yeast activity. The patterns of response to monovalent cations are generally similar for the two enzymes. The liver enzymic activity is almost completely dependent upon the presence of a sulfhydryl compound, for example, glutathione (14).The latter compound stabilizes the yeast enzyme to heat inactivation (14) but has no effect upon activity measured under the usual conditions (27,30). Reports of extensive searchs for specific inhibitors of methionine adenosyltransferase activity have not been published. The liver (14), but not the yeast (14,27) enzyme is quite sensitive to inhibition by fluoride ion. This reagent has been used to inhibit traces of pyrophosphatase present as a contaminant in some preparations of yeast adenosyltransferase (27).A rapid inactivation of the yeast enzyme by small amounts 40.

J. D. Finkelstein and S. H. Mudd, JBC 242, 873 (1967).

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of iodine has been reported, for example, 0.2 pmole I, per milligram of protein for less than 1 min a t 0” and 7.6. Under certain conditions A T P and Mg2+are said to protect the enzyme from this inactivation, but details of these observations have not been published (31).Tripolyphosphate is a potent inhibitor of methionine adenosyltransferase. The results of studies of this inhibition, as well as inhibition resulting from products formed by the enzyme (PPi and S-adenosylmethionine) , and inhibition resulting from a number of structural analogs of methionine or ATP will be discussed in the next. section. 4. Substrate Specificities and Inhibition by Substrate Analogs

Methionine adenosyltransferase of yeast demonstrates a high degree of specificity for ATP. Adenosine tetraphosphate, ADP, 2’-deoxyATP, ITP, and G T P are inactive as substrates (14) ; GTP inhibits in a manner which is competitive with ATP (Ki= 2.6 d ) and noncompetitive with methionine (Ki= 8.8 mM) (30). It has recently been shown that both the a$- and P,y-methylene analogs of ATP are also inactive. The a$methylene compound inhibits adenosyltransferase activity. An inhibition of 45% was observed in the presence of equimolar concentrations of analog and ATP (61).Mouse liver enzyme also has a strict specificity for ATP. Adenosine monophosphate, ADP, 2'-deoxy ATP, ITP, GTP, and CTP are inactive. Uridine triphosphate sustains 4% the rate achieved with ATP (35). The specificity with respect to methionine is not as strict. The results of extensive studies of this aspect of the enzymes purified from yeast, liver, and E . coli (11, 1.6, 60J SO, 3gJ 33, 41-44) may be summarized as follows. Active substrates are thioethers in which the sulfur atom is separated by two methylene groups from a carbon substituted in the L-configuration by a free hydrogen, an amino group, and a carboxyl (or its ester). Alteration of the number of methylene carbons, reduction of the carboxyl group, or replacement of the amino group with a hydroxyl moiety all lead to complete loss of activity. Partial or complete loss results from alkylation of the amino group with a formyl or acetyl substituent. The preferred second substituent of the thioether sulfur atom is a methyl group. Replacement of this group by a hydrogen atom causes almost complete loss of activity. Replacement by ethyl decreases activity, 41. S. H. Mudd and G . L. Cantoni, Nature (London) 180, 1052 (1957). 42. A. Peterkofsky, in “Transmethylation and Methionine Biosynthesis” (S. K. Shapiro and F. Schlenk, eds.), p. 136. Univ. of Chicago Press, Chicago, Illinois, 1965. 43. J. A. Stekol, in “Transmethylation and Methionine Biosynthesis” (S. K. Shapiro and F. Schlenk, eds.), p. 231. Univ. of Chicago Press, Chicago, Illinois, 1965. 44. R. Cox and R. C. Smith, ABB 129, 615 (1969).

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S. HARVEY MUDD

most markedly for the enzyme from E . coli. Compounds containing higher substituents in this position are probably almost completely inactive ( 4 3 ) . The trifluoromethyl analog has been reported to be a substrate ( 4 3 ) ,but this finding could not be confirmed in a latter study (33).The sulfur atom itself may not be oxidized to a sulfoxide or sulfone but may be replaced with selenium. The resulting compound, selenomethionine, sustains a reaction rate with yeast enzyme approximately 1.8 times that observed with methionine (8'7,41). With selenomethionine as substrate, the pH vs. activity curve is virtually identical to that for tripolyphosphatase activity, leading Greene to suggest that in this case the overall reaction is limited solely by the tripolyphosphatase activity and the adenosyl transfer reaction itself may proceed relatively rapidly (2"). Many studies have been performed of inhibition of methionine adenosyltransferase activity by structural analogs of methionine (20, 29, 30,33,34).In the most thorough and extensive studies of this type, carried out by Lombardini, Talalay, and their colleagues, use was made of analogs restricted in conformation by unsaturation or by ring formation and of analogs endowed with varying degrees of electronegativity in the region of the molecule corresponding spatially to that occupied by the sulfur atom of L-methionine. The trans, but not the Cis, isomer of D , L - ~ amino-Chexenoic acid is a potent inhibitor ( K i = 14.3 mM) (45). The corresponding acetylenic compound (L isomer) is almost four times more inhibitory ( K i = 4.0 mM) . L-Norleucine, the analogous saturated amino acid is 3 4 times less active, and the 5-carbon analog, L-norvaline, is a very poor inhibitor. It was inferred that a t the active site on the enzyme methionine lies in a conformation in which the spatial relationships between its amino and carboxy groups, the sulfur atom and the methyl moiety correspond respectively to the amino and carboxy groups, the electronegative zone of unsaturation, and the terminal methyl group of 2-amino-trans-4-hexenoicacid. In this conformation the sulfur atom and the terminal methyl group of methionine are in a somewhat extended position rather than curled back toward the remainder of the molecule. Both the methyl group and the electronegativity of the sulfur are thought to contribute to efficient binding. It was further observed that l-aminocyclopentane carboxylic acid is a strong inhibitor (Ki = 6.7 mM). Ring size is critical for satisfactory inhibition. These results provide further support for the conformation of L-methionine in question since the car45. For this compound, as for others for which values for Kc are presented here, inhibition waa shown to be competitive vs. cmethionine. With respect to ATP, inhibition was noncompetitive and, in each case, the Kt value was higher, providing evidence that the inhibitors in question are specific conformational analogs of me thionine .

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boxyl and amino groups and the methylene carbons in this conformation show good spatial correspondence to the similar groups and ring carbons of 1-aminocyclopentane carboxylic acid. Finally, Lombardini et a2. (33) found that, whereas L-serine and L-cysteine are weak inhibitors, homoserine and homocysteine are stronger, and 0-acetylserine and 0-carbamylserine are rather good inhibitors. These results were interpreted on the assumption that inhibition is generally proportional to the ability of a compound to provide an electronegative center in a position closely corresponding to that occupied by the sulfur atom of methionine, a postulate which also agrees with the model based on the results obtained with 2-amino-4-hexenoic acid and related compounds. 5. Kinetics

Kinetic aspects of the reaction catalyzed by methionine adenosyltransferase have been studied by a number of workers. Unfortunately, S-adenosylmethionine exerts opposing effects on the rate of this reaction. On the one hand, as discussed above, presumably as a result of stimulation of the tripolyphosphatase activity, low concentrations (i.e., below approximately 0.05 mM) of S-adenosylmethionine may increase the rate of its own synthesis in the overall reaction, On the other hand, higher concentrations of S-adenosylmethionine decrease reaction rates as a result of product inhibition (24, 27, SO). The difficulties posed by this situation are illustrated by the careful work of Chou and Talalay (SO). These authors noted sizable deviations from linearity of Lineweaver-Burk plots of reciprocal initial velocity with respect to reciprocal concentration of either methionine or ATP. The deviations were such as to suggest that a t high concentrations of either substrate the velocities become abnormally high. A reasonable explanation for these deviations is provided by the stimulatory effect of S-adenosylmethionine on its own synthesis. At best, therefore, in the studies published until now a steady state kinetic analysis of methionine adenosyltransferase has been only approximated. Considerable caution is indicated in interpretation of such studies, most especially those in which the S-adenosylmethionine concentrations in the medium are likely to have been passing through the critical range of concentrations during a significant portion of the experimental period. Of the two most recent and extensive studies of the kinetics of yeast methionine adenosyltransferase, one was carried out with a spectrophotometric assay (27).The sensitivity of this assay makes it likely that, at least in some experiments, 8-adenosylmethionine accumulated to concentrations above 0.05 mM. Chou and Talalay used a more sensitive assay and calculated kinetic constants by approximating regression lines

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S. HARVEY MUDD

to their experimental curves determined a t low substrate concentrations (SO). In spite of these differences in experimental design, the K , values calculated as a result of these two studies are in good agreement. For L-methionine, the values were 0.45-0.55 mM a t pH 7.6 and high ionic strength, and 0.31-0.42 mM a t pH 9 and lower ionic strength. For ATP, the corresponding K , values were 0.62-0.80 and 0.28-0.36 mM. Of the mammalian adenosyltransferases, the rat liver enzyme has been reported to have a K , for L-methionine of 0.56 (44) or 0.91 mM (39) and, for ATP, a K, of 2.3 mM (4.4). The rabbit liver preparation has a K , for L-methionine of 2.2 mM (14). Attempts have been made to utilize kinetic studies of product inhibition according to the methods of Cleland (46) to determine whether there are ordered sequences of addition of substrates to, and release of products from, the yeast enzyme. Tripolyphosphate is the most potent known inhibitor of methionine adenosyltransferase. Both Greene (27) and Chou and Talalay (SO) found the inhibition to be competitive with ATP ( K i = 1.4 X 10M5M)and noncompetitive with L-methionine. Greene observed that PPi exerts the same type of inhibition but is considerably weaker (Kivs. ATP = 7.1 X 10-4M).The inhibition resulting from Pi is too weak to be accurately measured without complications resulting from formation of insoluble Mg2+complexes. These inhibition patterns, as well as the inhibition by GTP which is competitive with ATP (SO), are most plausibly interpreted according to a model involving sequential addition of substrates, with ATP binding first to the free enzyme (27, SO). However, the observations of Chou and Talalay that a number of “deadend inhibitors” which are structural analogs of methionine inhibit competitively vs. L-methionine, but noncompetitively vs. ATP suggested to these authors a model involving the opposite order of addition, i.e., addition of methionine to free enzyme. To reconcile these observations, Chou and Talalay suggested a random-ordered sequence in which either L-methionine or ATP may be the first substrate (SO). With regard to possible ordered sequences of product release, Greene interpreted his data on PPi inhibition as suggesting this product was the last to be released. This author observed also that S-adenosylmethionine inhibited in an uncompetitive manner when either ATP or methionine was the variable substrate, suggesting to him that S-adenosylmethionine is the second product to be released and that S-adenosylmethionine does not bind to the free catalytic site (27). As has been emphasized by Chou and Talalay, the biphasic action of S-adenosylmethionine complicates 46. W. W. Cleland, BBA 67, 104, 173, and 188 (1963).

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analysis of the effects of this compound (So).The role of such complications in Greene’s experiments is not clear. The suggestion that S-adenosylmethionine does not bind to the free catalytic site (27)appears to be at odds with the binding of S-adenosylmethionine by free enzyme observed directly during gel filtration experiments (24,68),although it is, of course, possible that in the latter studies S-adenosylmethionine was binding a t a location other than the catalytic site. This possibility, namely, that there is more than one site for S-adenosylmethionine binding to yeast methionine adenosyltransferase, was invoked also by Chou and Talalay to exT plain the fact that this sulfonium compound first stimulates, then inhibits, the hydrolysis of tripolyphosphate, a reaction in which S-adenosylmethionine is not obviously either a substrate or a product (SO). An alternative explanation of this observation was set forth by Greene (27), who proposed a model involving obligatory association of S-adenosylmethionine with, and dissociation from, the enzyme during each catalytic tripolyphosphatase cycle. Formally, this is equivalent to treating S-adenosylmethionine as both a substrate and a product and yields rate equations predicting both stimulatory and inhibitory effects, as actually observed. Thus, in these recent studies no agreement appears to have been reached as to possible ordered sequences of substrate addition or product release. This problem, and questions as to the location and number of sites for S-adenosylmethionine binding, the requirements for binding, and the effects of this compound, once bound, stand out as aspects requiring more investigation before it is possible to provide a more detailed and convincing picture of the mechanism of methionine adenosyltransferase. 6. Energetics

On the basis of enthalpy changes measured during methyl transfer reactions, Durell and Cantoni were the first to calculate that sulfonium bonds are likely to be among those which have high free energies of hydrolysis ( 4 7 ) . Similar studies were subsequently extended to S-adenosylmethionine itself (48) and make it appear that, if the assumptions of the original analysis are correct, the sulfonium links of S-adenosylmethionine are, indeed, “high-energy” bonds. It was therefore of some interest to understand why the reversal of the enzymic synthesis of S-adenosylmethionine proceeds so slowly. With all substrates saturating, adenosyl transfer from S-adenosylmethionine to PPPi to form ATP occurs 5-6 orders of magnitude less quickly than does the adenosyl transfer from 47. G. L. Cantoni, Comp. Biochem. 1, 181 (1960). 48. S. H. Mudd, W. A. Klee, and P. D. Ross, Biochemistry 5, 1653 (1966).

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ATP to methionine to form S-adenosylmethionine. Mudd and Mann (36) pointed out that this reverse reaction could be considered as the sum of three steps:

+ +

(-)AMe E PPPi e (-)AMe-E..PPPi (-)AMe-E..PPPi ATP..E..met,hionine ATP-.E..methionine: ATP E methionine (-)AMe

+ PPP,

+ + ATP + methionine

(3%) (2b) (24 (3)

It seems likely that in both the forward and reverse reactions, as written, the actual adenosyl transfer on the enzyme surface, step (2b), is rate limiting ( 3 6 ) .If so, the relative rates of the forward and backward reactions, a t saturation, will be determined by the free energy change a t this step. This free energy change, AFLb, may be calculated as the sum of -AF:,, the total energy necessary to dissociate PPPi and S-adenosylmethionine from their ternary complex with the enzyme, AF’,, the free energy of adenosyl transfer from S-adenosylmethionine to PPPi, and -Mi,, the energy of binding ATP and methionine to the enzyme to form a ternary complex. The free energy changes in question (all in kilocalories per mole) were approximated as follows. The free energy of dissociation of S-adenosylmethionine from PPPi * *enzymewas calculated as 6.0 on the basis of the concentration of S-adenosylmethionine required to stimulate the tripolyphosphatase activity to 50% of maximum. The energy of dissociation of PPPi from enzyme was estimated as 9.4. This estimate was arrived a t by taking into account the K , for PPPi in the tripolyphosphatase reaction and the amount of PPPi dissociating from the enzyme prior to hydrolysis (38).Combining these two energies of dissociation, -AF:, = 15.4. On the basis of literature values, A F ’ ~was calculated to be 0.6. The free energy of binding ATP to enzyme was estimated as - 3.7,based on the inhibition of the tripolyphosphatase activity by ATP. The energy of binding methionine to ATP..enzyme was estimated as -3.8, assuming the K , for methionine in the overall methionine adenosyltransferase reaction is equal to the dissociation constant in question. Together, these two binding energies permit calculation that -AF;~= -7.5. Then, AFLb = 15.4 0.6 - 7.5 = 8.5. On this basis it was predicted that the initial rate of the forward reaction a t saturation would be 1.1 X lo8 times the initial rate of the back reaction a t saturation, an estimate which, within the limits of uncertainty of the estimates, agreed with the experimentally determined ratio of 0.5 X lo6 ( 3 6 ) .This admittedly approximate analysis suggests that the major reason for the very slow reversal is to be found in the tight binding of S-adenosylmethionine and PPPi to the enzyme. In the reverse reaction these binding

+

4. THE

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energies make unfavorable contributions which are not compensated by the favorable contributions resulting from binding of ATP and methionine. The result is that the enzyme, in effect, catalyzes a virtually irreversible reaction which allows accumulation of S-adenosylmethionine. The tight binding of the product, PPPi, however, creates a reciprocal problem. This compound is a very strong inhibitor. This inhibition is avoided in the present instance by hydrolysis of PPPi to PPi and Pi, compounds which are much less strongly bound and therefore commensurately less inhibitory ( 4 9 ) .

E. REGULATION AND GENETICS 1. Microorganisms

Although studies of regulation of methionine adenosyltransferases are as yet in their early stages, there is already considerable diversity in the patterns reported for various microorganisms. Addition of methionine to the growth medium induces an increase in adenosyltransferase activity in the yeast, Saccharomyces cerevisiae ( 5 0 ) ,but represses the level of this enzyme in E. coli K-12 ( 6 1 ) . Constitutive mutants of E. coli K-12 have been isolated. These strains have abnormally high, and poorly repressible, concentrations not only of methionine adenosyltransferase but also of such enzymes involved in methionine biosynthesis as cystathionine-ysynthase and cystathionase (51, 6 2 ) . These changes result from mutation a t a single genetic locus, suggesting that the regulatory systems for these three enzymes, and perhaps for others of the methionine biosynthetic pathway, have a t least some elements in common ( 5 3 ) . Mutants of both the mold, Neurospora c~assa(54,56), and E . coli K-12 (66) have been described which have abnormally low specific activities of methionine adenosyltransferase. Such mutants may contain altered 49. Some of the estimates of free energy changes used in this analysis could now be revised on the basis of more recent experimental results. Such revisions have not been made here since the relatively small changes required would fall within the uncertainty of the original values and the thrust of the argument would remain unaltered. 50. C. J. Pigg, W. A. Sorsoli, and L. W. Parks, J. Bacteriol. 87, 920 (1964). 51. C. T. Holloway, R. C. Greene, and C. H. Su, J. Bacteriol. 104, 734 (1970). 52. C. H. Su, R. C. Greene, and C. T. Holloway, Bacteriol. Proc. p. 136 (1970). 53. C. H. Su and R. C. Greene, Proc. N a t . Acad. Sci. U.S. 68, 367 (1971). 54. D. Kerr and M. Flavin, BBA 177, 177 (1969). 55. D. S. Kerr and M. Flavin, JBC 245, 1842 (1970). 56. R. C. Greene, C. H. Su, and C. T. Holloway, BBRC 38, 1120 (1970).

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structural genes for this enzyme (56, 57) or, in some instances, the situation may be more complicated ( 5 7 ) . In any case, these strains have been useful in defining the regulatory role of S-adenosylmethionine itself since they contain abnormally low pools of this compound (54, 5 6 ) . Such mutants of both N . craSsa and E . coli K-12 contain unusually high activities of cystathionine-y-synthase as well as, perhaps, other methionine biosynthetic enzymes. As a result, they overproduce methionine (52, 58) thereby presumably accounting for the ethionine resistance which served as the original basis for their selection. The mechanisms of these effects, however, have been shown to differ. In N . crassa, S-adenosylmethionine functions as a feedback inhibitor of cystathionine-y-synthase activity (54,55) but does not affect the level of this enzyme itself. By contrast, in E. coli K-12, S-adenosylmethionine, or one of its metabolites, represses accumulation of cystathionine-y-synthase and cystathionase ( 5 6 ) . Reference has already been made to additional studies of the regulatory effects of S-adenosylmethionine upon other enzymes of the methionine biosynthetic pathway as well as upon apparently unrelated enzymes (20, 2 1 ) . The biphasic action of S-adenosylmethionine on the rate of its own synthesis by yeast methionine adenosyltransferase has been discussed in previous sections of this chapter. Depending upon its concentration, S-adenosylmethionine may stimulate or inhibit. The role of such effects in various physiological states of intact yeast remains to be clarified as does the question of whether similar phenomena are manifested by the methionine adenosyltransferases of other organisms.

2. Mammals a. Effects of Age. Methionine adenosyltransferase activity has been reported to be present only in trace amounts in livers of fetal rabbits, mice ( 3 5 ) , and rats ( 5 9 ) . The activity is low but may not be entirely absent. since the assay methods used in these studies were relatively insensitive ( 2 0 ) .Hepatic adenosyltransferase activity is present in each of these species shortly after birth (35, 6 0 ) , increasing in mouse liver until 21 days of age, more slowly in rabbit liver until 100 days ( 3 6 ) , and slowly declining in liver of rats as the animals increase in weight from 6 to 280 g.

b . Dietary Factors. Finkelstein observed that the methionine adenosyl57. R. C. Greenc, C. H. Su, and E. H. Coch, Fed. Proc., Fed. Amer. SOC. E x p .

Biol. 30, 1261 (1971). 58. S. B. Galsworthy and R. L. Metzenberg, Biochemistry 4, 1183 (1965). 59. B. Shield and E. Bilik, Cancer Res. 28, 2512 (1968). 60. J. D. Finkelstein, ABB 122, 583 (1967).

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transferase activities of rat liver, kidney, pancreas, and brain were little affected by fasting if the animals had previously been fed a standard diet. Activities increased as much as 2-fold on a low protein diet, and fasting now brought about further increases (3-fold in liver) (60). I n contrast, Pan and Tarver noted decreases in adenosyltransferase activities in livers of fasted rats or rats fed a low protein diet (61). The reasons for these discrepancies are not clear. Addition of cystine to a low sulfur amino acid diet brings about a very moderate decrease in rat liver methionine adenosyltransferase activity. The decrease is prevented if methionine is added also (40). Methionine, ethanol, or choline, added singly to normal rat diets, led to small increases (6.2). c. Hormonal Effects. Natoni reported in 1963 that liver methionine adenosyltransferase activities of female rats were about twice as high as those of males. Orchiectomy raised, and administration of androgen to castrated males lowered the activity ( 6 3 ) . Subsequently, hepatic activities in females of a number of inbred strains of mice were found t o be 2.61.2 times the activities in males of these strains. In Race I11 rabbits, however, little sex difference was detected (5.5). Finkelstein made the interesting observation that hormonal regulation of methionine adenosyltransferase, as well as other enzymes involved in the conversion of methionine to cysteine, may differ from tissue to tissue. For example, treatment of male rats with estradiol caused a 1.8-fold increase in the adenosyltransferase specific activity of liver, a 1.2-fold increase in kidney, and slight decreases in pancreas and brain (60). Glucocorticosteroid administration raises the hepatic methionine adenosyltransferase activity 1.5-2-fold in normal (60, 61) and adrenalectomized (64) rats, whereas adrenalectomy leads to a slight decrease (61). Activities in other tissues are little affected by hydrocortisone (60). Alloxan brings about a 3-fold, and growth hormone a l.&fold, increase in activity of liver but not of kidney, pancreas, or brain (60). The physiological repercussions of these, on the whole rather minor, hormonal effects remain to be worked out, although Natori has suggested that the increased methionine adenosyltransferase activity of female rats may account for the higher susceptibility of this sex t o ethionine toxicity (63) 61. F. Pan and H. Tarver, J. Nutr. 92, 274 (1967). 62. J. D.Finkelstein and W. E. Kyle, Proc. SOC.Ezp. Biol. Med. 129, 497 (1968). 63. Y. Natori, JBC 238, 2075 (1963). 64. F.Pan, G . G . Chang, S. C. Lee, and M. S. Tang, Proc. SOC.Em. Biol. M e d . 128, 611 (1968).

144

111.

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B,,,

Adenosyltransferase

A. SIGNIFICANCE AND DISTRIBUTION Following the isolation of crystalline vitamin B,, in 1948 (66, 66),and the elucidation of the complete structure of this cobalt-containing substance by X-ray analysis ( 6 7 , 6 8 ) ,it was shown by Barker and his associates that for the enzyme-catalyzed isomerization of glutamate to /3-methylaspartate a Blz-related compound was necessary, not as the vitamin itself, but rather in the form of a derivative, L‘coenzyme-B12’’ (6 9, 70) .X-Ray diffraction analysis revealed that the central cobalt atom of the crystalline coenzyme was bonded to the 5‘-methylene carbon atom of an adenosyl group ( 7 ) . Subsequent investigations have greatly expanded the list of adenosyl-BIZ-dependent enzymic reactions. A recent review discusses eleven such reactions catalyzed by bacterial enzymes ( 4 ) . On the other hand, in mammals only one adenosyl-Blz-dependent reaction, the isomerization of methylmalonyl-CoA to succinyl-CoA, is known to play a role ( 4 ) . Almost simultaneously with recognition of the presence of the adenosyl group in this coenzyme form of B,, ( 7 ), the first detailed reports appeared describing cell-free systems capable of synthesizing adenosyl-B,, derivatives (71, 7 2 ) . These enzyme systems have been purified from the bacteria, Propwnibacterium shermanii (73) and Clostridium tetanomorphum (74, 7 6 ) . Mammals are also capable of converting vitamin forms of B,, 65. E. L. Rickes, N. G. Brink, F. R. Koniuszy, T. R. Wood, and K. Folkers, Science 107, 396 (1948). 66. E. L. Smith, Nature (London) 161, 638 (1948). 67. D. C. Hodgkin, J. Pickworth, J. H. Robertson, K. N. Trueblood, R. J. Prosen, J. G. White, R. Bonnett, J. R. Cannon, A. W. Johnson, I. Sutherland, A. Todd, and E. L. Smith, Nature (London) 17% 325 (1955). 68. D. C. Hodgkin, J. Kamper, J. Lindsey. M. MacKay, J. Pickworth, J. H. Robertson, C. B. Shoemaker, J. G. White, R. J. Prosen, and K. N. Trueblood, Proc. Roy. Soc., Ser. A 242, 228 (1957). 69. H. A. Barker, H. Weissbach, and R. D. Smyth, Proc. Nut. Acad. Sci. U. S.

44, 1093 (1958). 70. H. Weissbach, J. Toohey, and H. A. Barker, Proc. Nut. Acud. Sci. lJ. S. 45, 521 (1959). 71. R. 0. Brady and H. A. Barker, BBRC 4, 464 (1961). 72. H. Weissbach, B. Redfield, and A. Peterkofsky, JBC 236, PC40 (1961). 73. R. 0. Brady, E. G. Castanera, and H. A. Barker, JBC 237, 2325 (1962). 74. A. Peterkofsky and H, Weissbach, JBC 238, 1491 (1963). 75. E. Vitols, G. A. Walker, and F. M. Huennekens, JBC 241, 1455 (1966).

4.

THE ADENOSYLTRANSFERASES

145

to adenosyl-B,, (76, 7 7 ) . Homogenates of liver and kidney (78) and extracts of HeLa cells grown in tissue culture (79) can catalyze this transformation, but the enzyme system from mammalian sources has not yet been highly purified. The physiological importance of this enzyme system for mammals is suggested by the demonstration that humans unable to accumulate adenosyl-B,, excrete abnormally elevated amounts of methylmalbnic acid (80, 81). B. THENET REACTION Early studies of the enzyme systems converting vitamin forms of B,, (HO-B,, or CN-B,,) to adenosyl-B,, revealed a complex set of requirements. I n addition to a cobamide derivative and ATP, a divalent metal ion (Mn*+),a monovalent metal ion (K+),reduced flavin adenine dinucleotide (FAD), and a sulfhydryl compound were all necessary ( 7 3 ) .It is now clear that reduced FAD was required to permit reduction of the cobalt atom of B,,. I n HO-B,, (i.e., B,,,), for example, the cobalt is in the 3' valence state. Successive one-electron reductions yield B,,, (Co2+) and BIz8(Co+).The latter is the true substrate for the adenosyltransferase; B,,, itself is not a substrate (73-75, 82). There is evidence that two enzymic reducing systems are present in crude extracts of C . tetanomorphum, one of which catalyzes reduction of B,, to B,,,, whereas the second catalyzes reduction of B,,,. to BlZs (83).I n several studies the adenosyltransferase activity has been accompanied during fairly extensive purification by a t least the B,,, reductase activity (73, 74, 85), suggesting the two may exist under certain conditions as a structural complex (84). Nevertheless, it has recently been possible to obtain preparations of the adenosyltransferase physically separated from B,,, reductase activity (75, 83). The role of ATP as the adenosyl donor in the overall reaction was elucidated by the investigations of Peterkofsky, Weissbach, and their 76. H. Uchino, Y. Yagiri, T. Yoshino, M. Kondo, and G. Wakimka, Nature (London) 2Q5, 176 (1965). 77. Y . Yagiri, J . Vitaminol. (Kyoto) 13, 228 (1967). 78. J. Pawelkiewicz, M. Gorna, W. Fenrych, and S. Magas, Ann. N . Y . Acad. sci. 112, 641 (1964). 79. S. S. Kerwar, C. Spears, B. McAuslan, and H. Weissbach, ABB 142, 231 (1971). 80. S. H. Mudd, H. L. Levy, and R. H. Abeles, BBRC 35, 121 (1969). 81. L. E. Rosenberg, A. C. Lilljeqvist, Y. E. Hsia, and F. M. Rosenbloom, BBRC

37, 607 (1969). 82. E. Vitols, G. Walker, and F. M. Huennekens, BBRC 15, 372 (1964). 83. G. A. Walker, S. Murphy, and F. M. Huennekens, A B B 134, 95 (1969). 84. A. Peterkofsky, R. Redfield, and H . Weissbach, BBRC 5, 213 (1961).

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8. HARVEY MUDD

colleagues (74,84-86). During adenosyl-B12 formation there is a transfer of one adenosyl group from ATP to each molecule of the cobalamin substrate, as shown by the use of randomly labeled [14C]ATP or [S-l*C]ATP. The stoichiometric transfer of both the adenine and the ribose portions (73, 84, 8 5 ) , as well as the fact that neither free adenine nor ribose dilute the radioactivity incorporated from ATP-14C (73) , support the hypothesis that the adenosyl group is transferred as an intact unit. The fate of the phosphate residues of ATP during adenosyl-B12 formation was clarified by these same studies (74, 86). The enzyme used had been extracted from C . tetanomorphum and subjected twice successively to chromatography on DEAE-cellulose in a manner which removed most of the contaminating ATPase. As a result, the ratio of ATPase to adenosyltransferase activity, initially 149: 1, decreased to 0.35: 1. With this relatively ATPase-free enzyme preparation, it was possible to demonstrate a B,,-dependent dephosphorylation of ATP. Throughout the course of the reaction there was a 1 :1 stoichiometric relationship between BIZ-dependent removal of radioactivity from [/3,y-"P] ATP and incorporation of [ 14C]ATPinto adenosyl-BIZ.I n contrast to the findings with the methionine adenosyltransferase, none of the radioactivity removed from [/3,y-"'P] ATP appeared as 32Pi.Fractionation of the radioactive products of the reaction on Dowex-1 (Cl-) demonstrated that almost all the B,,-dependent radioactivity released from [ / ~ , Y - ~ ~ P ] Awas TP found in "PPPi. Further proof that this material was 32PPPiwas obtained by submitting the isolated 32P-labeledmaterial to alkaline hydrolysis. Under the conditions used, the radioactive product of the enzymic reaction yielded both s2Piand 32PPjin the same proportions as resulted from alkaline hydrolysis of authentic carrier PPPi (74, 85). These findings contrast with those reported in an earlier study in which a B,,, adenosyltransferase extracted from P . shermanii was used. Although this enzyme had been purified 337-fold1 it was still contaminated with an excess of Al'Pase. When [/3,y-3ZP]ATPwas used as substrate, there was an enzyme and BI2-dependent formation of 32PPiapproximately equivalent to the dmount of adenosyl-BIZ formed. For reasons which remain obscure, there was a concomitant B1,-depei?dent formation of 3zPiseveralfold in excess of adenosyl-B12 formation (73). The paper reporting these results was submitted for publication before appearance of the studies implicating PPP, as an intermediate in the inethionine adenosyltransferase reaction (24, 28) or as a product in the BIze adenosyltransferase system of C . tetanomorphum (74, 85). I n retrospect, the observations described do not permit a decision as to 85. A. Peterkofsky and H. Weissbach, Ann. N . Y . Acad. Sci. 112, 022 (1964). 88. A. Peterkofsky, BBRC 24, 310 (1966).

4.

147

THE ADENOSYLTRANSFERASES

whether the B,,, adenosyltransferase of P. shermanii is mechanistically more closely related to the BIZ,adenosyltransferase of C . tetanomorphum or to methionine adenosyltransferase. Assay of the purified P . shermanii preparation for PPPase activity and determination of the pattern of radioactivity in the products obtained with, for example, ATP labeled in the y phosphate only, would be helpful in this respect. If PPPase activity is present but is a contaminant, [ Y - ~ ~ P I A will T P yield both s2Pi and 32PPi.If PPPase activity is present and is physically and functionally associated with the P . shermanii enzyme as is PPPase with methionine adenosyltransferase, [y-”’P] ATP should yield 32Pi as the sole radioactive product. Finally, there is an unlikely possibility that no PPPase activity is present in the P. shermanii B,,, adenosyltransferase preparation a t this stage of purification and that the mechanism of this enzyme differs from that of either of the other two known adenosyltransferases. Bearing in mind this uncertainty, the net reaction catalyzed by the systems converting vitamin forms of B,, to adenosyl-B,, may tentatively be formulated as follows:

BIZ^ Biz.

+ ATP

Bm reductase

’ Bib

Rla adenosyltransferaae 9

adenosyl-Bla

(5)

+ PPPi

(6)

Reactions (4)and ( 5 ) each are coupled to NADH oxidat,ion by flavindependent enzymes. These activities have been separated from the enzyme catalyzing reaction ( 6 ) , which represents the adenosyl transfer step per se (83).By reducing Blznchemically, for example with borohydride, it is possible to study the isolated adenosyltransferase activity without a complicating requirement for either the B,,, or B,,, reductase (76, 83).

C. PURIFICATION AND PHYSICAL PROPERTIES Several of the early studies of the adenosyl-Blz synthesizing system were complicated because enzymes capable of catalyzing the overall reaction, including the reductive steps (4) and ( 5 ) , were purified together with the adenosyltransferase. In only one study has the adenosyltransferase been obtained free of the reductase and the properties of the enzyme defined by use of an assay in which only reaction (6) was measured (75). In this study the enzyme extracted from sonicated C . tatanomorphum cells was purified as much as 300-fold by use of protamine followed by heat treatment (60” for 5 min) and two successive

148

S. HARVEY MUDD

chromatographic procedures on DEAE-cellulose. An overall yield of a%, based on the initial activity, was obtained. The purified B,,, adenosyltransferase was shown by starch gel electrophoresis to carry a moderate negative charge a t pH 7.0. The enzyme had a single absorption maximum a t 280 nm with a ratio of absorbance at 280 nm to that a t 260 nm of 1.6. The activity was easily lost in the absence of reducing agents. Mercaptoethanol, 5 X 10-3M final concentration, was added to all solutions of the enzyme during purification and storage. D. CATALYTIC PROPERTIES 1. Assay Several assays of Blzs adenosyltransferase activity have been described. As noted above, if it is desired to eliminate complications, care must be taken to supply B,,,, the true substrate for the transfer reaction. Since this compound is very easily oxidized, the B,,, must be generated in situ, for example by chemical reduction of B12, with borohydride. In the most sensitive assays, the amount of adenosyl-B,, formed is quantitated by means of a second, adenosyl-BIZ-dependent, enzymic reaction. The glutamate mutase system, measured spectrophotometrically according to Barker et al. (87), has been used for this purpose ( 7 5 ) . An adenosylcobamide containing benzimidazole in place of the 5,6-dimethylbenzimidazole moiety of adenosyl-B,, has a more favorable K , for the glutamate mutase system; thus, it may be advantageous to use benzimidazolecobamide rather than cobalamin as initial substrate for the adenosylating enzyme if this assay system is employed ( 7 1 ) .An alternative system which has been used for the enzymic assay of adenosyl-B1, is the dioldehydrase as described by Abeles et al. (88) (see, e.g., reference 79). Several simpler assays of B,,, adenosyltransferase activity have been reported but are applicable chiefly to more purified preparations: 1. Conversion of B,,, to adenosyl-B,, may be followed spectrophotometrically by measurement of increase of light-labile absorbance a t 525 nm ( 7 5 ) . 2. Transfer of radioactivity from [ l'C]ATP into [ "C] adenosyl-BIZ may be followed. The latter is easily separated from [14C]ATP by passage through a small column of Dowex-1 (Cl-) (7'4). 87. H. A. Barker, R. D. Smyth, H. Weissbach, A. Munch-Petersen, J. I. Toohey, J . N. Ladd, B. E. Volcani, and R. M. Wilson, JBC 235, 181 (1960). 88. R. H. Abeles, C. Meyers, and T. A. Smith, Anal. Biochem. 15, 192 (1966).

4. THE ADENOSYLTRANSFERASES

149

3. B,,-dependent release of radioactivity from ["PIATP may be assessed by the formation of noncharcoal-adsorbable 3zP ( 7 4 ) . This assay, of course, requires an enzyme preparation virtually free of contaminating ATPase. 2. Reversibility, Partial Reactions, and Mechanistic Considerations

Peterkofsky (86) investigated the reversibility of the adenosyltransferase step by seeking an adenosyl-B,,-dependent incorporation of 32PPPj into charcoal-adsorbable material, catalyzed by the virtually ATPase-free enzyme preparation from C. tetanomorphum. This reaction was measurable in a system composed of enzyme, 32PPPi,potassium phosphate buffer, MgCl,, mercaptoethanol, FMN, and adenosyl-B,,. The enzymic reaction was almost completely dependent upon addition of adenosyl-B,, and partially dependent upon FMN and mercaptoethanol. The charcoal-adsorbable radioactive material produced during the adenosyl-BIZ-dependent reaction was shown to contain [ szP]ATP by chromatography on Dowex-1 (Cl-) . No similar reaction occurred when 32PPi replaced 32PPPi.These findings constitute strong evidence that reversal of the adenosyl transfer reaction [Eq. (6)] had been achieved. The comparative initial rates at saturation of the forward and backward reactions were not reported, but calculations based on published (74, 86) and unpublished (89)data indicate that under the conditions used the back reaction was proceeding at at least $lo% of the initial rate of the forward reaction. This rate of reversal is very much more rapid than that attained with the yeast methionine adenosyltransferase ( 3 6 ) . As noted above, evidence for adenosyl-enzyme formation in the case of the methionine adenosyltransferase of yeast was sought but not found. However, with the B,,, adenosyltransferase of C. tetanomorphum, there is some evidence suggestive of the existence of such an intermediate (86). When the C. tetanomorphum enzyme preparation was incubated in the presence of ATP, 3zPPPi,mercaptoethanol, FMN, CN-B,,, MgCL, and potassium phosphate buffer there was an incorporation of radioactivity into charcoal-adsorbable material. Omission of B,, did not diminish the rate of this reaction. These data gave rise to the suggestion that an adenosyl-enzyme is formed as an early step in the adenosyltransferase reaction.

+ + + + ATP + Bla, 2 adenosyLB11 + PPPi

Enzyme ATP $ adenosyl-enzyme PPPi Adenosyl-enzyme BIZ.$ adenosyl-Blp enzyme

89. A. Peterkofsky, personal communication (1972).

(64 (6b) (6)

150

S. HARVEY MTJDD

Reversible formation of adenosyl-enzyme [reaction (6a) ] would account for exchange of s2PPPiinto ATP (86). In view of the importance of this observation in suggesting a feature unique to the B,, metabolizing enzyme, it would be well if additional studies were performed to answer several questions left open by the present evidence : 1. The enzyme used for this work was relatively crude (3-fold purified). There is thus some chance the exchange observed was catalyzed by an enzyme other than the adenosyltransferase in question. This possibility is especially pertinent because, in contrast to the other studies carried out with this enzyme preparation, in the present instance there is necessarily no B,, dependence, thus removing a powerful means of relating the activity measured to the B,, metabolizing enzyme. 2. Since 32PPPipreparations are likely to be contaminated with szPi and 32PP+, or to give rise during incubation to these compounds, there is some possibility that the observed incorporation of radioactivity into charcoal-adsorbable material resulted from an exchange of 32PPior rather than 32PPPi.A demonstration of uniform labeling in the three phosphates of the [52P]ATPformed during the exchange would meet this objection. 3. It was observed that the exchange reaction was not dependent upon mercaptoethanol addition (86). This is somewhat disturbing since mercaptoethanol is required to maintain enzyme capable of catalyzing reaction (6) (75). Of course, it is possible mercaptoethanol is required merely to maintain the capacity to catalyze the partial reaction (6b) and is not involved in partial reaction (6a). 3. Kinetics and Substrate Specificities The purified BlZsadenosyltransferase of C. tetanoniorphum has a K , M ) . Concenfor ATP of 1.6 X M (measured with B,,, a t 4 X trations of ATP above M depress activity. Adenosine triphosphate cannot be replaced by ADP, AMP (75) , or S-adenosylmethionine (90). Considerable nonspecificity with respect to the nucleoside triphosphate substrate exists since, when each was tested a t 2 X lO-‘M, CTP, ITP, UTP,and G T P supported reaction rates of 1.0, 0.53,0.39, and 0.22relative to the rate with ATP. With CTP as susbtrate, no compound supporting activity of the glutamate mutase system was formed, suggesting 90. H. Weissbach and A. Peterkofsky, personal communication to G. L. Cantoni, quoted by G. L. Cantoni, in “Transmethylation and Methionine Biosynthesis” (5. K. Shapiro and F. Schlenk, eds.), p. 21. Univ. of Chicago Press, Chicago, Illinois. 1965.

4. THE ADENOSYLTRANSFERASES

151

that cytosinyl-B,, cannot substitute for adenosyl-B,, as a cofactor for the latter system (75). The enzyme purified from P . shermanii was reported to be inactive with UTP, GTP, or ITP (73), but it is not possible to judge whether this resulted from a difference in the two bacterial enzymes or from the facts that the assay used in these studies depended upon the formation of a B,, derivative active in the glutamate mutase reaction and that nucleoside-B,, derivatives other than adenosyl-B,, may be inactive, as demonstrated above for cytosinyl-B,,. The C. tetanomorphum adenosyltransferase has a K , for BlZ8of 1 x M (measured with ATP at 1.3 X lO-'M) (75). The specificity of this enzyme with respect to the cobamide substrate was not reported. The enzyme from P. shermanii was shown to utilize CN-B,, a t a rate of 77% of the rate with HO-B,,. The hydroxo- and cyanocobamides in which benzimidazole replaced 5,6-dimethylbenzimidazolewere used a t rates of 51 and 3376, respectively. The cyanocobamide, in which an adenine group replaced the 5,6-dimethylbenzimidazole, reacted a t a rate of 133% (73). All these relative rates may have been affected by the requirement in the assay system used for an enzymic reduction of the cobalt of the cobamide substrate prior to adenosyl transfer. Of considerable interest was the demonstration that hydroxocobinamide [the corrinoid lacking all the components of the nucleotide structure "below" the ring ( I ) ] is a very satisfactory substrate, supporting a reaction rate equal to 117% that attained with HO-B,,. The adenosyl derivatives of cobinamide and other compounds containing only a portion of the cobamide structure may be the most abundant corrinoids accumulating in P. shennanii. These facts suggest that adenosyl-B,, synthesis in this organism may normally involve adenosylation of an early intermediate, such as cobinamide, to which other moieties are subsequently added to complete the adenosylcobamide structure (73). This is not likely to be the case for mammals which require as a vitamin some compound containing intact cobalamin or a closely related structure. 4. Activators and Znhibitors

The activity of the C. tetanomorphum BlZ8adenosyltransferase is stimulated about 3-fold by addition of MnY+(3 X lO-'M). In the absence of added Mnz+, activity can be abolished by addition of EDTA, 5 X 10-3M. Partial activity was obtained when Coz+,Mg2+,or Zn2+replaced &In2+,whereas none was obtained with Caz+or Cd" (75). The same general order of activities was observed with the adenosyltransferase from P. shermanii. In addition, activity of the latter enzyme was shown to be completely dependent upon the presence of K+.Ammonium ion was par-

152

S. HARVEY MUDD

tially effective in replacing K+; Li+ and Na+ were less so (73). Again, it is not possible to be sure whether these monovalent cations were acting chiefly upon the adenosyltransferase itself or upon a step in the reduction of the hydroxocobamide used as substrate in these experiments. The requirement for a reducing agent (satisfied by mercaptoethanol) for maintenance of activity of B,,, adenosyltransferase (75) has been noted above. The C . tetanomorphum adenosyltransferase activity is inhibited by PPi, PPPi, and trimetaphosphate to the extent of 80, 100, and 60%, respectively, when each compound was tested a t M. Inorganic orthophosphate did not inhibit a t this concentration (76).

IV. Conclusion

In this chapter an attempt has been made to review existing knowledge about the two enzymes known to catalyze adenosyl transfer reactions. Both the enzyme responsible for the formation of 8-adenosylmethionine and the one responsible for the formation of adenosyl-Blz are widespread in biological systems. Studies of the purified enzymes have demonstrated that there are many features common to the two reactions. I n each case ATP is the adenosyl donor. The adenosyl acceptor in the first instance is the sulfur atom of methionine; in the second instance, the reduced cobalt atom of cobalamin. The sulfur of methionine possesses an unshared pair of electrons which presumably spearhead a nucleophilic attack upon the 5/-carbon of ATP, resulting in transfer of the adenosyl moiety and displacement of PPPi. The cobalt atom of Blz, after reduction, similarly possesses an unshared pair of electrons ; thus, in this instance, too, the reaction may be visualized as a nucleophilic attack upon the 5/-carbon of ATP with PPPi serving as the leaving group. For methionine adenosyltransferase there is a good deal of evidence that adenosyl transfer takes place in a ternary complex on the enzyme surface such that the transfer is direct, i.e., from ATP to methionine without intermediary formation of an adenosylated form of enzyme. For Blz. adenosyltransferase there is some evidence suggestive of formation of an adenosyl-enzyme complex, but this evidence has yet to be subjected to critical experimental evaluation, and it is quite possible that this enzyme, also, mediates direct adenosyl transfer. Subsequent to the adenosyl transfer event itself, the reaction pathways catalyzed by the two enzymes diverge. Methionine adenosyltransferase

4. THE

ADENOSYLTRANSFERASES

153

retains PPPi firmly bound in an orientation determined by its origin in ATP. By virtue of the tripolyphosphatase activity of this adenosyltransferase, PPPi is hydrolyzed to PPi and Pi. The S-adenosylmethionine also formed during the adenosyl transfer event specifically stimulates this tripolyphosphatase activity. Thus, methionine adenosyltransferase catalyzes two sequential steps and utilizes a product of the first step to accelerate the rate of the second step. This situation is unusual, perhaps unique, among enzymes. The tight binding of tripolyphosphate to the enzyme has been shown to make a major contribution to retarding the ability of the enzyme to catalyze adenosyl transfer in the “reverse” direction (i.e., from 8-adenosylmethionine to PPPi). In effect, methionine adenosyltransferase catalyzes rapid accumulation of 8-adenosylmethionine but will not metabolize this important methyl donor. The intrinsic ability of the adenosyltransferase to cleave PPPi enables it to avoid product inhibition by this condensed phosphate. In more customary terminology, S-adenosylmethionine synthesis is obZigatori2y coupled to, or “pulled” by the favorable free energy of hydrolysis of the pyrophosphate bond of PPPi. In contrast, B,,, adenosyltransferase simply releases PPPi from the enzyme surface; thus, PPPi is a product of the overall reaction. As might be predicted, the reaction is relatively easily reversed and the enzyme catalyzes transfer of adenosyl from adenosyl-B1, to PPPi a t an appreciable rate. It remains to be established whether, in more intact biological systems, additional enzymes remove PPPi and, in so doing, “pull” adenosyl-B,, accumulation. The conclusions reviewed here as to the reaction mechanism of methionine adenosyltransferase are based chiefly upon studies of the enzyme from baker’s yeast. However, sufficient evidence is available to justify the inference that the analogous enzymes from mammalian liver and E. coli have basically similar features and that the unusual reaction mechanism of methionine adenosyltransferase, once developed biologically, has been retained during prolonged periods of evolution. It is, of course, an intriguing possibility that further search among additional biological forms will uncover a methionine adenosyltransferase which diff ers, for example, by releasing PPPi rather than hydrolyzing this compound. To date, B,,, adenosyltransferases have been purified only from two species of bacteria. Studies of the mechanism of this enzyme have therefore been limited to a relatively restricted portion of the phylogenetic map. The conclusions reviewed in this section are based on studies of the enzyme from Clostridiurn tetanomorphum. Available evidence does not permit a firm conclusion as to whether the analogous enzyme from

154

S. HARVEY MUDD

Propionibacterium shermanii releases free PPPi as does C. tetunomorphum B,,, adenosyltransferase, or hydrolyzes PPPi as do the various methionine adenosyltransferases. Finally, it is clear that the several methionine adenosyltransferases differ from one another in many relatively minor details of substrate specificity and the effects of some activators and inhibitors. Further studies may be expected to expand knowledge of such differences. Although less is known about these aspects of the B,,, adenosyltransferases, it is likely that many differences of this kind exist among this group of enzymes and are awaiting discovery. ACKNOWLEDQMENT The author wishes to thank Dr. Alan Peterkofsky for helpful discussions during preparation of the manuscript for this chapter.

Acyl Group Tranger (Acyl Carrier Protein) P. ROY VAGELOS I . Introduction

. . . . . . . . . . . . . . . . . . A . Historical Background . . . . . . B . Distribution and Intracellular Localization . C . Function in Fatty Acid Biosynthesis . . . D . Molecular Properties . . . . . . . E . Synthesis and Turnover of Prosthetic Group .

I1. Acyl Carrier Protein

. . . .

. .

. . . .

. . . . . . . .

. . . . . .

.

. . . .

187 188

. . .

188 189 190

. .

. .

I11. Malonyl CoA-ACP Transacylase . . . . . . A . Historical Background, Distribution, and Metabolic . . . . . . . . . Significance B . Molecular Properties . . . . . . . . C . Catalytic Properties . . . . . . . . I V . Acetyl CoA-ACP Transacylase . . . . . . . A . Historical Background, Distribution. and Metabolic Significance . . . . . . . . . B . Molecular Properties . . . . . . . . C . Catalytic Properties . . . . . . . . V . p-Ketoacyl ACP Synthetase . . . . . . . . A . Historical Background, Distribution, and Metabolic . . . . . . . . . Significance B . Molecular Properties . . . . . . . . C . Catalytic Properties . . . . . . . .

155 156 156 158 164

. . . . .

.

*

. .

188 173 176 176 178 179 185

185 186

.

1 Introduction

Acyl carrier protein (ACP) was discovered as a heat-stable protein in extracts of Clostridium kluyveri where it was shown to function in the 155

156

P. ROY VAGELOS

condensation reaction of fatty acid biosynthesis (1).It was subsequently isolated from Escherichia coli (2-’7), and it is the E . coli ACP which has been most extensively studied. Acyl carrier protein is involved as the acyl carrier in all the reactions of fatty acid biosynthesis (8-lo), where it has a function analogous to that of CoA in the /3 oxidation of fatty acids. Acyl group transfers involving ACP are catalyzed by three enzymes of fatty acid synthesis, acetyl CoA-ACP transacylase, malonyl CoA-ACP transacylase, and P-ketoacyl ACP synthetase. The present report will attempt to summarize the studies that have elucidated the structure and function of these three enzymes as well as of ACP. The reader is referred to the second edition of this series for a more general discussion of acyl group transfer (11) and to another review dealing specifically with acyl group transfer reactions involving CoA (12).

II. Acyl Carrier Protein

A. HISTORICAL BACKGROUND Although the initial observations on the function of ACP were made with extracts of C. kluyveri (1), the decisive experiments that demonstrated the role of this protein in the synthesis of fatty acids followed the isolation of ACP from E . coli (2-6). In the first of these experiments a condensation-decarboxylation reaction was demonstrated which involved acetyl CoA, malonyl CoA, and substrate amounts of ACP. The products of this reaction were COz, CoA, and acetoacetate bound through thioester linkage to ACP. This reaction, utilizing carboxyl-labeled malonyl CoA, is illustrated in reaction (1). P. Goldman, A. W. Alberts, and P. R. Vagelos, BBRC 5, 280 (1961). P. Goldman and P. R. Vagelos, BBRC 5, 414 (1962). P. Goldman, A. W. Alberts, and P. R. Vagelos, JBC 238, 1255 (1963). P. Goldman, A. W. Alberts, and P. R. Vagelos, JBC 238, 3579 (1963). P. Goldman, JBC 239, 3663 (1964). A. W. Alberts, P. Goldman, and P. R. Vagelos, JBC 238, 557 (1963). 7. W. J. Lennarz, R. J. Light, and K. Bloch, Proc. N a t . Acad. Sci. U . S. 48, 840

1. 2. 3. 4. 5. 6.

(1962).

8. P. W. Majerus, A. W. Alberts, and P. R. Vagelos, Proc. Nat. Acad. Sci. U . S. 51, 1231 (1964). 9. S. J. Wakil, E. L. Pugh, and F. Sauer, Proc. N a t . Acad. Sci. U.S. 52, 106 (1964). 10. P. R. Vagelos, P. W. Majerus, A. W. Alberts, A. R. Larrabee, and G. P. Ailhaud, Fed. Proc., Fed. Amer. SOC.E x p Biol. 25, 1485 (1966). 11. W. P. Jencks, “The Enzymes,” 2nd ed., Vol. 6, p. 339, 1962. 12. P. Goldman and P. R. Vagelos, Compr. Biochem. 15, 71 (1964).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

157

+ 6OOCH2CO-S-CoA + ACP-SH

CHsC0-S-CoA

CHSCOCH~CO-S-ACP + 602 + 2CoA-SH (1) Of major importance was providing proof that acetoacetyl ACP is an intermediate in fatty acid synthesis, and this proof came from the experiment illustrated in Fig. 1. As shown here, when acetyl CoA and radioactive malonyl CoA (labeled in both the free and esterified carboxyl carbons) were incubated with ACP and the E. coli fatty acid synthetase, radioactive acetoacetyl ACP (5610 cpm), was formed. When this radioactive acetoacetyl ACP was subsequently incubated with unlabeled malonyl CoA, TPNH, and the partially purified fatty acid synthetase, radioactive vaccenic acid was formed. Vaccenic acid is a major product of the E . coli fatty acid synthetase ( r ) ,and therefore it appeared that acetoacetyl ACP was converted to a fatty acid normally produced by this enzyme system. In order to be sure that acetoacetyl ACP was incorporated intact, without randomization of the isotope, into the four carbons at the methyl terminus of the vaccenic acid (bold print), the vaccenic acid was degraded, Oxidation of the vaccenic acid by permanganate-

CHSCOSCOA

+

*

*

COOHCH&OSCoA

CH,COC&&S-ACP

ll +

2CoA

+

ACP-SH

+

&I2

1

(5610 cpm)

(2~$H,COBCoA

*

CHs~2CH,~H,CH2C&CH=CHCKC&CKC&C~CH2C~C&C&COH (1300 cpm)

Vaccenic acid

Ixuno4 NaIO,

*

CH&&C&C&C&C&COOH

(1380 cpm)

+

COOHCH2C&C€I&H2CI&C&C~C&C&COOH (10 cpm)

FIQ.1. Conversion of acetoacetyl ACP to vaccenic acid. Initial incubation mixture contained [1,3-1'Clmalonyl CoA, acetyl CoA, ACP, and E . coli fatty acid synthetase preparation. [l-"Cl-Acetoacetyl ACP that was formed was isolated and then incubated with malonyl CoA, NADPH (TPNH), and E . coli fatty acid synthetase. ["CIVaccenic acid was isolated and degraded by permanganate-periodate oxidation. Radioactivity was determined in the resulting mono- and dicarboxylic acids. Carbon atoms originating in acetoacetyl ACP are shown in bold print.

158

P. ROY VAGELOS

periodate yielded a monocarboxylic acid which contained essentially all the radioactivity of the unsaturated fatty acid. Additional degradation experiments confirmed that acetoacetyl ACP was incorporated intact into vaccenic acid. It was thus established that acetoacetyl ACP is an intermediate in fatty acid synthesis in E. coli.

B. DISTRIBUTION AND INTRACELLULAR LOCALIZATION While the majority of studies attempting to delineate the structure and function of ACP have been carried out with E. coli ACP, ACP or a protein which functions like ACP has been identified in every biological system which catalyzes the de novo synthesis of fatty acids. Homogeneous preparations of ACP have been obtained from Arthrobacter, avocado, spinach (13), Mycobacterium phlei (14), Clostridium butyricum (16), and yeast (18).The amino acid compositions of ACP’s from these sources are shown in Table I (13,15-18). The most striking feature of these compositions is the presence of 1 mole each of taurine (the oxidation product of 2-mercaptoethylamine) and /3-alanine, representing the 4‘-phosphopantetheine prosthetic group (see below). Five of the seven ACP’s have only a single sulfhydryl group, that contributed by the prosthetic group. Avocado and yeast ACP both have an additional sulfhydryl group contributed by a cysteine residue; however, Simoni et al. (1s) have shown conclusively that the cysteine of avocado ACP does not function as an acyl carrier. All the proteins except that derived from yeast are rich in glutamic and aspartic acid residues, thus accounting for the acidic nature of the proteins. Molecular weights of these proteins range from approximately 8600 for C. butyricum ACP (16) to 16,000 for yeast ACP (18). The report of the isolation of yeast ACP represents the first isolation of ACP from one of the multienzyme complexes. Very small amounts of material were available for amino acid composition studies, and therefore the molecular weight estimation, which was based on the amino acid composition, must be considered only tentative (18).Future work on ACP from multienzyme complexes may reveal the reasons for the extremely tight association exhibited by these systems. Examination of the amino acid composition of yeast ACP reveals some important differences from ACP’s isolated from the easily dissociated fatty acid synthetase systems. 13. R. D. Simoni, R. S. Criddle, and P. K. Stumpf, JBC 242, 573 (1967). 14. S. Matsumura, BBRC 38, 238 (1970). 15. G. P. Ailhaud, P. R. Vagelos, and H . Goldfine, JBC 242, 4459 (1967). 16. T. C. Vanaman, S. J. Wakil, and R. L. Hill, JBC 243, 6420 (1968). 17. S. Matsumura, D. N. Brindley, and K. Bloch, BBRC 38, 369 (1970). 18. X. Willecke, E. Ritter, and F. Lynen, Euro. J . Biochem. 8, 503 (1969).

c”

TABLE I AMINO ACID COMPOSITION OF ACP FROM SEVERAL SOTJRCES~.~ Amino acid

E. colic

Arthrobacterd Avocadod

Spinachd

M. p h l d

C. butyricumf

Yeast0

4 d -4

r 0

Cysteic acid Taurine j3-Alanine Aspartic Threonine Serine Glutamic Proliie Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lysine Histidine Arginine

0 1

1 9

6 3 18 1 4

7 7 1

7 5 1

2 4 1 1

0 1 1

14 2 6 10

1 1 1

0 1

0 1

0 1

1

1

1

1

1

12

12 6 5 16 2 4 9 7 1 5

11

13 1 3 14 1 1 6

14 8 12 19

7

1

10 22 3

5

7

12 6 1 6 6 0 3 5 1 1

11

10 1

5 9 1

3 10 1 1

7 0 2 9 1 0

5 5 20 4 6 13 8 1 6 8 2

2 5 0 3

7 4 8

7 1

3 4 2 0

1

7 13 14 8 2-3 7 13 4-5 6 10 2-3 5

a Values for amino acid composition of yeast ACP were calculated assuming 1 mole of p-alanine/mole protein and rounding to the nearest integer. b Nearest integer values. Vanaman et al. (16). Smoni el a2. (13). * Matsumura et al. (17). Ailhaud et al. (16). 0 Willecke et a€. (18). J

t? d v

85 crl

2!

*

h

d

r: r

z

d

+d

3!

Z

v

CI

cn CD

160

P. ROY VAGELOS

The proportion of acidic residues is lower than that for ACP of other sources; the proportion of aromatic residues and proline is higher. Whether these differences are significant in the association of yeast ACP with the enzymes of yeast fatty acid synthetase must await further investigation. Mycobacterium phlei contains two types of fatty acid synthetase systems (17).One system exists as a tightly associated multienzyme complex of molecular weight 1.7 X lo6 (I@, which is similar to the complex isolated from yeast (ZOO).The other system consists of individual enzymes that can be purified separately, and it resembles the E . coli system in the types of reactions that are catalyzed, with one notable exception. While the E . coli system utilizes acetyl groups effectively as primer in fatty acid synthesis, the Mycobacterium nonassociated system utilizes palmityl or stearyl groups exclusively while short chain precursors are inactive (17). Therefore this fatty acid synthetase functions in chain elongation only. The amino acid composition of the ACP from Mycobacterium listed in Table I is that of ACP derived from the fatty acid synthetase which is not associated in a multienzyme complex. Whether this ACP might be derived from dissociation of the multienzyme complex has not been established (17).Acyl carrier protein from E . coli was active in the Mycobacterium fatty acid synthetase system, although some differences in rate of elongation were noted compared to Mycobacterium ACP. It should be noted that all preparations of ACP studied thus far, except yeast, have been obtained from organisms which contain fatty acid synthetase systems that are found nonassociated when the cell membrane is ruptured. Preliminary studies of yeast ACP have been reported (It?), but ACP has not been isolated from other organisms that have a fatty acid synthetase multienzyme complex. However, the presence of proteinbound 4'-phosphopantetheine has been shown in the multienzyme complexes isolated from adipose tissue (21),pigeon liver (ZZ), rat liver (23), lactating rat mammary gland (Zg), and M . phlei (19), and this implies the presence of ACP or a protein that functions as ACP in each of these systems. I n addition, the presence of ACP in the multienzyme complex isolated from etiolated cells of Euglena gracilis is indicated by the fact 19. D. N. Brindley, S. Matsumura, and K. Bloch, Nature (London) 224, 666 (1969). 20. F. Lynen, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 20, 941 (1961). 21. A. R. Larrabee, E. G . McDaniel, H . A Bakerman, and P. R. Vagelos, Proc. Nat. Acad. Sci. U.S. 54, 267 (1965). 22. P. H. Butterworth, P. C. Yang, R. M. Bock, and J. W. Porter, JBC 242, 350s (1967). 23. D. N. Burton, A. G . Haavik, and J. W. Porter, A B B 126, 141 (1968). 24. S. Smith and R. Dils, BBA 116, 23 (1966).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

161

that this complex catalyzes de novo fatty acid synthesis from acetyl CoA and malonyl CoA in the absence of added ACP ( 2 6 ) . The fatty acid synthetases of bacteria and plants are exemplified by the system studied in detail in E . coli; they are nonassociated in that the component proteins fail to show any tendency to associate in vitro after the cell membranes of the organism are disrupted. The individual soluble enzymes can be isolated by conventional means; ACP is present in the free state, associated with neither the cell membrane nor the enzymes of the fatty acid synthetase system. Yet in these fatty acid synthetases, as in the case of the fatty acid synthetase multienzyme complexes, ACP must interact specifically and consecutively with all the biosynthetic enzymes. I n spite of this, the process of fatty acid biosynthesis in E. coli is apparently very efficient since no acyl ACP intermediates are found in extracts made from cells grown in normal conditions. The possibility was considered that the fatty acid synthetase of E. coli might exhibit some kind of structural organization in vivo. Experiments were designed to localize ACP, a component of the fatty acid synthetase with a unique prosthetic group that facilitates detection, in the bacterial cell. For the localization of ACP, E . coli auxotrophs, requiring either pantothenate or p-alanine, were utilized. When these mutants were grown on limiting concentrations of radioactive pantothenate or p-alanine and allowed to remain in stationary phase for some time, the level of CoA dropped dramatically and the pantothenate or p-alanine was present almost exclusively in ACP (26,27). Initial experiments were conducted to determine if the labeled ACP was present in the periplasmic space (98). Cells, subjected to the osmotic shock treatment described by Heppel (as), retained 92% of the radioactivity inside the shocked cells, while 81% of the 5'-nucleotidase, a typical periplasmic enzyme, was released into the osmotic shock fluid. Thus, ACP is not in the periplasmic space. In order to localize ACP within the cell by electron microscopy and autoradiography, the cells were grown in the presence of p-alanine of very high specific radioactivity (5.2 Ci/mmole) in such a way that 11, 68, or 89% of the radioactivity was in ACP, the remainder being in CoA in each instance. The distribution of grains, representing p-particle tracks, was determined by electron microscopy and autoradiography. The distance from the grains to the cell surface was measured, and the data were 25. J. Delo, M . L. Ernst-Fonberg, and K. Bloch, ABB 143, 385 (1971). 26. A. W. Alberts and P. R. Vagelos, JBC 241, 5201 (1966). 27. G. L. Powell, J. Elovson, and P. R. Vagelos, JBC 244, 5616 (1969). 28. H. van den Bosch, J. R. Williamson, and P. R. Vagelos, Nature (London) 228, 338 (1970). 29. L. A . Heppel, Scknce 156, 1451 (1967).

162

P. ROY VAGELOS

FIG.2. Electron microscopy and autoradiography of tritiated cells. Washed cells were fixed in Os01 and processed for autoradiography, exposure time 6 months (288). Cells were grown in the presence of the following label: (A) [methyl-*Hlthymidine (6.7 Ci/mmole), (B) ~-['Hltryptophan (2.5 Ci/mmole), (C) ['HIP-alanine (5.2

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

163

Ci/mmole) with cells containing 11% of the 'H in ACP and 89% in CoA, and (D) ['HIP-alanine (5.2 Ci/mmole) with cells containing 89% of the 'H in ACP and 11% in CoA.

164

P. ROY VAGELOS

treated statistically. I n order to ascertain that the methods utilized would adequately delineate specific cellular areas, a number of controls were included. Figure 2A demonstrates that cells in which the DNA was labeled with [ m e t h ~ l - ~ Hthymidine ] had grains predominating over nuclear areas. Figure 2B shows that cells grown on [3H]tryptophan, in order to label all proteins in general, contained grains over the cytoplasm. In Fig. 2C are shown the cells grown on [3H]P-alanine with 11% of the 3H in ACP and 89% in CoA; in Fig. 2D are shown the cells grown in [3H]/3-alanine with 89% of the 3H in ACP and 11% in CoA. It is apparent that the grain distribution in the cells with 89% of the ["IFalanine in CoA was very similar to the distribution of grains in [3H]tryptophan cells. In other words, CoA is distributed in the cytoplasm as are the general proteins of the cell. The grain distribution in the cells with 89% of the [3H]P-alanine in ACP (Fig. 2D) is different from all the others since the majority of the grains are close to the surface of the cell. Statistical analyses of all data verified the fact that ACP is located on or near the inside surface of the plasma membrane. This location of ACP is consistent with its role in the synthesis of fatty acids and phospholipids that are present almost exclusively in the envelope of E . coli. I n addition, the fact that ACP is not randomly distributed in the cell suggests a certain type of organization for a critical component of the fatty acid synthetase. This finding suggests that the E . coli fatty acid synthetase may be organized in zlivo, perhaps in a typical multienzyme complex that includes ACP. IN FATTY ACIDBIOSYNTHESIS C. FUNCTION

The role of ACP in the biosynthesis of saturated fatty acids is illustrated in reactions (2) through (8) that have been studied in detail in the E . coli fatty acid synthetase. In the initial reaction [reaction (2)]

+ + +

+ +

CHsCO-SCoA HS-ACP CHsCO-S-ACP COA-SH (2) CHaCO-S-ACP HS-Ec,,d CH,CO-S-Ee,,d ACP-SH (3) HOOCCHzCO-S-CoA HS-ACP HOOCCHtCO-S-ACP CoA-SH (4) HOOCCHICO-S-ACP CH&O-f!&E&,d COP HS-&,,d CHaCOCHzCO-S-ACP (5) CHsCOCHCO-S-ACP NADPH H+ NADP+ CHjCHOHCHtCO-S-ACP

+ +

+

CHsCHOHCHzCO-S-ACP CHaCH=CHCO-S-ACP NADPH

+

+

+

+

+ +

+

(6)

HZ0 CHsCHICHCO-S-ACP (7) H+ + CHaCHaCHnCO-S-ACP NADP+

+

(8)

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

165

the acetyl group of acetyl CoA is transferred to the sulfhydryl group of ACP by acetyl CoA-ACP transacylase, forming acetyl ACP. The acetyl group is then transferred to the sulfhydryl group of the condensing enzyme ( E d to form an acetyl-enzyme intermediate and liberate ACP [reaction (3) 1. Malonyl CoA-ACP transacylase then catalyzes the transfer of a malonyl group from CoA to ACP in reaction (4) ; this is followed by the condensation reaction [reaction (5) 1, which takes place between malonyl ACP and acetyl-enzyme to produce acetoacetyl ACP, COz, and the free condensing enzyme. Acetoacetyl ACP is then reduced by NADPH to form specifically D- ( - ) -P-hydroxybutyryl ACP [reaction (6) ] ; the latter is dehydrated to form the trans unsaturated thioester, crotonyl ACP [reaction ( 7 ) ] ; and crotonyl ACP is reduced by NADPH to form the saturated thioester, butyryl ACP [reaction (8)1. I n the normal biosynthetic sequence, butyryl ACP reacts with the condensing enzyme to form butyryl enzyme [reaction (3) ] thereby liberating ACP, which can accept another malonyl group and thus initiate another elongation, reduction, dehydration, and reduction sequence. After appropriate repetitions of this series of reactions, the normal product palmityl ACP is formed. Small amounts of myristate and stearate are also produced, but the major saturated fatty acid produced in vivo by most fatty acid synthetase systems contains 16 carbon atoms. The in vitro products of the E . coli fatty acid synthetase are free fatty acids; this is probably a result of thioester hydrolysis catalyzed by a specific palmityl thioesterase that has been recently characterized (SO). I n vivo palmityl ACP and other long chain acyl ACP's produced by the synthetase probably react directly with sn-glycerol 3-phosphate and a specific membranous acyltransferase to form lysophosphatidic acid, the first intermediate in the pathway of phospholipid synthesis ( 3 1 ) . The similarity between the function of ACP and CoA in fatty acid metabolism is obvious. Acyl carrier protein functions as an acyl group carrier in fatty acid biosynthesis, and the acyl groups are bound covalently to a sulfhydryl group of the protein, forming thioesters. As noted above, the critical sulfhydryl group of ACP is contributed by 4'-phosphopantetheine, a covalently bound prosthetic group. I n fatty acid p oxidation, acyl groups are bound as thioesters to the sulfhydryl group of CoA, and the sulfhydryl group of CoA is contributed by 4'-phosphopantetheine. These two acyl carrier coenzymes are also related to each other biosynthetically (see below). 30. E. M. Barnes, Jr. and S. J. Wakil, JBC 243, 2955 (1968). 31. G . P. Ailhaud and P. R. Vagelos, JBC 241, 3866 (1988).

166

P. ROY VAGELOS

D. MOLECULAR PROPERTIES 1. Physical Properties

Escherichia coli ACP exhibits properties expected of a globular protein (32). The ratio of the frictional coefficient observed to the minimum frictional coefficient in aqueous solution is 1.12. The optical rotatory dispersion (ORD) curve of the protein in the ultraviolet region has a negative trough a t 232 nm, a shoulder near 213 nm, and a positive peak a t 198 nm (32),all typical of cu-helical polypeptides. The mean residue rotation a t 233 nm has been reported as approximately -4500" (32) and approximately - 8400" (33).The reason for this discrepancy is not understood. The sedimentation constant of reduced ACP is 1.34 S (32). An earlier report of 1.44s (8) probably reflected the presence of dimers of ACP resulting from disulfide bonds through the prosthetic group. Guanidine hydrochloride causes reversible denaturation of ACP. At high concentrations of denaturant (6 M ) , the protein exhibited ORD properties similar to those of a random coil, but the ORD spectrum returned to that of the native protein upon dilution of the guanidine hydrochloride to 0.6 M (39).The molecular weight of E. coli ACP, determined from the primary structure, is 8847 (16). The physical properties of M . phlei ACP resemble those of E . coli ACP. The ratio of frictional coefficients is 1.12, indicating that M . phlei ACP is also a globular protein in solution (14). The sedimentation coefficient is 1.49S, which, when used with the diffusion constant of 13.1 X lo-' cm2/sec, yielded a molecular weight of 10,450. The ORD curve of this protein also has characteristics of cu-helical proteins. The mean residue rotation a t 233 nm is -7800" ( 1 4 ) . Mycobacterium phlei ACP also exhibited ORD curves characteristic of a random coil a t high concentrations of guanidine hydrochloride. Thus, the size and physical properties of M . phlei ACP suggest that it closely resembles E . coli ACP. Numerous attempts have been made to obtain E . coli ACP crystals for X-ray diffraction studies. Adequate crystals have not yet been obtained. 2. Prosthetic Group and Primary Sequence

Sulfhydryl titrations of E. coli ACP showed that it contains 1 mole of sulfhydryl group per mole protein ( 8 ) , and the residue containing the 32. T. Takagi and C. Tanford, JBC 243, 6432 (1988). 33. D. J. Prescott, J. Elovson, and P. R. Vagelos, JBC 244, 4517 (1969).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

167

sulfhydryl group was identified as 2-mercaptoethylamine ( 3 4 ) . Other components of CoA were identified in the ACP preparation. Thus, 1 mole of ACP was found to contain 1mole each of p-alanine (8,34, 35), organic phosphate, and pantoic acid (35). The structure of the prosthetic group containing these components was established enzymically. Mild alkaline treatment resulted in loss of the prosthetic group from ACP. The alkaline cleavage product could be converted to CoA in the presence of dephospho CoA pyrophosphorylase, dephospho CoA kinase, and ATP. The structure of the prosthetic group was thus established as 4’-phosphopantetheine (35). Acyl derivatives of ACP, which are intermediates in fatty acid synthesis, are thioesters in which the sulfhydryl group of the 4‘-phosphopantetheine is esterified. The attachment of the prosthetic group exhibited acid stability and alkali lability; thus, the possibility of a phosphodiester link through a serine hydroxyl was proposed. This was established through studies of peptic peptides prepared from enzymically synthesized [2-14C]malonyl ACP. Two radioactive peptides were purified and analyzed with the following results (35, 3 6 ) . Peptide PA-3 contained 1 mole each of [14C]malonate, phosphate, pantolactone, p-alanine, taurine (the oxidation product of 2-mercaptoethylamine), and the amino acids, serine, aspartic acid, and leucine; while PA-1 contained an additional residue each of glycine and alanine. Thus, these peptides both contained a serine residue. The properties of the covalent bond joining the prosthetic group to the protein identified this linkage as a phosphate ester through the serine hydroxyl group (35, 36). The alkali stability of the phosphate ester linkage to pantetheine indicated that the phosphate is in the 4’, rather than the 2’, position of the pantothenic acid (37). The primary sequence of the isolated radioactive peptic peptide was determined by Edman degradations, dinitrophenylation, hydrazinolysis, and nitrous acid treatment, and the structure of peptide PA-1 is shown in Fig. 3 ( 3 6 ) .When this peptide was treated under mild alkaline conditions, 4’-phosphopantetheine was released. Since the phosphate of the prosthetic group is a good leaving group, a p-elimination mechanism was proposed (Fig. 4). The observation that serine disappearance was accompanied by an equivalent appearance of pyruvate supports the proposed mechanism ( 3 6 ) . 34. F. Sauer, E. L. Pugh, S. J, Wakil, R. Delaney, and R. L. Hill, Proc. Nut. h a d . Sci. U.S. 52, 1360 (1964). 35. P. W. Majerus, A. W. Alberts, and P. R. Vagelos, Proc. Nut. Acad. Sd.U.S. 93, 410 (1985). 36. P. W. Majerus, A. W. Alberts, and P. R. Vagelos, JBC 240, 4723 (1965). 37. J. Baddiley and E. M. Thain, JCS, London 246, 2253 (1951).

168

P. ROY VAGELOS

- 1I

4 '-Phosphopantetheine

-1

4'-Phosphopantothenic acid

I I

L

I1 0

---+

I I

H,C OH I I I1 I I I O-P-O-C&-C-CH-C-NH-CH2-CH2-C-NH-CH2-CH2-SH

1

I1 0

I1

I

bH

HSC

0

I

Gly-Ala- Aep-Ser-Leu

FIQ.3. The structure of peptide PA-1 isolated from a peptic digest of E . coli ACP ( 3 6 ) .

The complete amino acid sequence of E. coli ACP has been determined (16). Several features of the amino acid composition and sequence are

noteworthy. The molecule contains 14 residues of glutamic acid and 8 residues of aspartic acid out of a total of 77 residues. There are only 4 lysine residues and 1 each of arginine and histidine. Therefore, ACP is quite acidic in nature with an isoelectric point of about, pH 4.2, and a t this pH the protein is probably least soluble (16). As is apparent in Fig. 5 , the acidic residues occur throughout the sequence, while the basic residues appear clustered a t both amino and carboxyl ends. Residues 4 7 4 9 are in the sequence of Glu-Glu-Glu and 56-58 are Asp-Glu-Glu. In fact, 9 out of 14 residues between residues 47 and 60 are acidic, while there are no extended sequences rich in hydrophobic side chains. The possible exception is the sequence from residues 62 to 69. The prosthetic group is attached to serine 36 and thus lies midway along the sequence. In view of the preponderance of charged residues and the fact that ACP behaves as a typical globular protein (3%'), the three-dimensional structure may show a preponderance of charged residues on the surface (16). This may Y P

-

Y P

NH

H-!+hi2-d-

0-Pantetheine

c=o

70°C 1 hr p H 12

Leu

NH I C=CH, I

c=o

6 N HC1 100"C

NH I C=CH2 I

+

4'- Phosphopantetheine

+

Leu

Y=O Leu

0 0 II

II

CH,-C-C-OH

+

Asp

+

NH,

Leu

Fm. 4. The &elimination mechanism of removal of 4'-phosphopantetheine and acid conversion of dehydroalanine to pyruvate.

5.

169

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

1 6 10 NH,- Ser-Thr ne Clu - Glu -A=-Val -Lys-Lys - Ile Ile - Cly -Glu-

- -

-

20 Gln -Leu-Cly -Val -Lys- Cln Clu -Glu -Val -Thr-Asp-Asn- Ala -Ser

-

-

(PI-Pantetheine-SH 30 I Phe-Val -Glu -Asp-Leu- Gly -Ala -Asp -Ser Leu-Asp-Thr-Val -Glu 36

-

44

50

Leu-Val -Met -Ala-Leu-Glu -Glu -Clu -Phe - Asp-Thr-Glu

55

- Ile - Pro-

60 Asp-Clu-Glu- Ala-Glu-Lys- Ile- Thr-Thr- Val-Gln-Ala-Ala- ne

-

70 77 A s p - Q r - Ile -Asn -Cly - H i s -Cln-Ala - O H

FIQ.5. The complete amino acid sequence of E . coli ACP. From Vanaman et al. (16). Three lysine residues are underlined.

have important implications for the interactions of acyl ACP’s with the various enzymes of the fatty acid synthetase system. Detailed information on the structure of ACP’s isolated from other sources is lacking. As mentioned above, all biological systems that catalyze de novo biosynthesis of fatty acids contain protein-bound 4’-phosphopantetheine, and in all cases that have been examined the 4’-phosphopantetheine is linked to a serine residue of the protein which has the function of ACP. The amino acid sequence around the 4’-phosphopantetheine has been studied in Arthrobacter and spinach ACP’s (Table 11). Based on the results of compositional analyses and partial Edman degradations of peptides, Matsumura and Stumpf (98) suggest that 9 residues around the prosthetic group appear to be identical in Arthrobacter ACP, spinach ACP, and E. coli ACP. The similarity in the primary structure about the prosthetic group is perhaps not surprising since in all three systems the fatty acid synthetase (FAS) is the nonassociated variety. There is less structural information concerning ACP’s which are components of the tightly associated FAS complexes of yeast and animals. As mentioned above, yeast ACP has been isolated. The amino acid sequence around the prosthetic group (Table 11) (16, 98-40) appears to have no resemblance to the corresponding segment of E. coli ACP. The animal FAS is more resistant to dissociation, and the protein con-

s.

38. Matsumura and P. K. Stumpf, ABB 125, 932 (1968). 39. D. A. K. Roncari, R. A. Bradshaw, and P. R. Vagelos, JBC 247, 6234 (1972). 40. J. Ayling, R. Pirson, and F. Lynen, Biochemistry 11, 526 (1972).

170

P. ROY VAGELOS

TABLE I1 PRIMARY SEQUENCE OF 4’-PHOSPHOPANTETHEINE PEPTIDES‘ Source

Sequence of peptides

* E . CON

-ibn-Al~Ser-Ph~Val-Glu-ib~Leu-Gly-Al~AspSer-Leu-~pThr-Val-Glu-

*

Gly-Ala-ibp,Ser,Leu,Asp,Thr,Val,Glu

Arthro-

--A_-----

bud@

*

Spinach” Rat liverd

--* Leu-Gly-Ser-Leu-Asx-Leu-Gly-Glx-Cly-Glu-ibp-Ser-Leu - ---

-

Lys-Gly-Ala, -.- Asp, Ser,Leu,Asp,Thr,Val,Glu

*

Yeast.

-

Lys-GIy-Ser-Val-Pro-Ala ~~

~

The asterisk denotes the serine to which the prosthetic group, 4’-phosphopantetheine, is bound; underlined amino acids are identical with the amino acids in the corresponding sequence-of E. c o ~ACP. i * From Vanaman el a2. (16). From Makumura and Stumpf (38). d From Roncari et al. (39). From Ayling et al. (40). 4

(I

taining 4’-phosphopantetheine has not yet been isolated. Because of the difficulties encountered in the isolation of ACP from animal FAS, the amino acid sequence of rat liver peptides containing 4’-phosphopantetheine was determined. The isolation of these peptides was facilitated by the use of FAS labeled with either 3H or 14C in the prosthetic group (39). The structure around the 4’-phosphopantetheine group of rat liver FAS is shown in Table 11. Five of the 13 residues in the region of the prosthetic group of rat liver FAS are identical with those found in the corresponding region of E . coli ACP. In particular, 4 of the 5 residues around the 4’-phosphopantetheine are identical. Since the FAS of yeast and animals behave as stable complexes, strong heteromeric contacts between the constituents of these complexes are implied and substantial differences in the primary structure of the corresponding subunits relative to the corresponding bacterial and plant subunits are expected. It is not known at the present time whether the regions of the mammalian and yeast FAS which differ in amino acid sequence from those of E . coli ACP form stronger heteromeric contacts with the component enzymes of the complex. 3. Structure-Activity Relationships

Both ACP and CoA contain 4’-phosphopantctheine as the component which carries acyl groups linked as thioesters. The fact that thioesters

5.

171

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

of ACP are obligatory intermediates in fatty acid biosynthesis, whereas thioesters of CoA are relatively inactive with the biosynthetic enzymes, suggests that the protein structure of ACP is important in its biological function. With the elucidation of the primary sequence of ACP, investigations were initiated to delineate those parts of the protein structure that are important for the activity of ACP with the ten enzymes with which it is known to react. These studies have indicated that few alterations of the polypeptide chain of ACP are tolerated by the enzymes of fatty acid biosynthesis or by the enzyme, holo-ACP synthetase, which catalyzes the synthesis of holo-ACP from apo-ACP and CoA, [reaction (9)] (Section I1,E). Preparations of modified ACP were assayed in the sensitive malonyl CoA-C02 exchange reaction, which is dependent .upon two enzymes of the fatty acid biosynthetic sequence, malonyl CoA-ACP transacylase [reaction (4) 1, and P-ketoacyl ACP synthetase [reactions (3) and ( 5 ) ] ( 4 1 ) . Preparations of modified apo-ACP were tested in the holo-ACP synthetase reaction. As seen in Table 111, treatment of ACP with carboxypeptidase A led to the removal of the 3 carboxyl terminal residues of the polypeptide chain, yielding peptide 1 through 74. This peptide was fully active in fatty acid synthesis and in the malonyl CoA-C02 exchange reaction (&) . In addition, the apopeptide functioned as well as apo-ACP in accepting the prosthetic group from CoA in the presence of holo-ACP synthetase (33).The K , values for apo-ACP and apopeptide 1 through 74 are 5.5 X lo-' M and 2.0 X lo+' M , respectively. Treatment of ACP with trypsin yielded peptide 19 through 61, which TABLE I11 STRUCTURE-ACTIVITY RELATIONSHIPS OF SEVERAL PEPTIDES OF ACPa

Peptide

Method of preparation

ACP (1 + 77)

Carboxypeptidase A Trypsin CNBr Trypsin treatment of acetylated ACP

1 + 74 19 + 61 1-44 7 + 77

4

Conversion of apoActivity of holopeptide in COZ peptide to holoexchange reaction peptide in presence pmole C02/(min)/ of ACP synthetase (mg protein) (%I 6.9 7.0 0 0 0

100 100 <5

-

<5

From Majem (48) and Prescott et al. (SS).

41. A. W. Alberts, R. M. Bell, and P. R. Vagelos, JBC 247, 3190 (1973). 42. P. W. Majerus, JBC 242, 2325 (1967).

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P. ROY VAGELOS

was inactive in fatty acid synthesis and in the malonyl CoA-C02 exchange reaction (&) ; the apopeptide 19 through 61 was inactive in the holo-ACP synthetase reaction. Malonylpeptide 19 through 61 was synthesized by chemical acylation of the peptide in order to determine whether it was active in the condensation reaction of fatty acid synthesis, catalyzed by P-ketoacyl ACP synthetase [reactions (3) and (5) 3 . This enzyme was found to be completely inactive with the peptide substrate, thereby explaining the inactivity of peptide 19 through 61 in fatty acid synthesis. Acetoacetyl peptide 19 through 61 was also tested with the enzyme, P-ketoacyl ACP reductase [reaction ( 6 ) ] , and found to be only slightly active compared to native substrate. The peptide substrate had a K , of 1.7 X M and a V,, of 0.21 pmole/(min) (mg protein) compared to 6.6 X M and 1.36 pmoles/(min) (mg protein) for acetoacetyl ACP. This difference suggested that the site which confers the high reactivity of the ACP substrate was missing in the tryptic peptide. The existence of such a site was indicated by the fact that free ACP competitively inhibited (Ki = 2.2 X lO+ M ) the reduction of acetoacetyl ACP by P-ketoacyl ACP reductase, whereas the tryptic peptide did not inhibit reduction of acetoacetyl ACP. The results of the above experiments suggested that ACP interacts with P-ketoacyl ACP reductase a t a site different from the prosthetic group-active site interaction and that this site was altered or lost in the tryptic peptide (49). Other experiments investigating P-ketoacyl ACP reductase with model compounds, as well as with ACP substrates, have led the authors (4.3) to the same conclusion. Simoni et al. (IS) also concluded that there may be more than one specific site on the ACP molecule after studying ACP’s purified from several sources. Although ACP from E . coli, Arthrobacter, avocado, and spinach have strikingly similar amino acid compositions, subtle structural differences were apparent. Although the ACP from these various sources functioned interchangeably in the E . coli or plant fatty acid synthetase systems, the products of fatty acid synthesis varied according to the ACP used. This suggested that slight structural variations in ACP can alter the specificity of ACP substrates in some, but not all, reactions of fatty acid synthesis. I n addition, it has been reported that ACP of M . phlei does not function as well as E. coli ACP in the E. coli malonyl CoA-C02 exchange reaction (14). I n fact, ACP (M. phlei) is an inhibitor of ACP ( E . coli) in the fatty acid synthesizing system of E . coli. Treatment of ACP with cyanogen bromide yielded peptide 1 through 44 which is inactive. On the other hand, acetylation of the 4 lysine resi43. H. Schulz and S. J. Wakil, JBC 246, 1895 (1971).

5.

ACYL GROUP TRANSFER (ACYL CARRIER

PROTEIN)

173

dues and the amino group of the terminal serine residue had no effect on the ability of ACP to function in fatty acid synthesis (4). Trypsin treatment of fully acetylated ACP yielded peptide 7 through 77, since cleavage occurred only at the single arginine residue. This peptide, lacking the amino terminal hexapeptide, was completely inactive in fatty acid synthesis and in the malonyl CoA-C02 exchange reaction (44) ; moreover, the apopeptide was inactive in the holo-ACP synthetase reaction (33). Thus, it is obvious that the peptide structure is very important for the activity of ACP both with enzymes of fatty acid synthesis and with halo-ACP synthetase. Abita et al. (46) have attempted to extend the understanding of the structure and function of ACP by investigating the effects of chemical modifications of ACP on both its biological activity and its structure as monitored by optical rotatory dispersion measurements. Nitration of tyrosine led to a decrease in melting temperature, determined by optical activity measurements. No effect on the malonyl CoA-COa exchange reaction or fatty acid synthesis was observed, while acylation of glycerol 3-phosphate by the palmitate derivative of the modified ACP, catalyzed by a purified E. coli membrane fraction, was decreased. Alkylation of methionine had only minor effects on structure and activity measured in the malonyl CoA-C02 exchange reaction. However, fatty acid synthesis and acylation of glycerol 3-phosphate were decreased. The modification of surface carboxyl groups with glycine ethyl ester and dicyclohexylcarbodiimide led to slight structural changes, accompanied by a complete loss of biological activity. Finally, acetylation of the a-amino group strongly affected the structure, as measured by the melting temperature, but did not affect biological activity. The removal of the NH2terminal hexapeptide led to the loss of all organized structure as determined by optical measurements. This finding led the authors to propose that the terminal hexapeptide stabilizes the structure, possibly through a salt bridge with the sequence Ala-Asp-Ser-Leu of the substrate binding site.

E. SYNTHESIS AND TURNOVER OF PROSTHETIC GROUP 4’-Phosphopantetheine is a component of both CoA and ACP, and CoA is the 4’-phosphopantetheine donor in the synthesis of holo-ACP ( 4 6 ) . 44. P. W. Majerus, Science 159, 428 (1968). 45. J. P. Abita, M. Lazdunski, and G. P. Ailhaud, Eur. J . Biochem. 23, 412 (1971). 46. J. Elovson and P. R. Vagelos, JBC 243, 3603 (1968).

174

P. ROY VAGELOS

The enzyme, holo-ACP synthetase, catalyzes the synthesis of holo-ACP from apo-ACP and CoA according to reaction (9) : Apo-ACP

+ CoA M g * + holo-ACP + 3’,.5’-adenosinediphosphate +

(9)

Initial attempts to purify this enzyme from extracts of E. coli failed because of enzyme instability. After the discovery that ACP synthetase is markedly stabilized by the presence of 10-’M CoA, a 780-fold purification was achieved (46). This protective effect was specific for CoA, since CoA analogs, dephospho CoA, or oxidized CoA did not serve as protective agents. The purified enzyme was not homogeneous, but the preparation was free of apo-ACP, holo-ACP, and ACP hydrolase. The enzyme has apparent K , values of 4 X lO-’M for apo-ACP and 1.5 X M for CoA. Dephospho CoA and oxidized CoA were inactive as substrates. Although the enzyme requires either Mg2+or Mn2+for activity, both cations gave complex saturation curves, with Mgz+ showing substrate activation between lo-* and 1O-lM. The specificity of the enzyme toward apo-ACP and apopeptides is demonstrated in Table 111, where it is noted that apopeptide 1 through 74, which is active as the holopeptide in the malonyl CoA-C02 exchange reaction, is also active in the holo-ACP synthetase reaction. On the other hand, apopeptides 19 through 61 and 7 through 77 are completely inactive with the synthetase, and this parallels the inactivity of the analogous holopeptides in fatty acid synthesis. Although holo-ACP synthetase represents a very small proportion of soluble protein of E. coZi, in the order of O.Ol%, the enzymic activity is sufficient to convert 10 mpmoles of apo-ACP to holo-ACP per minute per gram of cells, and E. coli contains only 50 mpmoles of ACP per gram of cells. The intracellular concentration of CoA varies from 20 to 200 ,.& depending I on growth conditions ( 4 7 ) , and while these concentrations would not be saturating for the enzyme ( K , for CoA about 0.15 mM) , the intracellular enzymic activity would be sufficient to account for all synthesis of ACP in exponentially growing cells (46). Another enzyme that is involved in the metabolism of ACP is ACP hydrolase, which catalyzes the removal of 4‘-phosphopantetheine from ACP according to reaction (10) :

-

+ H20 Mn*+ 4’-phosphopantetheine + apo-ACP

(10) The enzyme is cytoplasmic and has been partially purified from extracts of E. coli (48).One reaction product, apo-ACP, was isolated and identiHolo-ACP

47. G. M. Brown, JBC 234, 379 (1959). 48. P. R. Vagelos and A. R. Larrabee, JBC 242, 1776 (1967).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

175

fied by amino acid composition analysis. The other product, 4’-phosphopantetheine, was identified by demonstrating that it was converted to CoA by the enzymes, dephospho-CoA pyrophosphorylase and dephosphoCoA kinase. Acyl carrier protein hydrolase is completely dependent on divalent metal cations for activity with manganous ion giving maximal activity a t 2.5 X M . Other divalent metal cations such as magnesium, cobalt, iron, and zinc were effective activators a t higher concentrations. However, CuSOa, CdCl,, and CrCI, did not stimulate the reaction at any concentration tested, nor were trivalent metal cations, such as FeC13 or AICl,, effective. The most striking characteristic of ACP hydrolase is its specificity for hydrolysis of the phosphodiester of intact ACP. The enzyme is inactive with large peptides of ACP, including the tryptic peptide 19 through 61 described above. Neither CoA nor glycerylphosphorylserine was split by the enzyme. However, ACP from C. butyricum did serve as the substrate for ACP hydrolase from E. coli, although the rate, of hydrolysis was approximately one-third that observed with E. coli ACP (48).Thus, the enzyme is a phosphodiesterase with strict specificity for ACP. The discovery of the two enzymes, holo-ACP synthetase and ACP hydrolase, led to a series of experiments that attempted t o assess the in vivo significance of the reactions that they catalyzed. Since pantothenic acid is a component of 4’-phosphopantetheine, an E. wli pantothenate auxotroph was utilized in kinetic isotope experiments with radioactive pantothenate ( 2 7 ) . The results of these studies indicated the following relationship between the cellular pantothenate-containing compounds : 4‘-phosphopantetheine+ dephospho CoA -+ CoA -+ ACP .--f 4’-phosphopantetheine

CoA is synthesized from 4’-phosphopantetheine by the established series of reactions ( 4 7 ) ,and CoA is the obligatory precursor of holo-ACP. Thus, all of holo-ACP synthesis in vivo is dependent upon the enzyme, holoACP synthetase. The pulse and pulse-chase experiments also demonstrated the in vivo hydrolysis of ACP to 4’-phosphopantetheine that is catalyzed by ACP hydrolase. The turnover of the prosthetic group of ACP, demonstrated in exponentially growing cells, caused 47% of the cellular ACP to be hydrolyzed to 4’-phosphopantetheine and apo-ACP per minute. Since in exponentially growing cells the increase in mass of the ACP compartment is 1% per minute, i t appears that the rate of ACP synthesis is about five times what it would be if turnover did not occur. Pulse experiments have also been performed with radioactive pantothenic acid in animals. Tweto et al. (49) found with the rat liver 49. J. Tweto,

M.Liberti, and A. R. Larrabee, JBC 246, 2468 (1971).

176

P. ROY VAGELOS

fatty acid synthetase complex that protein turnover of the enzyme complex is much slower than the turnover of the 4'-phosphopantetheine prosthetic group. In steady state conditions rat liver FAS exhibited a half-life of 71-108 hr. The exchange rate of covalently bound 4'-phosphopantetheine in the complex was a t least an order of magnitude greater. Thus, prosthetic group turnover has been observed both in ACP of E. coli, which has a nonassociated fatty acid synthetase, and in the FAS of rat liver, which is a tight multienzyme complex. It should be mentioned that ACP hydrolase has not yet been identified in animal tissues. Although the turnover of the 4'-phosphopantetheine is striking in all biological systems that have been examined, the specific role of this turnover is not understood. Since apo-ACP would be inactive in fatty acid biosynthesis, it is tempting to postulate that removal of 4'-phosphopantetheine from ACP might be a means of controlling overall fatty acid synthetase activity. However, studies attempting to correlate the relative cellular concentrations of apo-ACP and holo-ACP with in uiuo fatty acid synthesizing activity have not yet been reported.

111. Malonyl CoA-ACP Tranracylase

A. HISTORICAL BACKGROUND, DISTRIBUTION, AND METABOLIC SIGNIFICANCE Malonyl CoA-ACP transacylase catalyzes the transfer of malonyl groups from CoA to ACP forming malonyl ACP as shown in reaction (4). The first report of an isolated enzyme which catalyzes this reaction followed the fractionation of the E. coli fatty acid synthetic enzymes (8, 5 0 ) . Using standard protein purification procedures on bacterial extracts in which all the fatty acid synthetic enzymes are found nonassociated, malonyl transacylase was isolated and shown to be distinct from the enzyme that catalyzes transfer of acetyl groups from CoA to ACP. The purification of ACP and isolation of the other biosynthetic enzymes from E. coli extracts permitted the demonstration that malonyl ACP is the substrate required in the condensation reaction of fatty acid biosynthesis, catalyzed by P-ketoacyl ACP synthetase [reaction ( 5 ) ] (8, 9, 50, 61). The importance of malonyl transacylase in fatty acid synthesis was thereby defined. Although the dissociation and isolation of a separate malonyl trans50. A. W. Alberts, P. W. Majerus, B. Talamo, nnd P. R. Vagelos, BiochemistTy 3, 1563 (1964). 51. A. W. Alberts, P. W. Majerus, and P. R. Vagelos, Biochemistry 4, 2265 (1965).

5.

177

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

acylase has not been achieved in the fatty acid synthetase multienzyme complexes, the role of this enzyme in the complexes of yeast and pigeon liver has been actively investigated. The first report of an enzyme that catalyzes the transfer of malonyl groups between different thiol acceptors was made by Lynen who reported this activity in the purified yeast FAS (5’U). Incorporating the observation that malonyl pantetheine and malonyl N-acetylcysteamine could substitute for malonyl CoA in fatty acid synthesis, Lynen demonstrated that the synthetase complex catalyzed the transfer of [“Clmalonyl groups from CoA to pantetheine according to reaction (11).

+

[14C]malonylS-CoA pantetheine-SH

[1F]malonyl-S-pantetheine

+ CoA-SH

(11) In addition, he reported that the ratio between FAS activity and malonyl transacylase remained constant during the course of a 200-fold purification of the enzyme. Thus, it was evident that the transacylase activity was intimately associated with the FAS multienzyme complex. In spite of the fact that malonyl transacylase has not been isolated from the yeast FAS complex, Lynen’s laboratory has reported important experiments on the mechanism of malonyl transacylation in this system, and the conclusions drawn from these experiments have been corroborated by studies of the pigeon liver FAS and also by studies of the isolated E. coli malonyl transacylase. I n the initial experiments, yeast FAS was incubated with labeled malonyl CoA, and this resulted in the labeling of two classes of binding sites on the complex, one labile and another stable to performic acid oxidation. Identification of the first class as sulfhydryl sites was based on the fact that sulfhydryl groups are converted to sulfonic acid derivatives by performic acid oxidation with the consequent release of the acids which are present as thioesters (55’).The second class of malonyl linkages, that were stable to performic acid oxidation, were termed “nonthiol sites” (53, 5 4 ) . By labeling the FAS complex with malonyl CoA and digesting with proteolytic enzymes, Lynen and co-workers attempted to identify the chemical nature of the malonyl binding sites through a characterization of the labeled peptides (53-56). Two different groups of [14C]malonylpeptides were isolated after peptic hydrolysis. One class of peptides contained [ “ c ] 52. I. Harris, B. P. Meriwether, and J. Hastings-Park, Nature (London) 198, 154 (1963). 53. F. Lynen, BJ 102, 381 (1967). 54. F. Lynen, D. Oesterhelt, E. Schweizer, and K. Willecke, in “Cellular Compartmentalisation and Control of Fatty Acid Metabolism” (F. C. Gran, ed.), p. 1. Academic Press, New York, 1968. 55. E. Schweizer, F. Piccinini, C. Duba, S. Gunther, E. Ritter, and F. Lynen, Eur. J. Biochem. 15, 483 (1970).

178

P. ROY VAGELOS

malonyl groups that were labile to oxidation by performic acid, and these peptides contained 2-mercaptocthylamine, phosphate, and p-alanine, indicating the presence of 4’-phosphopantetheine of ACP. The other group of [l*C]malonyl peptides was resistant to performic acid oxidation. One of these peptides was a pentapeptide containing the amino acids, leucine, alanine, glycine, serine, and histidine. Because of the performic acid stability of malonya groups bound to this peptide, the lability of the malonyl groups to alkaline conditions, and the absence of cysteine in the peptides, it was proposed that serine was the most probable nonthiol malonyl binding site in the yeast FAS and that serine is in the active site of the malonyl transacylase ( 5 6 ) . The observation of nonthiol sites involved in malonyl transacylation in the yeast complex was extended by similar studies of Porter and coworkers (56-61) and of Wakil and co-workers (62-64) which produced nearly identical results in the pigeon liver FAS complex. Thus, it is apparent that E. coli, yeast, and pigeon liver fatty acid synthetases contain malonyl transacylase as an enzyme component. The enzyme appears to function exclusively in the synthesis of malonyl ACP, an intermediate in fatty acid biosynthesis. Although all fatty acid synthetase systems contain this enzyme, only the E. coli malonyl transacylase has been isolated and studied as a homogeneous protein.

PROPERTIES B. MOLECULAR Malonyl CoA-ACP transacylase of E. coli has been purified 4800-fold by procedures which included chromatography on DEAE-cellulose, Sephadex G-100, Sephadex G-75, DEAE-Sephadex, and preparative polyacrylamide gel electrophoresis ( 6 5 ) . This procedure produced pure enzyme that had a specific activity of 1.85 mmoles/min/mg protein. The 56. E. J. Jacob, P. H. W. Butterworth, and J. W. Porter, ABB 124, 392 (1968). 57. C. J. Chesterton, P. H. W. Butterworth, and J. W. Porter, ABB 126, 864 ( 1968).

58. G. T. Phillips, J. E. Nixon, A. S. Abramovitz, and J. W. Porter, ABB 13& 357 (1970). 59. J. E.Nixon, G. T. Phillips, A. S. Abramovitz, and J. W. Porter, ABB 138,372 (1970).

60. G.T. Phillips, J. E. Nixon, J. A. Dorsey, P. H. W. Butterworth, C. J. Chesterton, and J. W. Porter, ABB 138, 380 (1970). 61. J. D. Brodie, G. Wasson, and J. W. Porter, JBC 239, 1346 (1964). 62. V. C. Joshi, C . A. Plate, and S. J. Wakil, JBC 245, 2857 (1970). 63. V. C. Joshi and S. J. Wakil, ABB 143, 493 (1971). 64. C. A. Plate, V. C. Joshi, and S. J. Wakil, JBC 245, 2868 (1970). 65. F. E. Ruch, Jr., Ph.D. Theuis, Washington University, St. Louid, Missouri, 1972.

5.

ACYL GROUP TRANSFER

179

(ACYL CARRIER PROTEIN)

enzyme was shown to be homogeneous by criteria th a t included electrophoresis on polyacrylamide gels both in the presence and absence of sodium dodecyl sulfate, sedimentation equilibrium, and velocity centrifugation, and by amino acid analysis. A molecular weight of 36,600, obtained with the carboxymethylated enzyme by sedimentation equilibrium measurements, agreed with determinations on the native enzyme by sodium dodecyl sulfate polyacrylamide gel electrophoresis (36,500) and by Sephadex G-100 column chromatography (37,000). These studies indicate that malonyl transacylase is a single polypeptide chain of approximately 36,700 molecular weight, and t.hey substantiate the molecular weight estimated earlier by Joshi and Wakil ( 6 3 ) .An s~~,,.,= 2.31 was determined for the enzyme by sedimentation velocity measurements. Amino acid analysis and determination of t.he isoelectric point (pH 4.65) both showed the enzyme to be acidic in nature. The presence of 6 half-cystine residues was indicated by the performic acid oxidation method of Moore (66) and confirmed by carboxymethylation using iodo[ 1-14C]acetic acid as described by Him (67).

C. CATALYTIC PROPERTIES 1. Assays

Two assay procedures are employed for measurement of malonyl transacylase. A coupled spectrophotometric assay conveniently measures the overall activity using the following series of reactions:

+ +

+ +

(12) (13) (14) The formation of malonyl ACP by malonyl transacylase is coupled with P-ketoacyl ACP synthetase [reaction (13) ] and pig heart p-hydroxyacyl CoA dehydrogenase [reaction (14)] which convert malonyl ACP to acetoacetyl ACP and P-hydroxybutyryl ACP, sequentially, with the consequent oxidation of NADH which is measured spectrophotometrically a t 340 nm (65). Alternatively, a precipitation assay is used in which malonyl transfer from [ 14C]malonyl CoA is followed directly by measuring the formation of acid insoluble [14C]malonyl ACP (68). Enzyme is incubated with [14C]malonyl CoA and ACP. The reactions are stopped by addition of Malonyl CoA ACP Fmalonyl ACP CoA Acetyl ACP malonyl ACP 5 acetoacetyl ACP COZ Acetoacetyl ACP NADH H+ &hydroxybutyryl ACP

+ +

+ ACP + NAD+

66. S. Moore, JBC 238, 235 (1963). 67. C. H. W. Him, “Methods in Enzymology,” Vol. 11, p. 199, 1967. 68. A. W. Alberts, P. W. Majerus, and P. R. Vagelos, “Methods in Enzymology,” Vol. 14, p. 53, 1969.

180

P. ROY VAGELOS

trichloroacetic acid, and the precipitated labeled product is counted directly. 2. p H Optimum, Substrate Specificity, and Kinetics The enzyme has a broad pH optimum from 6.5 to 8.5 ( 6 5 ) . It is very specific for malonyl thioesters ; acetyl thioesters are essentially inactive. Although the physiological thiols involved in the reaction are CoA and ACP, the enzyme also catalyzes malonyl transfer to pantetheine, N - ( N ncetyl-p-alanyl) cysteamine, and N-acetylcysteamine. Although acetyl CoA is inactive as a substrate, it is a competitive inhibitor of malonyl CoA with a K i value of 115 pM ( 6 3 ) . Kinetic studies of the transacylase suggested the formation of a malonyl-enzyme intermediate during the reaction. Lineweaver-Burk plots a t different concentrations of malonyl CoA and a t various fixed concentrations of ACP resulted in a series of parallel lines. I n addition, CoA is a competitive inhibitor of ACP, and malonyl ACP is a competitive inhibitor of malonyl CoA. These findings are consistent with a pingpong Bi-Bi kinetic scheme for the malonyl transacylation ( 6 3 ) . 3. Catalytic Mechanism (Malonyl-Enzyme Intermediate)

Experiments from two laboratories (63, 65, 69) have indicated that malonyl transacylation occurs via two partial reactions:

+

+

Malonyl CoA E c malonyl-E CoA Malonyl-E ACP malonyl ACP I?

(15) (16) I n reaction (15) malonyl CoA reacts with the malonyl transacylase to form a stable malonyl-enzyme intermediate, and the latter reacts with ACP in reaction (16) to form malonyl ACP and the free enzyme. The formation of the malonyl-enzyme intermediate after incubation a t 0" of [''C]malonyl CoA with enzyme was detected by trichloroacetic acid precipitation or by Sephadex G-50 chromatography of the reaction mixture (66). Based on the acid precipitation assay, it was determined that 0.75 mole of malonyl groups was bound by 1 mole of malonyl transacylase (MW 36,700). The covalent nature of the product was indicated by the fact that it was not dissociated by trichloroacetic acid or by 8 M urea, I n order to establish that CoA was not bound to the malonylenzyme complex, an incubation of enzyme with doubly labeled [14C]malonyl [3H]CoA was carried out. Isolation of the malonyl-enzyme by column chromatography indicated that the intermediate contained I4C but no 3H. Thus, CoA is displaced from malonyl CoA during formation

+

+

69. P. R. Vagelos, Curr. Top. Cell. Regul. 4, 119 (1971).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

181

of the malonyl-enzyme linkage. Incubation of the isolated malonylenzyme with CoA led to the formation of malonyl CoA, while incubation with ACP led to the formation of malonyl ACP. Thus, the isolated malonyl-enzyme was active in reactions (15) and (16). Malonyl-enzyme isolated in the presence of 8 M urea was enzymically inactive. However, when the urea was diluted to 0.1 M, at least 60% of the malonyl groups could be transferred to CoA, suggesting that malonyl-enzyme can be renatured after it has been denatured by urea. Attempts to measure the rate of malonyl-enzyme formation at 0" indicated that the reaction was too rapid to measure since it had gone to completion within 3 sec. The equilibrium constant for the reversible formation of malonyl-enzyme from malonyl CoA and enzyme, as defined by K,, = [ malonyl-enzyme] [CoA] /[enzyme] [ malonyl CoA] was determined to be 0.025. Investigation of the effect of pH on the formation of malonyl-enzyme indicated an optimum between pH 6.0 and 7.5. Salt concentrations above 0.1 M were inhibitory to malonyl-enzyme synthesis. Study of the properties of the malonyl-enzyme complex showed that it is unstable at pH values over 6. The half-life at 0" in 0.1 M potassium phosphate, pH 6.8, is 30-40 min. On the other hand, in the presence of 8 M urea malonyl-enzyme was stable a t pH 10; pH values of 12 or higher were required for rapid hydrolysis when the protein was denatured by urea. Treatment of native malonyl-enzyme with 2 M hydroxylamine at pH 7.0 caused the release of 85% of the malonyl groups within 20 sec. In the presence of urea a t concentrations of 2 M or higher, however, malonyl-enzyme did not react with hydroxylamine. Dilution of the urea to a concentration of 0.1 M caused the malonyl-enzyme to regain its sensitivity to neutral hydroxylamine, to alkaline hydrolysis, and its ability to react with CoA or ACP. These experiments indicated that malonyl-enzyme must be in its native conformation for enzymic activity as well as for reactivity with base or neutral hydroxylamine. As mentioned above, the lability of thioesters to performic acid oxidation is well established, and Lynen and co-workers (63-56)have shown that malonyl groups are bound to the yeast FAS complex through nonthiol sites. Therefore, the malonyl-enzyme intermediate was incubated with performic acid for 16 hr a t 4". No malonyl groups were released by this treatment. This result, in addition to the finding that malonylenzyme became insensitive to base hydrolysis and to reactivity with neutral hydroxyamine when it was denatured by urea, indicates that the malonate of malonyl-enzyme is bound to the protein as an oxygen ester rather than as a thioester. In order to identify the malonyl binding site of malonyl transacylase,

182

P. ROY VAGELOS

[ " C ] malonyl-enzyme was subjected to proteolytic digestion by thermolysin. A labeled tetrapeptide was isolated and this peptide contained nlanine, glycine, histidine, and serine in amounts stoichiometric with the amount of [ " C ] malonate. Further proteolysis of the isolated thermolytic peptides by exposure to ribosomal aminopeptidase and Streptomyces griseus pronase gave rise to a radioactive product that was identified as malonyl-0-serine by co-electrophoresis with synthetic malonyl-0-serine. As further support of the identity of the proteolytic product and synthetic malonyl-0-serine, the radioactive material eluted from paper after electrophoresis was chromatographed together with synthetic malonyl-0serine on the amino acid analyzer. As shown in Fig. 6, essentially all the radioactivity was recovered in the fractions containing malonyl-0-serine. Thus, esterification of malonatc to the hydroxyl group of serine was established for the first time as thc sitc of malonyl attachment in the E . coli malonyl transacylase. Although the site of attachment of the inalonyl group to the nonthiol peptapeptide isolated from the yeast FAS was not similarly established (66),there is a striking similarity between the E . coli malonyl tetrapeptide amino acid composition (alanine, glycine, histidine, and serine) and the composition of the yeast malonyl pentapeptide (leucine, alaninc, glycine, histidine, and serine). Although it has been clearly established that a malonyl-enzyme intermediate is formed in the malonyl transacylase reaction and that the malonyl group of the intermediate is bound to a hydroxyl group of a serine residue of the enzyme (65), sulfhydryl groups also appear to be involved in the enzymic reaction. The suggestion that sulfhydryl groups 10 1 Malonvl-0-serine 9

$1

'

I

I

1

p

i6 min

4

2 c

a

f

D

0

2

500

g

400

7

300

u

200

8

100

0

E

1 0

I

20

40.

80 60 100 Elution time (min)

120

140

FIa. 6. Chromatography of malonyl-0-serine and radioactive component derived from proteolytic digest of malonyl peptide. The radioactive peptide was treated with aminopeptidase and pronase. After paper electrophoresis, the radioactive compound was mixed with synthetic malonyl-0-serine and chromatographed on the long column of the automatic amino acid analyzer. Samples of the fractions were reacted with ninhydrin or analyzed for radioactivity (66).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

183

were involved in malonyl transacylation by this enzyme was first made by Alberts e t al. (50) and by Williamson and Wakil (70) based on inhibitor studies. Joshi and Wakil later reported that the enzyme is strongly inactivated by phenylmethanesulfonylfluoride, suggesting that it possesses an active serine residue but that it is insensitive to sulfhydryl inhibitors such as N-ethylmaleimide or iodoacetamide a t concentrations of 10 mM (65).Ruch has reinvestigated the possibility that malonyl transacylase has a critical sulfhydryl group (65).He found that enzymic activity was stimulated when the enzyme was incubated with dithiothreitol, suggesting that a reduced sulfhydryl group is involved in the reaction. The enzyme was inhibited by all sulfhydryl inhibitors tested, but the extent of inhibition was pH dependent with some of the inhibitors. N-Ethylmaleimide (11.5 mM) caused 100% inhibition of the enzyme a t pH 8.6 but only 37% inhibition at p H 7.3. A similar p H effect was noted when iodoacetamide and iodoacetic acid were tested as alkylating agents. The effect of sulfhydryl inhibitors was also noted on the formation of malonyl-enzyme (Table IV) . N-Ethylmaleimide (8.3 mM) caused 98% inhibition at p H 8.6 but only 47.4% at pH 7.3. p-Chloromercuribenzoate (0.5 mM) caused 92.8% inhibition of malonyl-enzyme formation, but this inhibition was not pH dependent. Inhibition by N-ethylmaleimide, p-chloromercuribenzoate, or iodoacetamide (not shown) was prevented by prior addition of malonyl CoA to the reaction mixtures. This indicates that conversion of enzyme to malonyl-enzyme renders it insensitive to sulfhydryl inhibitors. All these findings together suggest the involvement of a sulfhydryl group(s) in the active site of this enzyme. Although protection of the enzyme by preincubation with malonyl CoA strongly suggests that the reactive sulfhydryl residue is located within the active site, the position of such a residue a t a site removed from the catalytic region but structurally proximal to it would also serve to explain these results. The relationship of the sensitive sulfhydryl group (s) to the serine residue in the active site remains to be determined. The mechanism of malonyl transfer in the E. co2i malonyl transacylase, which involves the formation of an oxygen ester with a serine residue of the enzyme and transfer of the malonyl group from the serine to a sulfhydryl of ACP, forming a thioester, malonyl ACP, appears to be unique. The formation of acyl- or phosphoryl-enzyme intermediates in reversible reactions, such as those catalyzed by thiolase (71), succinyl CoA synthetase (7%’) , and glyceraldehyde-3-phosphate dehydrogenase (52) involves the transfer of substrates to these enzymes with the forma70. I. P. Williamson and S. J. Wakil, JBC 241, 2326 (1966). 71. V. Gehring and J. I. Harris, FEBS Lett. 1, 150 (1968). 72. J. L. Robinson, R. W. Benson, and P. D. Boyer, Biochemistry 8, 2503 (1969).

184

P. ROY VAGELOS

TABLE IV EFFECTSOF SULFHYDRYL INHIBITORS ON MALONYL-ENZYME FORMATION" Inhibition at

Inhibitor N-E thylmaleimide N-E thylmaleimide N-Ethylmaleimide N-E thylmaleimide N-Ethylmaleimide N-Ethylmaleimide 30 pM malonyl CoA p-Chloromercuribenzoate p-Chloromercuribenzoate p-Chloromercuribenzoate p-Chloromercuribenzoate p-Chloromercuribenzoate p-Chloromercuribenzoate p-Chloromercuribenzoate 30 pM malonyl CoA

+

+

Concn. (mM)

pH 8.6

pH 7.3

(%I

(%I

0 0.4 0.8 8.3 25.0 25.0 0 0.05

0 60.5 73.5 98.0

0 13.1 25.0 47.4 72.6

0.08

0.17 0.50 5.0 0.50

100.0 0 0

48.0 82.0 92.6 92.8 100.0 0

0 0

n.t.b n.t. n.t. 90.7 96.3 0

Malonyl transacylase wm pre-reduced by incubation in 0.1 M tris-HC1, pH 8.6, and 0.05 M dithiothreitol a t 25" and separated from the reducing agent by chromatography on Sephadex G-50. Enzyme, mixed with the indicated inhibitors a t pH 8.6 or 7.3 in 0.1 M tris-HC1, wm incubated for 20 min at 25", and the N-ethylmaleimide reactions were stopped by addition of 5 pmoles of dithiothreitol. Where indicated, malonyl CoA was added 3 min prior to the inhibitor. Treated enzyme wm tested for malonyl-enzyme formation by incubation with [14C]malonylCoA. [W]Malonyl-enzyme, formed after a 3-min incubation at 0", wm isolated by trichloroacetic acid precipitation and counted directly. Percent of inhibition of malonyl-enzyme formation is indicated above (66). *Here, n.t. stands for not tested.

tion of acyl-cysteine or phosphoryl-histidine intermediates, which are themselves high energy compounds. A comparison of the free energy of hydrolysis a t pH 7.0 of ethyl acetate (-4720 cal/mole) and S-acetylmercaptoacetate ( - 7200 cal/mole) indicates that an oxygen acylenzyme intermediate requires the input of additional energy for the subsequent formation of an acyl thioester (73). The transfer of malonyl groups from an oxygen ester to a thioester linkage via a serine hydroxyl group in malonyl transacylase was shown to require a certain tertiary structure of the enzyme. When dissolved in 2 M or higher concentrations of urea, malonyl-enzyme was insensitive to 2 M neutral hydroxylamine, stable to alkaline pH values up to 12.0, and enaymically inactive. Since dilution of the urea concentration restored the activity of the intermediate, it appears unlikely that an acyl shift under the high urea con73. W. P. Jencks and M. Gilchrist, JACS 86, 4651 (1964)

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

185

centrations would account for the altered reactivity. The observation that malonyl-enzyme was insensitive to neutral hydroxylamine in high urea concentrations also appears to rule out an equilibrium of malonyl0-serine-enzyme with a malonyl thioester-enzyme intermediate. A similar effect of urea in distorting tertiary structure and decreasing the reactivity of an acyl-enzyme intermediate was previously shown with chymotrypsin (74). A solution of acetyl-chymotrypsin in 8 M urea was reduced in its sensitivity to base-catalyzed hydrolysis and hydroxylamine cleavage to the level of N,O-diacetylserinamide, an oxygen ester active site analog. Studies of the mechanism of chymotrypsin, which has acyl0-serine in its active site, have indicated that certain ionizable groups function in the catalysis of this enzyme. Evidence for the participation of histidine and aspartic acid residues has been observed from pH and kinetic studies ( 7 5 ) , alkylation and affinity labeling experiments ( 7 6 ) , and X-ray crystallographic data ( 7 7 ) . Thus, the involvement of an ionizable sulfhydryl group in malonyl transacylation, suggested by the pH effect noted in the sulfhydryl inhibitor studies discussed above, would not be surprising.

IV. Acetyl CoA-ACP Transacylase

A. HISTORICAL BACKGROUND, DISTRIBUTION, AND METABOLIC SIGNIFICANCE Acetyl CoA-ACP transacylase catalyzes the transfer of acetyl groups from CoA to ACP forming acetyl ACP as shown in reaction ( 3 ) . As in the case of malonyl CoA-ACP transacylase, acetyl CoA-ACP transacylase was discovered as a component of the E . coli fatty acid synthetase (8, 5 0 ) . Separation of the various enzymes of this fatty acid synthetase led to the observation that acetyl CoA was inactive in the synthesis of fatty acids until ACP and an enzyme fraction which catalyzed the synt.hesis of acetyl ACP from acetyl CoA was added to the reaction mixtures. Partial purification of the acetyl CoA-ACP transacylase allowed the demonstration that the transfer of acetyl groups from CoA to ACP was accompanied by an equivalent disappearance of sulfhydryl groups of ACP. Moreover, treatment of acetyl ACP with neutral 74. B. M. Anderson, W. V. Cordes, and W. P. Jencks, JBC 236, 455 (1961). 75. M. L. Bender, G. E. Clement, F. J. Kezdy, and H. Heck, JACS 88, 3880 (1964). 76. R. A. Osterbaan, M . VanAdrichem, and J. A. Cohn, BBA 63, 204 (1962). 77. J. J. Birktoft, B. W. Matthews, and D. M. Blow, BBRC 36, 131 (1969).

186

P. ROY VAGELOS

hydroxylamine released the acetyl groups from acetyl ACP, forming acetyl hydroxamic acid. Thus, acctyl ACP was shown to be a thioester (8).When the condensing enzyme of fatty acid biosynthesis, P-ketoacyl ACP synthetase, was isolated, it became apparent that acetyl ACP is the required substrate, along with malonyl ACP, in the initial condensation reaction of the biosynthetic sequence. The importance of acetyl CoA-ACP transacylase in fatty acid synthesis was thereby established. Investigations of acetyl &A-ACP transacylase as an isolated protein have been carried out only with the E . coli enzyme, although the presence of a similar enzyme has been implied by the results of studies of several other nonassociated fatty acid synthetases. As in the case of malonyl CoA-ACP transacylase, many attempts have been made to isolate the acetyl CoA-ACP transacylase from the multienzyme complexes of yeast and animal tissues without major succcss. Low yields of both these transacylases have been obtained from pigeon liver fatty acid synthetase after partial dissociation of the complex in the presence of 0.5 M guanidine HCl (62); however, the enzyme recovered in this manner was unstable, and it was not characterized. Lynen and co-workers (53, 54), studying the yeast FAS with [ “C] acetyl CoA, showed that [ 14C]acetyl groups were bound to the enzyme complex and that the [14C]acety1-FAS could be isolated by precipitation with trichloroacetic acid or by Sephadex filtration. Acetyl groups were shown to be bound to both sulfhydryl and nonthiol sites by studying the release of acetyl groups after performic acid oxidation, as discussed above. Proteolytic digestion of [ 14C]acetylFAS gave rise to [14C]acetyl-peptides that were analyzed. One group of labeled peptides was found to contain components of 4’-phosphopantetheine and was therefore derived from ACP. Another group contained the acetyl group on a nonthiol site. Similar studies by Porter and coworkers (56, 67) and by Wakil and co-workers (62, 64) with the pigeon liver fatty acid synthetase have corroborated the finding that acetyl groups are bound on both thiol and nonthiol sites in the multienzyme complexes. The nonthiol acetyl site is thought to be associated with the acetyl CoA-ACP transacylase. This nonthiol site of the multienzyme complexes is thought to be serine; therefore, the acetyl group would be bound as acetyl-0-scrine in the active site.

B. MOLECULAR PROPERTIES Acetyl CoA-ACP transacylase has been purified about 230-fold from E. coli extracts by procedures which included ammonium sulfate precipitation and chromatography on columns of DEAE-cellulose and Sephadex

5.

ACYL GROUP TRANSFER

(ACYL CARRIER PROTEIN)

187

G-100 (78). The partially purified preparation had a specific activity of 0.3 pmole/min/mg protein. This enzyme is relatively unstable, and it has therefore not been obtained in a highly purified state. However, the purified preparation is free of malonyl CoA-ACP transacylase, P-ketoacyl ACY synthetase, and P-ketoacyl ACP reductase. Enzymic activity is destroyed by boiling for 1 min.

PROPERTIES C. CATALYTIC 1. Assay The assay measures the production of acid-insoluble labeled acetyl ACP in which the label is derived from the transfer of radioactive acetate from acetyl CoA to ACP (78). After incubation of [ 14C]acetyl CoA with ACP and enzyme, reactions are stopped by addition of perchloric acid, and the radioactive precipitate is recovered by Millipore filtration and counted. 2. p H Optinaum and Substrate Specificity

Acetyl CoA-ACP transacylase exhibits a pH optimum a t approximately pH 6.5 a t which point there is essentially no detectable nonenzymic activity. High ionic strength is required for maximal activity. The K,, for the system [acetyl ACP] [CoA]J[acetyl CoA] [ACP] is 2.09 (70). The enzyme is relatively specific for the acetyl moiety. It has been reported (70) that longer chain fatty acyl thioesters up to C 8 can replace acetyl CoA, but with much slower rates. Thus, the enzyme can transacylate CoA esters of propionic, butyric, and hexanoic acids a t a rate of 23.4, 9.8, and 4.5% that of acetyl CoA, respectively. Malonyl CoA is inactive. Pantetheine can replace ACP, and acetyl pantetheine readily substitutes for acetyl CoA. Mercaptoethanol does not substitute for ACP. 3. Catalytic Mechanism (AcetyGEnzyme Intermediate)

Acetyl transacylase was inhibited 89% by 0.1 mM N-ethylmaleimide and 83% by 0.1 mM iodoacetamide, and this inhibition was prevented by prior incubation of the enzyme with acetyl CoA (70).This prompted the suggestion that the reaction, as in the case of malonyl CoA-ACP transacylase, occurs as two partial reactions with an acyl-enzyme intermediate as indicated in reactions (17) and (18). 78. A . W. Alberts, P. W. Majerus, and P. R. Vagelos, “Methods in Enzymology,” Vol. 14, p. 50, 1969.

188

P. ROY VAGELOS

+

+

Acetyl CoA E + acetyl-E CoA Acetyl-E ACP acetyl ACP E

+

+

(17)

(18)

I n reaction (17) acetyl CoA reacts with acetyl transacylase to form a stable acetyl-enzyme, and the latter reacts with ACP in reaction (18) to form acetyl ACP and the free enzyme. Williamson and Wakil (70) have reported that incubation of the enzyme with [14C]acetyl CoA led to the formation of [14C]acetyl-enzyme, which was separated from the reaction mixture by filtration through Sephadex G-25. The [ W ] acetylenzyme was able to transfer the [14C]acetyl group to either CoA or ACP, supporting the role of an acetyl-enzyme intermediate in reactions (17) and (18). Based upon the inhibitions noted with N-ethylmaleimide and iodoacetamide, Williamson and Wakil have proposed that the enzyme intermediate is a thioester ( 7 0 ) . However, the nature of the acetylenzyme was not characterized. I n light of the information now available concerning the E. coli malonyl CoA-ACP transacylase (Section 111,C13), and also the evidence suggesting that acetyl CoA-ACP transacylase of the yeast and pigeon liver fatty acid synthetase complexes forms an acetyl-0-serine enzyme intermediate (see above), further information must be obtained with the E. coli isolated enzyme to substantiate the suggestion that the acetyl-enzyme intermediate in this reaction is a thioester.

V. P-Ketoacyl ACP Synthetare

A. HISTORICAL BACKGROUND, DISTRIBUTION, AND METABOLIC SIGNIFICANCE P-Ketoacyl ACP synthetase catalyzes the condensation reaction of fatty acid biosynthesis. The first condensation reaction of the biosynthetic sequence involves acetyl ACP and malonyl ACP, according to reaction (19).

+ malonyl ACP $ acetoacetyl ACP + COZ + ACP

(19) This enzyme also functions in all the subsequent condensation reactions with longer chain fatty acyl ACP's t,hat are intermediates in the biosynthetic sequence (see below). The enzyme was discovered in extracts of C . lcluyveri ( 1 , 79, 8 0 ) , but the pure protein has only been obtained Acetyl ACP

79. P. R. Vagelos, JACS 81, 4119 (1959). 80. P. R. Vagelos and A. W. Alberts, JBC 235, 2'786 (1980).

5.

ACYL GROUP TRANSFER (ACTL CARRIER PROTEIN)

189

from extracts of E. coli (81, 82). Like the other enzymes of the E. coli fatty acid synthetase, studies of P-ketoacyl ACP synthetase awaited the isolation of ACP since the substrates in the synthetase reaction are thioesters of ACP. When substrate quantities of acetyl ACP and malonyl ACP became available, it became obvious that these substrates, rather than acetyl CoA and malonyl CoA, are the reactants in the condensation reaction of fatty acid synthesis. Thus, this enzyme is responsible for acyl chain elongation during de novo fatty acid synthesis from acetyl CoA and malonyl CoA. Although the enzyme has been detected in all fatty acid synthetase systems that have been examined, it has only been extensively st,udied in E. coli. Condensing enzyme activity has been studied as a component of the multienzyme fatty acid synthetase complexes of yeast (20,53, 83) and pigeon liver (84), but the enzyme has not been isolated from either of these sources. However, sulfhydryl inhibition experiments, as well as acyl binding experiments, with these multienzyme complexes suggest that the P-ketoacyl ACP synthetase component of these complexes is functionally similar to the E. coli enzyme.

B. MOLECULAR PROPERTIES f3-Ketoacyl ACP synthetase of E. coli has been purified approximately 700-fold by procedures which included ammonium sulfate precipitation, chromatography on columns of DEAE-cellulose, hydroxylapatite and Sephadex G-200,and crystallization from ammonium sulfate solutions. This proeedurc produced pure enzyme that had a specific activity of 14 pmoles/min/mg protein (81).In standard 7.5% polyacrylamide disc gels a t pH 8.5, a single band of protein was observed with an electrophoretic mobility slightly greater than that of bovine serum albumin. The molecular weight of the enzyme, determined from equilibrium centrifugation studies, is 66,000. Treatment of the enzyme with 6 M guanidine HC1 causes it to dissociate into inactive subunits. The subunit size was determined to be 37,000 by agarose gel filtration chromatography. Enzymic activity was partially recovered from guanidine HC1-denatured enzyme when the guanidine solution was diluted 10-fold. Maximal recovery of activity was observed after 3-day incubation a t 4'. The enzyme was also rapidly dissociated by 0.1% sodium dodecyl sulfate, and the subunit 81. M. D. Greenspnn, A. W. Alberts, and P. R. Vagelos, JBC 244, 6477 (1969). 82. D. J. Prescott and P. R. Vagelos, JBC 245, 5484 (1970). 83. F. Lynen, I. Hopper-Kessel, and H. Eggerer, Biochem. 2. 340, 95 (1964). 84. S. Kumar, J. A. Dorsey, R. A. Muesing, and J. W. Porter, JBC 245, 4732 (1970).

190

P. ROY VAGELOS

weight, determined by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate, was 35,000. I n an effort to determine whether the subunits of the synthetase are identical, peptide mapping of trypsinhydrolyzed alkylated protein was performed, and the results of this experiment suggested that the subunits are identical (82). The amino acid composition of the enzyme has been determined (81). Amino acid analysis after performic acid oxidation showed that the protein contains 8 moles of cysteic acid per mole of protein. Titration of the protein with 5,5’-dithiobis (2-nitrobenzoic acid) after denaturation with sodium dodecyl sulfate showed 7.2 -SH groups per mole of protein. C. CATALYTIC PROPERTIES 1. Assays

Since the enzyme catalyzes the formation of acetoacetyl ACP from acetyl ACP and malonyl ACP, it is conveniently assayed by coupling the reaction with pig heart p-hydroxyacyl CoA dehydrogenase (85).The latter was shown to catalyze the NADH-dependent reduction of acetoacetyl ACP to form L- ( ) -p-hydroxybutyryl ACP; thus, the enzymic synthesis of acetoacetyl (or other p-ketoacyl) ACP can be followed spectrophotometrically by observing the decrease in absorption a t 340 nm as a result of the oxidation of NADH. In an alternative assay, the formation of P-ketoacyl ACP is measured directly by the increase in absorbance at 303 nm a t pH 8.5 in the presence of MgC1, (85).

+

2. pH Optimum, Substrate Specificity, and Kinetics The enzyme has a rather broad pH optimum between pH 7.0 and 7.8. The rate of the reaction falls off sharply below p H 6.5 and above pH 8.0. Optimal enzymic activity requires either 0.5 M 2-mercaptoethanol or 10 mM dithiothreitol. In addition, EDTA a t 7 m M stimulates the reaction. Studies of the substrate specificity have done much to elucidate the physiological function of this enzyme as well as to help unravel the details of the mechanism of action. The predominating saturated fatty acid in E . coli is palmitate (hexadecanoate) , whereas the predominating unsaturated fatty acids are palmitoleate (cis-9-hexadecenoate) and cisvaccenate (cis-11-octadecenoate) (7, 86). The fatty acid synthetase of this organism synthesizes both saturated and unsaturated fatty acids 85. A. W. Alberta, P. W. Majerus, and P. R. Vagelos, “Methods in Enzymology,” Vol. 14, p. 57, 1969. 86. V. A . Knivett and J. Cullen, BJ 103, 299 (1967).

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

191

in vitro, and, as shown by Bloch (87), the critical reaction in the biosynthetic pathway that leads to unsaturated fatty acids is catalyzed by P-hydroxydecanoyl thioester dehydrase. As discussed earlier (Section II,C) saturated fatty acids are synthesized from acetyl ACP and malonyl ACP by the repetition, sequentially, of reactions catalyzed by P-ketoacyl ACP synthetase, P-ketoacyl ACP reductase, P-hydroxyacyl ACP dehydrase, and enoyl ACP reductase. During the course of chain elongation P-hydroxydecanoyl ACP occurs as the 10-carbon intermediate. This intermediate lies a t the branch point between the pathways to saturated and unsaturated fatty acids. As shown in Fig. 7, ( ~ , pdehydration of P-hydroxydecanoyl ACP (left pathway) gives rise to trans-2-decenoyl ACP which is a normal intermediate in saturated fatty acid synthesis. The thioester intermediates expected to undergo chain elongation in the condensation reactions of this pathway include decanoyl ACP, dodecanoyl ACP, and tetradecanoyl ACP. On the other hand, the right pahhway is initiated by the Ply dehydration of p-hydroxydecanoyl ACP, catalyzed by P-hydroxydecanoyl thioester dehydrase, to form Cis-3decenoyl ACP, the first unique intermediate in the unsaturated fatty acid pathway. This thioester presumably condenses with malonyl ACP to initiate chain elongation of the &-unsaturated acyl ACP intermediates. Other hypothetical intermediates expected to undergo condensation with malonyl ACP in the unsaturated pathway include cis-5-dodecenoyl ACP, cis-7-tetradecenoyl ACP, and cis-9-hexadecenoyl ACP (palmitoleate). Figure 7 emphasizes the fact that the enzymes of the E. coli fatty acid synthetase catalyze a parallel series of reactions in the synthesis of saturated (left pathway) and unsaturated fatty acids (right pathway). The only difference between the substrates in the two pathways is the presence of a cis double bond in the precursors of unsaturated fatty acids. The availability of pure P-ketoacyl ACP synthetase and many of the postulated ACP intermediates has permitted investigations to determine whether this enzyme catalyzes all the condensations in the biosynthesis of both saturated and unsaturated fatty acids, and whether the specificity of this enzyme can explain the accumulation of fatty acids of particular chain lengths in the cell (88).As shown in Table V, reactions with acetyl ACP, decanoyl ACP, and dodecanoyl ACP gave approximately similar results for both K , and Vn,ax.The enzyme is slightly less active with tetradecanoyl ACP. However, it is completely inactive with hexadecanoyl ACP, the C,, saturated fatty acid that accumulates in the cell. Assay of the enzyme with cis-3-decenoyl ACP and cis-5-dodecenoyl 87. K. Bloch, Accounts Chem. Res. 2, 193 (1969). 88. M. D. Greenspan, C. H. Birge, G. Powell, W. S. Hancock, and P. R. Vagelos, Science 170, 1203 (1970).

192

P. ROY VAGELOS

R = CH&H&

S-ACP

/ R A S - A C P

1 R

0 R4

S -ACP

A

s

- ACP

0-ketoacyl ACP synthetase

p-ketoacyl ACP reductase

(3)

I.-

S-ACP

TPNH

S-ACP P-hydroxyacyl ACP dehydrase

RA

enoyl ACP reductase

- ACP

FPYH)

(5)

I

t

t R4

Cle-ACP

S

s

- ACP

cis-9-C,,- ACP and cis- 11 -C,,-ACP

FIR 7. Reactions beyond the divergence of the saturated and unsaturated fatty acid biosynthetic pathways in E . coli. Pathway on left leads to saturated fatty acids; pathway on right leads to unsaturated fatty acids.

5.

193

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

TABLE V ACTIVITY OF 8-KETOACYL ACP SYNTHETASE WITH VARIOUS INTERMEDIATES IN FATTY ACIDSYNTHESIS IN E. coZ+ V,X

Intermediate Acetyl ACP Decanoyl ACP Dodecanoyl ACP Tetradecanoyl ACP Hexadecanoyl ACP cis-3-Decenoyl ACP cis-5-Dodecenoyl ACP cis-9-Hexadecenoyl ACP cis-ll-Octadecenoyl ACP

K, (PM)

(#mole product/min/mg)

0.52

2.8

0.33 0.27 0.28

2.8 0.97 0.31 N.A.b

0.71 0.20 0.37

1.9

1.7 0.37 N.A.

Assays were carried out according to Greenspan et al. (88). Kinetic constants were determined from Lineweaver-Burk plots of the data obtained. * Here, N.A. stands for no activity.

ACP, two early intermediates in the synthesis of unsaturated fatty acids, indicated that they are both as active as acetyl ACP or decanoyl ACP and have similar K , values. The activity of cis-9-hexadecenoyl ACP is decreased to approximately one-fifth of the rate of the cis-5-dodecenoyl ACP. The cis-1 l-octadecenoyl ACP is completely inactive. These results indicate that this condensing enzyme can function in both saturated and unsaturated fatty acid synthesis in E . wli.Furthermore, the specificity of this enzyme explains how chain elongation is terminated specifically at C,, in the saturated pathway and a t C,, and CIS in the unsaturated pathway. It should be mentioned, from the point of view of the other biosynthetic enzymes (Fig. 7), that specificity studies with /3-ketoacyl ACP reductase and p-hydroxyacyl ACP dehydrase have indicated that both of these enzymes are also active with intermediates of the saturated and unsaturated fatty acid biosynthetic pathways (89). This suggests that the presence of the cis double bond in the substrates of these enzymes does not intefere with their reactivities. As opposed to the broad specificity demonstrated by P-ketoacyl ACP synthetase for the fatty acyl group of the ACP thioesters, the enzyme shows absolute specificity for the ACP portion of the substrates (51,90). Thioesters of CoA and pantetheine, even a t elevated concentrations, are completely inactive in the condensation reaction. As will be described 89. C. H. Birge and P. It. Vagelos, JBC 247, 4921 (1972). 90.R.. E. Toomey and S. J. Wakil, JBC 241, 1159 (1966).

194

P. ROT VAGELOS

below, however, thc cnzymc docs catalyze some reactions involving thioesters of CoA. Kinetic studies of p-ketoacyl ACP synthetase have indicated that the K , for either acetyl ACP or malonyl ACP is not affected by the concentration of the other substrate ( 5 1 ) . This suggests that the binding of the two condensing units is nonconsecutive and a t independent binding sites, and this is surprising since it is known that a stable acyl-enzyme is formed during thc course of thc condensation reaction (see below). 3. Catalptic Mechanism

a. Condensation Reaction. The major reaction catalyzed by p-kctoacyl ACP synthetase proceeds via two partial reactions:

+

+

Acetyl-S-ACP HS-E 2 acety1-S-E ACP-SH Acetyl-S-E malonyl-S-ACP : rtcetoacetyl-S-ACP COz

(20) (21) In reaction (20) acetyl ACP (or longer chain acyl ACP) reacts with the enzyme to form a stable acetyl-enzyme intermediate, which in this case is a thioester (81).The acetyl group is transferred from the enzyme to the methylene carbon of malonyl ACP, forming acetoacetyl ACP in reaction (21). The decarboxylation of the free carboxyl group of malonyl ACP occurs during this condensation reaction. The evidence supporting this mechanism comes from studies of the sulfhydryl groups of the enzyme as well as investigations of the acetylenzyme intermediate. As mentioned above the enzymic activity is stimulated by sulfhydryl rcagcnts such as 2-mercaptoethanol and dithiothreitol. In addition, the enzymc is very sensitive to alkylating agents such as iodoacetamide and N-ethylmaleimide. Kinetic studies of enzyme inhibition by iodoacetamide indicated that the enzyme was completely inactivated when only onc molecule of iodoacetamide had reacted with one molecule of p-ketoacyl ACP synthetase (81).To test this more directly, direct binding studies were done with [ 14C]iodoacetamide in the absence and in the presence of acetyl ACP, which was found to completely protect the enzyme from inhibition by alkylating agents. The results of this experiment indicated that about 0.7 mole of [ 14C]iodoacetamide was bound per mole of enzyme when the enzyme was 93% inhibited by iodoacetamide, whereas only 0.1 mole was bound when the enzyme was protected by acetyl ACP. Thus, inhibition occurred when approximately 1 mole of iodoacetamide was taken up per mole of enzyme. I n order to determine which amino acid was alkylated, the alkylated [ "C] enzyme was submitted to acid hydrolysis. Essentially all of the radioactivity of thc protein was associated with carboxymethylcysteine which was identified by thin layer chromatography, high voltage electrophoresis on

+

+

+ HS-E

5.

ACYL GROUP TRANSFER (ACYL CARRIER PROTEIN)

195

paper, and chromatography on the automatic amino acid analyzer. These experiments indicated that the enzyme has a cysteine residue that is essential in the reaction (81). Protection of the critical cysteine residue of the enzyme by acetyl ACP suggested that an acetyl-enzyme might form, thereby preventing the sulfhydryl group from reacting with the alkylating agent. I n order to test this, the enzyme was incubated with doubly labeled [ 3 H ] a ~ e t y l [14C]ACP, and the labeled acetyl-cnzyme intermediate was isolated by chromatography on Bio-Gel P-60 followed by sucrose density centrifugation. The acetyl-enzyme isolated by this procedure contained approximately 0.5 mole of [3HH]acetylper mole of enzyme but only about 0.05 mole of 14C. Thus, i t appears that CoA is not involved in the acetylenzyme intermediate. The [3H]acetyl group was released from the enzyme when the protein was treated with alkali, neutral hydroxylamine, or performic acid. These properties of the intermediate suggest that the acetyl group is bound covalently to a cysteine residue of the enzyme in thioester linkage (81). The isolated [ 3H]acetyl-enzyme was shown to be active in transferring the [3H]acetyl group either to ACP to form [ 3H]acetyl ACP [reaction (20) 3 or to malonyl ACP to form [ 3HJ acetoacetyl ACY [reaction (21) 1. Thus, it is suggested that acetyl-enzyme, in the case of the /!-ketoacyl ACP synthetase, is a thioester enzyme intermediate in the condensation reaction of fatty acid synthesis.

b. Fatty AcyZ CoA-ACP Transacylation. In addition to the acyl transfer reaction [reaction ( 2 0 1 , which is part of the normal overall condensation reaction catalyzed by /!-ketoacyl ACP synthetase, this enzyme also catalyzes fatty acyl CoA-ACP transacylation as show? in reaction (22) (41). RCO-S-COA + ACP-SH RCO-S-ACP + C O A S H (22) This reaction, the transfer of acyl groups from thioesters of CoA to ACP, was demonstrated in experiments where the enzyme was incubated with ['%] tetradecanoyl CoA and [3H]ACP. Filtration of the reaction mixtures through Sephadex G-50 (Fig. 8 ) led to the separation of a coincident peak of 3H and l*C radioactivity (fractions 2 4 3 4 ) from the bulk of the ["C] tetradecanoyl CoA radioactivity, indicating that [ 14C]tetradecanoyl [ 3H]ACP had been formed during the incubation. When either enzyme or ACP was omitted during the incubation, no peak of I4C appeared in this region. The small radioactivity peak in the region of the void volume of the column (fractions 18-22) was a contaminant of the [ 14C]tetradecanoyl CoA. /!-Ketoacyl ACP synthetase catalyzes fatty acyl transaeylation of tetradecanoyl CoA a t 0.5% of the rate of /!-ketoacyl ACP synthesis from tetradecanoyl ACP and malonyl

196

P. ROT YAGELOS

3000

2000

5

0

I000

20

40 Fraction number

60

FIG.8. Demonstration of the formation of tetradecanoyl ACP by P-ketoacyl ACP synthetase. A-Ketoacyl ACP synthetase (30 pg) was incubated with 5 nmoles of I"Cltetradecanoy1 CoA (10 pCi/pmole), 20 nmoles of I'HIACP, 2 pmoles of 2-mercaptoethanol, and 5 pmoles of imidasole-HCl buffer, pH 6.7, in 0.1 ml. Aftrr 30 min a t 37" the reaction was terminated by boiling. The reaction mixture was subjected to filtration chromatography on a Sephadex G-50 column equilibrated with 0.05 M imidazole-HC1, pH 6.7. Fractions were counted for radioactivity. ( 0 )"C "C radioacradioactivity with A-ketoacyl ACP synthetase or ACP omitted, (0) tivity in complete system, and ( A ) ['HIACP in presence or absence of P-ketoncyl ACP synthetase (41).

ACP. Thioester specificity studies of reaction (22) have indicated that acetyl CoA was inactive; however, CoA thioesters of 6, 8, and 10 carbon atoms were equally active, while tetradecanoyl CoA and cis-9-hexadecenoyl CoA were somewhat less active. Neither hexadecanoyl CoA nor cis-ll-octadecenoyl CoA was an active substrate, The acyl group specificity of p-ketoacyl ACP synthetase for various acyl ACP substrates (Table V) is very similar to the specificity for acyl CoA substrates with the exception that acetyl ACP is activc in the p-ketoacyl ACP synthctase reaction, whereas acetyl CoA is inactive in the fatty acyl CoA-ACP transacylation. It is apparent that fatty acyl transacylation occurs in two reactions catalyzed by p-ketoacyl ACP synthetase: the transfer from acyl ACP to a sulfhydryl group of the enzyme in reaction (20) and the transfer froin acyl CoA to ACP in reaction (22). The possibility was considered that acyl enzyme occurs as an intermediate in the transfer of acyl groups from CoA to ACP. If this were the case, it would be expected that acyl

5.

ACYL GROUP TRANSFER (ACTL C.4RRIER PROTEIN)

197

CoA thioesters would be substrates in P-ketoacyl ACP synthesis ; however, as noted above, CoA thioesters are completely inactive in this reaction. A specific cysteine residue of the enzyme is involved in reaction (20). This cysteine is preferentially alkylated a t low concentrations of iodoacetamide, and alkylation is prevented when the enzyme is initially reacted with fatty acyl ACP since the enzyme cysteine is converted to a thioester. Since fatty acyl CoA’s are not active substrates for P-ketoacyl ACP synthesis, it was anticipated that an enzyme thioester would not be formed during the fatty acyl CoA-ACP transacylation. In order to test this supposition, the ability of fatty acyl CoA’s to protect the enzyme against alkylation by iodoacetamide was tested. The results of the experiment showed that the thioesters active in fatty acyl CoA-ACP transacylation protected the enzyme against alkylation (41). These results suggest that the acyl CoA site of the enzyme is the same or close to the acyl ACP site. Although the catalysis of fatty acyl CoA-ACP transacylation by P-ketoacyl ACP synthetase has been demonstrated, the mechanism of this transacylation is not understood. However, a possible explanation for this reaction involves the specificity of this enzyme for the fatty acyl groups. As noted above, any even-numbered, straight chain saturated fatty acyl ACP from Cz to C,, as well as the cis-monounsaturated fatty acyl ACP thioesters from C,, through C,, are effective substrates in P-ketoacyl ACP synthesis. This broad range specificity for a fatty acyl chain suggests that there is a hydrophobic region on the enzyme that has affinity for the acyl group, in addition to a site for ACP. The usual thiol moiety of the thioester substrates is the protein, ACP. The acyl groups of thioesters of ACP are covalently bound to the protein through its prosthetic group, 4’-phosphopantetheine, and 4‘-phosphopantetheine also constitutes the component of CoA to which fatty acyl groups are esterified. TI~uY, fatty acyl thioesters of CoA are nucleotide analogs of thioesters of ACP, and therefore it is not surprising that P-ketoacyl ACP synthetase has a high affinity for thioesters of both ACP and CoA. Since the enzyme has a specific site for both the fatty acyl group and the thiol group, it is possible that at the C2level, affinity of the enzyme is high for acetyl ACP, which contains the physiological thiol, but that it is very low for acetyl CoA because the 2-carbon acyl group is too small for adequate hydrophobic interaction with the acyl site of the enzyme. With longer chain acyl groups, there is greater opportunity for hydrophobic interactions; therefore, the enzyme has a high affinity for longer chain fatty acyl CoA’s as well as ACP’s. Evidence that the enzyme has a high affinity for certain fatty acyl CoA thioesters is indicated by the finding that these thioesters are substrates in the fatty acyl CoA-ACP transacylase reaction

198

P. ROT VAGELOS

and that thcy can protect the critical cysteine of the cnzymc from alkylation by iodoacetamide. I n the first partial reaction catalyzed by P-ketoacyl ACP synthetase, reaction (20), the acyl group of a fatty acyl ACP is transferred to a sulfhydryl group of the enzyme, and therefore transacylation is a major function in the physiological reaction catalyzcd by this enzyme. I n spite of the fact that fatty acyl CoA’s protect the critical cystcine of the cnzymc; thcse compounds are not active substrates in P-kctoacyl ACP synthesis. Thus, it is possible that CoA thioestcrs occupy thc acyl ACP site imperfectly and that the acyl group of the CoA thioester may not be transferred to the cysteine of the enzyme. Thc suggestion that acyl transfer does not occur from CoA to the enzyme is based on lack of activity of CoA thioesters in the overall p-ketoacyl ACP synthetase reaction. If the cysteine of the enzyme is not esterified, protection by fatty acyl CoA’s of this cysteine against alkylation might occur through steric hindrance or conformational changes imposed on the enzyme when the fatty acyl thioester site is occupied, even imperfectly. An alternate explanation for the protection against alkylation is that the fatty acyl group is transferred from CoA to the cysteine of the enzyme, but that it cannot condense with malonyl ACP because that reaction requires a conformational change in the enzyme which occurs only when the acyl ACP site is occupied by ACP. I n this latter hypothesis it would be assumed that acyl CoA’s cannot induce the proper conformational changes. c. Malonyl ACP Decarboxylation. I n addition to the decarboxylation of malonyl ACP that is catalyzed by P-ketoacyl ACP synthetase as part of the overall reaction [reaction (21) ] , this enzyme also catalyzes malonyl ACP decarboxylation that is independrnt of fatty acyl ACP, as shown in rcaction (23) (41). Malonyl ACP acetyl ACP + CO? (23) P-Ketoacyl ACP synthetase catalyzes malonyl ACP decarboxylation a t 2 3 % of the rate of P-ketoacyl ACP synthesis. Although all the other reactions catalyzed by P-ketoacyl ACP synthetase are sensitive to sulfhydryl inhibitors (see above), this decarboxylase activity is not inhibited by iodoacetamide at concentrations that complctcly inhibit both overall P-ketoacyl ACP synthesis and fatty acyl CoA-ACP transacylation. The relationship of this decarboxylase activity to the physiological reaction is not clear. Two mechanisms havc been proposed for the condensation-decarboxylation reaction catalyzcd by the P-ketoacyl ACP synthetase (91). One involvcs a concerted condensation-decarboxylation; the --f

91. F. Lynen, in “Organizational Biosyntliesis” (H. J. Vogel, J. 0. Lampen, and V. Bryson, eds.), 11. 243. Academic Press, New York, 1967.

5.

ACPL GROUP TRANSFER (AWL

CARRIER PROTEIN)

199

second involves a two-step reaction characterized by thc initial formation of an enzymically stabilized carbanion by decarboxylation of malonyl ACP,followed by condensation of the carbanion with acetyl-enzyme. I n the former case, the decarboxylation which occurs independent of acyl ACP would represent an abortive reaction of the enzyme. I n the latter case, the decarboxylation would represent thc actual formation of the carbanion which in turn would react with an H+ to givc free acetyl ACP. ACKNOWLEDGMENTS The unpublished experimental work from this laboratory presented here and the preparation of this article have been assisted by grants from the National Institutes of Health (R01-HL-10406)and the National Science Foundation (GB-5142X).

This Page Intentionally Left Blank

Chemical Basis of Biological Phosphoryl Transfer S. J. BENKOVIC

0

K. J. SCHRAY

I. Introduction . . . . . . . . . , 11. Hydrolysis of Acyclic Phosphate Esters . . . . A. The Metaphosphate Mechanism for Monoesters B. Acyclic Phosphate Di- and Triesters . . . 111. Nucleophilic Reactions a t Acyclic Phosphorus . . IV. Pentacovalency and Pseudorotation . . . . . V. Catalysis of Phosphoryl Transfer or Ligand Loss . . A. Intramolecular Catalysis . . . . B. Metal Ion Catalysis . . VI. Enzymic Catalytic Mechanisms . . A. The Metaphosphate Mechanism . . B. Bimolecular or Associative Mechanisms

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201 202 202 207 208 214 219 219 227 232 233 235

1. Introduction

The aim of this chapter is to present an abbreviated but comprehensive discussion of the probable mechanisms involved in biological phosphoryl transfer reactions as disclosed by studies of model systems and their extrapolation to the enzyme mediated processes. A number of critical, extensive reviews on particular aspects of this topic have recently been written to which the reader who desires further detail and possibly clarification is referred (1-5). Recent examples will be emphasized. 1. F. H. Westheimer, Accounts Chem. Res. 1, 70 (1968). 2. A. J. Kirby and S. G. Warren, “The Organic Chemistry of Phosphorus.” Elsevier, Amsterdam, 1967. 201

202

S. J. BENKOVIC AND K. J . SCHRAY

II. Hydrolysis of Acyclic Phosphate Esters Although it is somewhat artificial to dissect hydrolysis from other nucleophilic reactions, the intensity of investigation on phosphohydrolase enzymes suggests such division may be useful. MECHANISM A. THEMETAPHOSPHATE

FOR

MONOESTERS

Various lines of evidence have led to a general acceptance of the dissociative monomeric metaphosphate mechanism for the hydrolysis of monoester monoanions derived from alcohols and phenols, thiols, and amines (6-9).The principal supporting data include: (a) a general observation of P-0, P-S, or P-N bond cleavage; (b) entropies of activation close to zero in contrast to bimolecular or associative solvolyses where entropies of activation are usually more negative by 20 eu (10);(c) molar product compositions (methyl phosphate :inorganic phosphate) in mixed methanol-water solvent which approximate the molar ratio of methanol: water or favor methyl phosphate formation inferring the presence of a highly reactive electrophilic species; and (d) the existence of linear free-energy relationships between the logarithmic rates of hydrolysis of the monoanions and the dissociation constants of the corresponding leaving group (Fig. 1 ) . The small slopes observed in the structure-reactivity correlations, - 0.3 and 0.08 for 0- and S-phosphate monoesters, respectively, are in accord with the departure of a neutral ligand suggesting that the mechanism generally involves proton transfer to the potential leaving group in a preequilihrium step ( 1 ) . Proton transOII

RX-p-o--I

OH

0 Kzw _ + I : RX-P-o-H 1:0

kr

RXH + (PO,]

X = 0,S, NH 3. J. R. Cox and B. Ramsay, Chem. Rev. 64, 317 (1964). 4. T. C. Bruice and S. J. Benkovic, “Bioorganic Mechanisms,” Vol. 11. Benjamin, New York, 1966. 5. S. J. Benkovic, in “Comprehensive Chemical Kinetics” (C. H. Bamford and C. F. Tipper, eds.), p. 1. Amer. Elsevier, New York, 1972. 6. C. A. Bunton, E. J. Fendler, E. Humeres, and K. V. Yang, J . Org. Chem. 32, 2806 (1967). 7 . A. J. Kirby and A . G. Varvoglis, JACS 89, 414 (1967). 8. S. Milstien and T. H. Fife, JACS 89, 5820 (1967). 9. S. J. Benkovic and E. J. Sampson, JACS 93, 4009 (1971). 10. F. A. Long, J. G. Pritchard, and F. E. Stafford, JACS 7Q,2362 (1957).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

-I

203

-

-2-

--

-3-

'c .-

4-

-31 - 5 E

n 0

X

0

0 -

-6-7-

-8-

2

-lo

4

6

10

8

12

14

16

PKa

FIG.1. Plots of log kobrd ( m i d ) for the hydrolysis of S-, N-, and O-phosphate ester monoanions a t 20" vs. the pK, of the leaving group (9).

fer apparently proceeds directly or through intervening solvent molecules and may become rate determining when the leaving group corresponds to a pK, <7 for 0 monoesters. For S monoesters this step may be partially rate determining throughout the pK, range depicted in Fig. 1 and, if so, might then account for the subtle change in the sensitivity toward pK, ( - to slope) for these two classes of esters. Some support for this suggestion is found in the deuterium solvent isotope effects (lcH,o/kD,o)of 1 .6 1 .8 that have been detected in the pK, regions of interest for both the 0 and S esters. The rate step itself, therefore, must involve diffusion apart of the two fragments and solvation of the metaphosphate species. The biphasic structure-reactivity correlation which describes phosphoramidate hydrolysis does not represent a gross change in mechanism. Unlike the 0- and S-phosphate monoanions, the zwitterions derived from phosphoramidates are readily isolable (11, 12) ; consequently, their aqueous solutions are anticipated to be mixtures of zwitterionic and non-

+

11. D. E. C. Corbridge and E. J. Lowe, JCS, London p. 493 (1954). 12. E. Jampel, M. Wakselmann, and M. Vilkas, Tetrahedron Lett. 31, 3533

(1968).

204

S. J .

BENKOVIC AND K. J. SCHRAY

ewitterionic species. The structure-reactivity correlation for these esters is the sum of the Br@nstedslope for k, (-1.0) and the dependency of K,, on the pK, of the amine ( 1.0). Since K,, is approximately unity for amine leaving groups of PI<,7.2 phosphoramidates with pK, <7.2 exist primarily in the nonewitterionic form, whereas those with pK, >7.2 are predominately ewitterionic in aqueous solution. As a result ( 9 ) the observed rate constant remains relatively constant for the former-a cancellation of effects-but depends directly on the pK, of the leaving amine for the latter. It is of interest to find that N-phosphoryl imidazolium monoanion is about 50-fold less reactive than predicted, perhaps as a result of increased delocalization in the ion ( 2 ) . The equality be-

+

tween its rate of hydrolysis and that for N-phosphoryl-”-methylimidaeolium ion confirms that proton transfer is not rate determining in this case and that hydrolysis proceeds solely through the ewitterion species. I n a similar context the rate of hydrolysis of N-phosphoryl creatine monoanion is predicted by the above relationship and therefore does not manifest unusually high hydrolytic lability as previously postulated (13). The metaphosphate mechanism also applies to the hydrolysis of the appropriate ionic species of polyphosphates. Kinetic studies of y-phenylpropyl di- and triphosphate revealed pH-rate profiles (pH 4-9) that are similar to those observed with adenosine di- and triphosphate, respectively (14). Thus, the hydrolytic mechanisms for these substrates are presumably the same with the adenosine residue having no influence. Since hydrolysis of monoprotonated 7-phenylpropyl diphosphate is some two thousand times faster than hydrolysis of the symmetrically substituted P,P‘-di-y-phenylpropyl pyrophosphate, a monomeric metaphosphate rather than a displacement mechanism appears operative for the monoester dianion ( 3 ) . Verification of this hypothesis is derived from the close agreement between the observed rate coefficient for the hydrolysis of the unsymmetrical pyrophosphate diester, P,,P,-diethylpyrophosphate, a model for (I) and the calculated rate coefficient for hydrolysis of (I) (16). Yet, it is interesting that solvent competition studies with mixtures of meth13. P. Haake and G . W. Allen, Proc. Nnt. Acad. Sci. U. S. 68, 2691 (1971). 14. D. L. Miller and F. H. Westheimer, JACS 88, 1507 (1966). 15. D. L. Miller and T. Ukena, JACS 91, 3050 (1969).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

0 II

0 11

C,H, (CH,) ,O -P -o -P-0-

b-

I

-

L

0 II

205

0 II

C,H, (CH,),O -P -0- P-0I

HO

OH

0 II

c,H,(cH,),o-P-oI OH

b-

t

+

IPO;]

anol and water show that no methyl phosphate is formed when pyrophosphate monoanion is solvolyzed. Although this does not rule against the metaphosphate mechanism, it is necessary to assume that the product-forming transition state is selectively solvated by water (16). Note that in (I) the proton is not bound internally to the bridging oxygen but a t the thermodynamically preferred site. I n contrast, the hydrolysis of P1,P,-diethylpyrophosphate monoanion, which most likely proceeds through the zwitterionic species is 300-fold slower in rate than the dianion. Presumably similar mechanisms apply to triphosphate ester hydrolyses, i.e., the di- and trianions. The hydrolytic lability of (I) is characteristic of dianions of phosphate monoesters possessing excellent leaving groups, e.g., carboxylate, phosphate monoanion, or dinitrophenolate. For 2,4- and 2,6-dinitrophenyl phosphate, the dianion, as in the aforementioned case, is more reactive than the monoanionic species (7, 1 7 ) . This spontaneous hydrolysis is attributed to a mechanism involving metaphosphate expulsion from the dianion and is supported by the usual experimental criteria-insignificant deuterium solvent isotope effects, entropies of activation near zero, and trapping experiments conducted in various alcohol-water mixtures which yield significant amounts of the corresponding phosphate esters. The latter product distributions, ester:inorganic phosphate, parallel the solvolysis under similar conditions of sulfur trioxide-isoelectronic with metaphosphate-enabling one to rely with some confidence on such trapping experiments as a sufficient but not necessary criterion for the metaphosphate mechanism (18). Dianion hydrolysis, as implied above, is highly sensitive to the pK, of the leaving group as measured by a slope of - 1.2 for the structurereactivity correlation (see Fig. 2). It is of interest that the rate CO-

16. C. A . Bunton and H. Chaimovich, Inorg. Chem. 4, 1763 (1965). 17. C. A. Bunton, E. J. Fendler, and J. H. Fendler, JACS 89, 1221 (1967). 18. S. J. Benkovic and P. A. Benkovic, JACS 90, 2646 (1968).

206

S. J . BENKOVIC AND K . J . SCHRAP

efficient for hydrolysis of (I) and the unsymmetrical pyrophosphate diester analog mentioned above do not correlate well with this relationship, being some 200-fold slower than predicted (19). This deviation along with the above collective data is consistent with a transition state that is well advanced, i.e., a high degree of P-0 bond cleavage; thus, resonance stabilization of the leaving group is important ( 4 ) .

Additional evidence, to be discussed subsequently, discloses that the sensitivity of dianion hydrolysis to the pK, of the departing anion is experimentally identical to that for the equilibrium or complete transfer of the phosphoryl nioiety in accord with (4). It is important to realize that, although metaphosphate is invoked as a probable product in (1) , ( 3 ) , and (4), there is no evidence for its existence as a fully independent, unsolvated ion. Perhaps an accurate assessment of the above hydrolyses is that they represent the limiting dissociative situations most likely to proceed via a metaphosphate species. An inquiry of interest is the rate of decomposition of the reactive zwitterionic species in several representative cases. A sample calculation, based on estimates of the zwitterion concentration, predicts the first-order, rate coefficient, lc,, to be lo7 min-' (39") for phenyl phosphate monoanion (7). For N-phosphoryl N'-methylimidazolium ion and phosphoramidate lc, is 10-3-10-4min-l (39") (9), and for phenylthiophosphate + monoanion [assuming a pK, -10 for RSHP02,- (ZO)], the value (35") is greater than diffusion-controlled (8). This fact supports the previous contention that the proton transfer may be rate determining in S-phosphate monoanion hydrolysis. The magnitude of lc, is sensitive to leaving group pK,; thus, the above should be viewed as median values. I n the absence of other effects these coefficients furnish a preliminary estimate as to the efficiency of enzymic catalysis provided the latter merely optimized the zwitterion concentration without changing the intrinsic pK, of the departing group. The overall order of hydrolytic reactivity (S 2 N > 0) phosphate esters, which extends a t least for the phosphoramidates and O-phosphate monoesters to other nucleophiles, offers a tentative rationale for the intermediacy of S and N esters as kinetically competent intermediates in enzyme mediated processes. 19. S. J. Benkovic and R. C. Hevey, JACS 92, 4971 (1970). 20. E. M. Amett, Progr. Phys. Org. Chem. 1, 223 (1963).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

207

B. ACYCLIC PHOSPHATE DI- AND TRIESTERS The spontaneous hydrolysis of a series of diaryl phosphate diester monoanions exhibits a high sensitivity to the nature of the leaving group ( - 1.2, 39"), but a decreased entropy of activation, -25 eu, and a small though significant deuterium solvent isotope effect, kH20/kU~o = 1.6 (21). Such evidence is in accord with a simple bimolecular nucleophilic attack by water on phosphorus perhaps subject to general base catalysis ( 5 ) .There

is no evidence for elimination of a metaphosphate ester ( 2 2 ) .On the basis of similar evidence, the identical mechanism may be written for the hydrolysis of a series of dialkyl aryl triesters; however, the aspects of general base catalysis are more prominent, ie., ICH2,,/kDZozz 2 (23). By comparing the various linear free-energy relationships for hydrolysis, it is seen that the order of reactivity is diaryl diester anions < dialkyl aryl triesters < monoester monoanions (Fig. 2 ) . Monoester dianions are more reactive than triesters and monoester monoanions when the leaving group of the former has a pK, <10 and <5, respectively. Comparable reactivity for all ester types except the monoester dianion is encountered with a leaving group of pK, z 2. It is obvious that efficient catalysis is associated with lowering the pKa of the departing group, especially if the reaction involves the dianionic species. The possibility that metal ions and suitably juxtapositioned acid groups may function in this manner has been borne out by model studies (Section V ) . A final point of emphasis is that the two hydrolytic mechanisms postulated to involve metaphosphate-like species proceed via the more reactive pathways. Thus, in terms of enzyme mediated processes, in particular direct rather than two-step hydrolysis reactions where the low phosphorylation selectivity of metaphosphate is relatively unimportant, these mechanisms are attractive from a kinetic standpoint but remain unproved possibilities. 21. A. J. Kirby and M. Younas, JCS, B p. 510 (1970). 22. C. A . Bunton and S. J. Farber, J. Org. Chem. 34, 767 (1969). 23. S. A . Khan and A. J. Kirby, JCS, B p. 1172 (1970).

208

S. J . BENKOVIC AND

K.

J . SCHRAY

PK

FIQ. 2. Linear free-energy relationships between rate constants for hydrolysis and pK,, values of the conjugate acids of the leaving groups, ROH, for the ionic spec& of phosphate mono-, di-, and triesters present at physiological pH. Broken lines represent extrapolations. A, ROPOi-; B, ROP(OH)02-; C. (RO),PO ; and D, (RO)?PO?- (23).

111. Nucleophilic Reactions at Acyclic Phosphorus

Studies of the reactions of nucleophiles, including water at the phosphorus atom of monoester dianions, are characterized by their marked lack of sensitivity to the basicity of the nucleophile (24-,$?6).The logarithms of the second-order rate coefficients for the attack of various amines on a series of 2-nitro-4-substituted-phenyl phosphate dianions have been correlated as a function of both amine and leaving group pK,. The slopes, ,8, of these linear plots are tabulated in Table I. Since there is no doubt that a bimolecular displacement reaction on phosphorus is involved-isolation of the anticipated phosphoramidate product, entropy of activation -20 eu, etc.-it is remarkable that the rates of the sub24. G. DiSahato nnd W. P. Jencks, JACS 83, 4393 and 4400 (1961). 25. A. J. Kirby and W. P. Jencks, JACS 87, 3209 (1965). 26. A. J. Kirby and A. G . Varvoglis, JCS, B p. 135 (1968).

6.

209

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

TABLE I A COMPARISON OF p VALUESFOR THE INCOMING AND LEAVING GROUPS FOR PHOSPHATE ESTERS

Type of reaction Phosphoramidaka plus amines Phosphate monoester dianions plus amines Phosphate diester monoanions plus amines Phosphate triesters plus oxyanions

p nucleophile

Ref.

p leaving group

Ref.

0.2 0-0.1

(28, 29) (26)

- 0 . 9 to - 1 . 1 -1.2

(29)

0. 3-0. 4

(32)

- 0 . 9 to - 1 . 0

(32)

0. 3-0. 6

(33)

- 0 . 3 to - 0 . 6

(33)

(26)

stitution reactions remain constant and completely independent of the basicity of the pyridines employed with the dianion of 2,4-dinitrophenyl phosphate, i.e., /I= 0 (26). The reaction obviously is a limiting case and may be described in terms of a specific molecular interaction more akin to solvation of the developing electrophilic center, i.e., pyridine replacing water, rather than the mechanism of displacement encountered with spa hybridized carbon. As the pK, of the leaving group is increased, p for the series of nucleophiles increases only slightly, 0-0.1. The rate coefficients for attack, however, have decreased significantly with the increasing basicity of the leaving group owing to the high sensitivity ( p = - 1 ) of the reaction to this variable. The p parameters, may be utilized as approximate indices of the fraction of charge transferred to the nucleophiIe or leaving group in the transition state and correspondingly as a measure of the degree of bond formation or breakage (27). Since one is correlating the kinetics of a phosphoryl transfer reaction with the equilibrium for proton transfer process, i.e., pK,,, it is necessary prior to analysis to recalibrate in terms of equilibrium phosphoryl transfer. The p values for complete transfer of [PO:-] between the donor phosphoramidate and a series of pyridines is 1.2 (28-30). Likewise, the equilibrium transfer of an 0-phosphate diester moiety between phenolic oxygens is characterized by /3 of 1.2 (31). This value, which should be independent of the chemical nature of the donor, will be employed in the ensuing discussion and should be a reasonable approximation for all classes of esters. The moieties [Poi-] or [ (RO),,PO,-], approximate the electropositive character of a proton, for which- p by definition is unity, rather than an acyl group where 27. 28. 29. 30. 31.

W. P. Jencks and M. Gilchrist, JACS 90, 2622 (1968). W . P. Jencks and M. Gilchrist, JACS 86, 1410 (1964). W. P. Jencks and M . Gilchrist, JACS 87, 3199 (1965). G. W. Jameson and J. M. Lawlor, JCS, B p. 53 (1970). R . H. Bromilow. S. A. Khan, and A. J. Kirby, JCS, B p. 1091 (1971)

210

S. J. BENKOVIC AND K. J . SCHRAT

p

= 1.6-1.7 (27). For the general case the unit charge transferred to or lost by the nucleophile may be estimated by ,Gnucleophile/l.2 with a group/l.2on the departing ligand. concomitant change in charge of pleavillg The description of the above reaction that emerges features a loose, uncoupled transition state; i.e., the degree of bond cleavage is not directly proportional to bond formation ( 6 ) . A similar situation obtains for nucleophilic attack on phosphoramidate monoanions (9).

6+

6-

X = NH;, 0-

As the degree of esterification or protonation is progressively increased, the p parameters gradually increase. For example, the logarithms of the rate coefficients for the reaction of the same series of pyridines with the monoanion of 2P-dinitrophenyl phosphate or 2,4-dinitrophenyl methyl phosphate are a linear function of nucleophile pK, with a p of 0.3-0.4 (32).As for the above case of the dianionic species, the rate of nucleophilic attack by amines on phosphorus is still extremely dependent on the pK, of the leaving group and rapidly becomes less significant as the pK, increases. Consequently, the transition state for these reactions still closely resembles ( 6 ) , although it is beginning to assume some characteristics of a displacement or associative process. I n the reactions of nucleophiles toward a series of 2- (substitutedphenyl) -1,3,2-dioxaphosphorinane-2-oxides(11), one now encounters ap-

(n) proximately equal values of p both for nucleophile and leaving group (33). For the series of pyridines, nucleophilic attac,k on the 2,4-dinitrophenyl ester of (11) defines a p of 0.6. General base catalysis now becomes important with pyridines of low pK, as anticipated from the chemistry of carboxylic acid esters, Other nucleophiles, including, for example, phosphate dianion, acetate, and trifluoroethoxide, exhibit their own linear free-energy relationships. The p for the leaving group changes 32. A. J. Kirby and M. Younas, JCS, B p. 1165 (1970). 33. S. A. Khan and A. J. Kirby, JCS, B p. 1172 (1970).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

211

from -0.65 to -0.88 for phosphate dianion and acetate, respectively, whereas pnucleoplli l e for the oxyanions increases from 0.30 to 0.48 with increasing pK, of the ester leaving group. Thus, the transition state (7) resembles that encountered in SN2 displacement and certain acyl transfer reactions (27) where bond formation and cleavage are coupled. 0.4/1.2 6-

0 0.8/1.2 6-

.. . .. .

I* RO p X RO’ ‘OR

(7)

The unimolecular dissociative aspect, as expected, has disappeared since the driving force for metaphosphate expulsion derived from excess electron density on the anionic oxygens has been abrogated by esterification or protonation. Although the possibility of a pentacovalent intermediate has not been considered within this analysis, it is obvious that regardless of the location of the rate-determining step leading to and from such an intermediate, /3 for the leaving group should not exceed p for the nucleophile. Consequently, displacement reactions on monoester mono- and dianions and diester monoanions are not anticipated to proceed via such species (Table I ) . It should also be noted that progressive protonation or esterification may lead in some cases to mechanisms involving C-0 bond cleavage, but these do not constitute phosphoryl transfer and will not be discussed here. It is useful to compare the reaction rate coefficients for several standard nucleophiles with the various classes of esters in order to visualize the effects of changes in pK,, state of ionization, and chemical identity of the nucleophilic heteroatom. The pertinent data are presented in Table 11. For the oxyanions, acetate and hydroxide, the ratio of rate coefficients for reaction with the negatively charged diester and neutral triester favors the latter by an average factor of about two thousand. For attack by amines, pyridine, and n-butylamine (not shown), the same ratio varies over a range of only two to one hundred. The latter is probably the result of the increased electropositive character of phosphorus upon esterification ( 3 4 ) .Assuming that a similar effect may be ascribed to the oxyanions, a retarding factor of ca. one hundred then may be attributed to unfavorable electrostatic repulsion per unit negative charge. The importance of this term will be considered further in Section V. A second point of interest is the higher reactivity of acetate or fluoride relative to pyridine a t the triester level despite their lower pK,. This is generally observed as a consequence of the greater strength of P-0 or P-F VS. P-N bonds being reflected in this transition state owing to the increased 34. R.

I,. Collin, JACS 88, 3281 (1966).

212

S. J. BENKOVIC AND K. J . SCHRAY

TABLE I1 RATECONSTANTS

REACTIONS OF VARIOUS 2,4-DINITROPHENYI, PHOSPHATE ESTERS"~~

FOR THE

ArOP0:-

Nucleophile

;

ArO -0-

I

Ad)-

OCH3

x x x

10-7 10-6 10-2

8.3.5 x 10-6 1.48 x 64

3.90 X

ti. 1.5

x

10-3

1.24 X

6.48 x -

5.29 x 1.92 x 10-3

-

NH2OH F5

-

3.22 7.4 2.8

1 . 9 x 10-4

H2O CHICOO-

10.7 9.36

From Kirby and Younas (32). min-l I.( = 1.0, 39".

M-1

importance of bond formation. Note that hydroxylamine only shows unusual reactivity (a effect) a t this level. A final important feature is that reactivity does not decrease monotonically from triester to monoester, an additional manifestation of the highly uncoupled, metaphosphate character of the transition state for the latter. Given the fact that common substrates for enzyme-catalyzed phosphoryl transfer reactions are either mono- or diesters, it is logical to conclude from the above that the transition state for a large body of these reactions must have metaphosphate character. Replacement of a phosphoryl oxygen atom with sulfur generates a class of phosphate esters possessing a P=S center. Such reagents have found increasing use as a means for probing enzyme-catalyzed phosphoryl transfer processes. The susceptibility of triesters containing the P=S center to nucleophilic displacement reactions is considerably lessened, i.e., k s / k o 0.03, owing to decreased polarization of phosphorus upon substitution of the less electronegative sulfur (34, 3 5 ) . For monoester hydrolysis where the metaphosphate mechanism presumably is operative, the inverse order is observed (36). The deduction of an enzyme mechanism based on determination of the same ratio in the enzyme mediated process, however, is subject to several pitfalls, not the least of which is the considerable difference in interatomic distances between

=

35. J. Ketelaar, H.Geismann, and K. Koopmans, Rec. Trav. Chim. Pays-Bas 71, 1253 (1952). 36. R.Breslow and I. Katz, JACS 90, 7376 (1968).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

213

P=S (1.94A) and P=O (1.57A) and van der Waals radii (37, 3 8 ) . Consequently, phosphorothioates may act either as good substrates, pancreatic ribonuclease (39) (Ics/ko = 0.02) ; poor substrates, alkaline phosphatase (ks/ko 0.001) (40) ; competitive inhibitors, snake venom and spleen phosphodiesterase ( 3 9 ) ; or not complex as for cyclic 3’,5’-nucleotide phosphorodiesterase (41, 42) . Their extreme usefulness, however, in stereochemical studies will be discussed in Section VI. The above displacement mechanisms have been illustrated to conform with the viewpoint that nucleophilic substitution a t a tetracovalent phosphorus atom, in the absence of geometric constraints, involves preferential apical attack and departure for ligands which are electronegative (434 5 ) . In brief this is an “in-line” mechanism. The transition state (8) may be envisioned as a trigonal bipyramid whose two apical positions are occupied by the entering nucleophile and departing group with the remaining ligands bound a t equatorial or basal positions. I n accordance

with this model are numerous observations of inversion of configuration at phosphorus for nucleophilic displacement reactions by alkoxide or hydroxide ion on optically active phosphonium and thiolphosphinate esters (46, 4 7 ) . The plausibility that under as-yet unspecified conditions an apical-equatorial displacement mechanism may operate, however, has been raised by the discordant finding of racemization in the basic hydrolysis of a thiolphosphonothionate (48). Such a result is consistent, 37. W. Saenger and F. Eckstein, JACS 92, 4712 (1970). 38. D. E. C. Corbridge, in “Phosphorus Chemistry” (D. E. C. Corbridge et al., rds.), Vol. 111, pp. 211, 293. Wiley (Interscience), New York, 1966.

F. Eckstein, JACS 92, 4718 (1970). P. Mushak and J. E. Coleman, Biochemistry 11, 201 (1972). F. Eckstein and H. P. Bar, BBA 191, 316 (1969). A similar criticism may be leveled a t rate ratios determined with O-phosphate and phosphoramidate pairs [A. Williams and R. A. Naylor, JCS, B p. 1967 ( 1971) 1. 43. S. I. Miller, Advan. Phys. Org. Chem. 6, 253 (1968). 44. P. C. Van der Voorn and R. S Drago, JACS 88, 3255 (1966). 45. W. C. Hamilton, S. J. LaPlaca, F. Ramirez, and C. P. Smith, JACS 89, 39. 40. 41. 42.

2268 (1967). 46. K. E. De Bruin, K. Naumann, G . Zon, and K . Mislow, JACS 91, 7031 (1969). 47. W. B. Farnham, K. Mislow, N. Mandel, and J. Donohue, Chem. Commun. p. 120 (1972). 48. L. P. Reiff, 1,. J. Szafraniec, and H. S. Aaron, Chem. Commun. p. 366 (1971).

214

S. J . BENKOVIC AND K. J . SCHRAY

in fact commonplace, with the intervention of metastable pentacovalent species capable of pseudorotation, a significant aspect deferred until now.

IV. Pentacovalency and Pseudorotation

Intramolecular ligand reorganization is a particular feature of the pentacoordinate family of phosphorus compounds, the phosphoranes. Such compounds may undergo a nondissociative intramolecular exchange of sites provided that the lifetime of the phosphorane is sufficient relative to decomposition and the energy barrier to ligand exchange is surmountable. Phosphoranes exhibit trigonal-bipyramidal geometry (49, 5 0 ) . The structure of a typical oxyphosphorsne, the phenanthrenequinone-triisopropyl phosphite adduct, has the following pertinent features : (a) the phosphorus atom lies within a triangle defined by three bonding “equatorial” ligands which form the basal plane of the trigonal bipyramid and their bonds subtend an angle of ca. 120”, (b) the remaining two “apical” ligands are situated above and below the basal plane and their bonds subtend an angle of ca. 180”, and (c) an 0-P-0 ring bond angle of ca. 90” with the five-membered ring spanning one apical and equatorial position. Feature (c) applies to adducts which possess a fivemembered ring and is probably a general phenomenon ( 6 1 ) . Note the resemblance of this structure to the transition states proposed above. Within (111) are, however, extensive steric interactions which occur owing to several short nonbonded distances; i.e., the apical oxygens are within 2.7 A of the first carbon of the equatorial isopropyl groups. Nonetheless, cyclic oxyphosphoranes are generally more stable than acyclic ones owing to the constraints imposed by the five-membered ring which minimize additional steric repulsions.

49. E. L. Muetterties and R. A. Schunn, Quart. Rev.,Chem. SOC.20, 245 (1966). 50. R. Schmutaler, Angew. Chem., Znt. Ed. Engl. 4, 496 (1965). 51. F. Ramirea, Accounts Chem. Res. 1, 168 (1968).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

215

Phosphoranes may undergo intramolecular ligand exchange by a process termed "pseudorotation," or more precisely the Berry mechanism (52). With the latter mechanism, pairwise exchange of apical and equatorial ligands takes place in a concerted fashion by way of a tetragonal-pyramidal transition state ( 5 3 ) . The pivotal point, an equatorial ligand-arbitrarily 3-remains equatorial in the process and occupies the apex position in the transition state (9). Proceeding from left to right the two apical groups designated 1 and 2 may be viewed as undergoing a motion which results in closing the original apical-apical bond angle from 180" to 120", whereas the remaining equatorial groups 4 and 5 appear to open the initial 120" bond angle to 180". The result is a new

trigonal bipyramid in which the pivotal group has remained in an equatorial position, but the other four groups have exchanged their positions. The Berry mechanism originally was proposed to account for the observed resonances in the NMR spectra of various phosphoranes. It has proved, after refinements imposed by the preference rules, remarkably adapted to rationalize a diverse body of experimental observations. Although an alternate means for ligand exchange, the turnstile mechanism, has recently been proposed, it offers at present no additional or more penetrating insights into the processes which will be discussed here ( 5 4 ) . It should now be readily perceived that pseudorotation may profoundly affect the stereochemical outcome of reactions involving pentacoordinate phosphorus. Without presenting the experimental evidence dictating their formulation, the preference rules for pseudorotation may be listed as follows. First, the more electronegative atoms preferentially occupy the apical positions-the Muetterties or stereoelectronic rule (49).Theoretical arguments indicate that apical bonds are generally longer, weaker, and more ionic than equatorial bonds consistent with the transition state idea depicted in (8) (4,5 6 ) . The phosphorus orbitals leading to equatorial bonds, on the other hand, are presumed to possess greater s character which would favor bonding t o electron-donating atoms ( 4 4 ) . Second, 52. R. S. Berry, J . Chem. Phys. 32, 933 (1960). 53. G. M. Whitesides and H. L. Mitchell, JACS 91, 5384 (1969). 54. P. Gillespie, P. Hoffman, H. Klisacek, D. Marguarding, S. Pfohl, F. Ramirea,

S. A. Tsolis, and I. Ugi, Angew. Chem., Znt. Ed. Engl. 10, 687 (1971). 55. H. Bent, Chem. R e v . 61, 275 (1961).

216

S. J. BENKOVIC AND K. J. S C H R A T

five-membered rings generally will span equatorial and apical positions which was illustrated above. Such rings will not usually bridge equatorialequatorial positions since the distortion of the phosphorus-ring ligand bond angle from 90" to 120" will be accompanied by a significant increase in ring strain. Apical-apical bridging is geometrically impossible. Pseudorotations which fall within this framework will require the minimal activation energy to occur. These concepts were first applied to the acid- or base-catalyzed hydrolysis of five-membered cyclic phosphates (1, 56). Several examples will serve to illustrate their employment. The hydrolysis of the methyl ester of propylphostonic acid (IV) occurs almost exclusively with ring

o \, O"P'oCH,

(Iv)

opening (57). Assuming that the addition of water is a t an apical position the intermediate derived from (IV) may be depicted (V) where the methylene group of the ring is equatorial and the oxygen atom is apical (the more electronegative ligand). The fact that loss of the methoxyl

(V)

group is negligible may be rationalized in terms of the preference rules which restrict pseudorotation of (V) since (a) pseudorotation about the equatorial carbon atom would expand the ring angle to 120", generating strain, and (b) pseudorotation about either of the other equatorial substituents as pivot would force the methylene into an apical position. Consequently, ring opening alone would be found after subsequent proton transfer provided that decomposition of (V) proceeds through apical expulsion. If both entering and leaving groups were situated equatorial, then methanol loss would be expected which is contrary to experiment. Thus, to the list of pseudorotation preference rules, a stereochemical 56. N. K. Hamer, JCS, C p. 404 (1966). 57. E. A. Dennis and F. H. Westheimer, JACS 88, 3431 (1966).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

217

rule governing the mode of reaction may be added which states that electronegative entering and leaving groups depart from apical positions. This proclivity has been verified by molecular orbital calculations. A second example is found in the base-catalyzed opening of (VI) which proceeds with almost exclusive P-0 fission whereas analogous acyclic esters undergo P-S cleavage (68). Addition of hydroxide to (VI)

is anticipated to generate (VII) with the oxyanions and sulfur located

I

OH

equatorially because of their relatively greater electron density. All pseudorotations available to (VII) would violate one or more of the above preference rules so that decomposition proceeds via P-0 bond cleavage. This example also stresses the need for monoanionic or neutral (nonzwitterionic) species for allowed pseudorotations. A final example is the acid-catalyzed hydrolysis of ethylene hydrogen phosphate which, unlike the acyclic esters treated above, is accompanied by almost equally rapid oxygen exchange into the unreacted substrate (59).A possible intermediate presumably would have structure (VIII) which upon protonation could decompose directly to the observed ring opened products. Oxygen exchange requires pseudorotation followed by protonation and loss of water. I t stands to reason that, if mechanism (10) applies, a t sufficiently high acidity the pathway leading to ring opened products should completely predominate. This in fact has been found for methyl ethylene phosphate where the fraction of exocyclic cleavage (loss of methoxy) falls to zero (60). Thus, under these conditions of high acidity, pseudorotation becomes rate limiting. 58. D. C. Gay and N.K. Hanier, JCS,B p. 1123 (1970). 59. P. C. Haake and F. H. Westheirner, JACS 83, 1102 (1961). 60. R. Kluger, F. Covitz, E. Drnnis, L. D. Williams. and F. H. Westheimer, JACS 91, 6066 (1969).

218

S. J. BENKOVIC AND K. J. SCHRAY

_t

Ring opened products

*OH

:I OH I

I

0

9

1 9

0

The above examples suffice to demonstrate the effects of pseudorotation of pentacoordinate intermediates on the stereochemical course of the reaction. Retention of configuration at phosphorus may be conceived as proceeding through a pathway involving apical attack, pseudorotation of the incipient leaving group from an equatorial position, and apical departure. Topological representations dealing with more complex situations have been dealt with elsewhere (61). It should be emphasized that the preference rules are not inviolate, but merely describe the lowest free energy pathway for the process. Numerous examples are known where these barriers are penetrated (61). However, these are energy requiring with the barrier imposed by stereoelectronic strain alone being variously estimated a t 6-12 ltcal mole-' (61, 62). The concept of pseudorotation per se does not explain the rapid rates of base- and acid-catalyzed hydrolysis of the five-membered cyclic phosphates which proceed some lo6 times faster than their acyclic analogs or higher ring homologs (63). A considerable portion of this increased reactivity resides in the strain of the five-membered ring based on evidence provided by measurements of the heats of saponification of the 61. K. Mislow, Accounts Chem. Res. 3, 321 (1970). 62. D. B. Boyd, JACS 91, 1200 (1969). 63. H. G. Khorana, G. M. Tena, R. S. Wright, and J. G. Moffatt, JACS 79, 430 (1957).

6. CHEMICAL

BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

219

acyclic and five-membered cyclic compounds (64, 66) and supported by calculation of the strain energy (66). The thermochemical strain energy is 5-6 kcal mole-l. Relief of strain in the phosphate ring on passing from the ground state with a ring angle a t phosphorus of 99" to LI trigonal-pyramidal transition state (ring angle, 90') would account for about lo' of the increased reactivity. It should be noted that the observed acceleration may be attributed to a-bond strain imposed by the ring or differences in x energy between cyclic and acyclic esters owing to distortion of the x bonding in the former (67).Regardless of the actual cause, it is obvious that a factor of lo2 remains unaccountable. This figure, then, may represent a measure of the lower free energy of a hydrolysis pathway proceeding through a metastable pentacovalent intermediate with a lifetime of >1O-l3 sec, relative to one in which only the transition state has pentacoordinate character. The translation of ring strain and pentacovalency by pseudorotation to the exocyclic ligand then accounts for the commensurate lability of P-0 bonds both within and without the five-membered ring structure. Although it is readily perceived that the probable involvement of pentacoordinate species will occur in enzyme-catalyzed processes involving cyclic esters, e.g., ribonuclease (Section VI), the question of the operation of this mechanism in phosphoryl transfer where mono- and diesters function as substrates will now be considered.

V. Catalysis of Phosphoryl Transfer or Ligand Loss

A. INTRAMOLECULAR CATALYSIS Catalysis by neighboring groups in phosphate ester hydrolysis falls into two general categories : (a) reactions which involve nucleophilic attack on phosphorus leading to the formation of cyclic five- or sixmembered rings and are general acid-base catalyzed, and (b) reactions which involve general acid catalysis of phosphoryl group expulsion. These types are comprehensively exemplified in systems where the carboxyl moiety serves as the neighboring group; consequently, the majority of illustrations will be selected from there. A particularly well known example in category (b) is the hydrolysis 64. J. R. Cox, R. E. Wall, and F. H. Westheimer, Chem. Znd. (London) p. 929 (1959). 65. E. T. Kaiser, M. Panar, and F. H. Westheimer, JACS 85, 602 (1963). 66. D.A. Usher, E. A. Dennis, and F. H. Westheimer, JACS 87, 2320 (1965). 67. M. G. Newton, J. R. Cox, and J. A. Bertrand, JACS 88, 1503 (1966).

220

S.

J. BENKOVIC AND K. J. SCHRAY

of salicyl phosphate dianion (68). Nucleophilic catalysis by the carboxylate is ruled out by the absence of an acyl phosphate intermediate and the requirement of an endocyclic displacement a t the phosphorus center of an anion (see later discussion) ; general base catalysis by the carboxylate is also unlikely owing to the lack of a significant deuterium solvent isotope effect and, more convincingly, the absence of a similar pathway for the related diester, methyl 2-carboxyphenyl phosphate. Thus, the key to mechanism lies in the positioning of the remaining proton. Its location may be probed by evaluating the effects of 4 and 5 substituents on the rate of hydrolysis of salicyl phosphate dianion in order to establish two linear-free energy correlations-one for the dependency of the rate coefficient on the pK, of the carboxyl moiety and the second on the leaving group pK, (69).The results suggest that P-0 cleavage is well advanced (the /3 leaving group is closer to that for dianion rather than monoanion hydrolysis) and that proton transfer (Br@nsteda = 0 ) is far from complete (11).This conclusion is somewhat surprising, in view of mechanism (1), which features preequilibrium proton transfer ; however, it appears that the carboxylate group initially may lie outside the plane of the benzene ring to minimize nonbonded

repulsions with the adjacent phosphate group. Its rotation upon protonation into the plane, therefore, constitutes a part of the rate-determining step. The efficiency of the catalysis markedly depends on the reference standard selected. If viewed as a dianion species then the rate of hydrolysis is some lo1"greater than predicted, as a monoanion ca. lo2 faster than anticipated. The effectiveness of general acid catalysis-expressed as the ratio of the rates of hydrolysis of the o and p isomers-may be roughly estimated from the A ~ K or , difference in the corresponding pK, values for the o and p isomers (70).The argument, in essence, is that these reactions follow Hammond behavior; thus, the increased stability of the product owing to hydrogen bonding is partially reflected in an early, lower freeenergy transition state. Whereas k , / k , for salicyl phosphate is ca. two 68. M. L. Bender and J. M. Lawlor, JACS 85, 3010 (1963). 69. R. H.Bromilow and A. J. Kirby, JCS, B p. 149 (1972). 70. S. J. Benkovic and L. K. Dunikoski, Jr., Biochemistry 9, 1390 (1970).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

221

, ; lc,/lc, for salicyl thiophosphate (IX) is only five hundred, A ~ K (3.0) where ApK, (1.9) ( 7 1 ) . The absence of significant catalysis in the hydrolysis of phosphoenolpyruvate (X) and 2-carboxyphenyl phosphoramidate ( X I ) may be accounted for in a similar manner (72, 73). I

l

0

/ ‘*‘H H,C=C I \ o C’ II 0

I1

0

II

0

Upon esterification or protonation the more favorable hydrolytic mechanism changes from catalysis of metaphosphate expulsion to one incorporating cyclization as the prevalent feature. The hydrolysis of monobenzyl phosphoenolpyruvate proceeds via loss of benzyl alcohol (90%)despite the fact it is the poorer leaving group-and minor amounts (10%) of monobenzylphosphate, respectively ( 7 4 ) . The pH-rate profile implicates the carboxyl group as being catalytically active. In the presence of hydroxylamine the course of the reaction is unchanged although pyruvate oxime hydroxamate is produced. The proposed scheme (12) incorporates pentacovalent species (XII) in order to rationalize the for-

/ -

R = -CH,C,H,

71. T. H. Fife and S. Milstein, J . Org. Chem. 34, 4007 (1969). 72. S.J. Benkovic and K. J. Schray, Biochemistry 7, 4090 (1968). 73. S. J . Benkovic and P. A. Benkovic, JACS 89, 4714 (1967). 74. K. J. Schray and S. J. Benkovic, JAGS 93, 2522 (1971).

222

S . J. BENKOVIC AND K. J. SCHRAY

mation of the acyclic acyl phosphate and to avoid an energetically less favorable apical-equatorial displacement. Either acyl phosphate species (XIII) or (XIV) would, in the presence of hydroxylamine, ultimately lead to pyruvate oxime hydroxamate. The fact that the product distribution remains unchanged under these conditions implies that benzyI alcohol loss from (XII) is nearly ten times greater than formation of acyclic acyl phosphate. Preferential exocyclic group loss is a characteristic of carboxyl group catalysis of diester hydrolysis although a persuasive rationale is lacking. It follows from the above, that the parent compound, phosphoenolpyruvate, also should undergo ring closure under the appropriate conditions. Experiments conducted in IsO-enriched water demonstrate that incorporation of label into unhydrolyzed phosphoenolpyruvate parallels hydrolysis presumably via mechanism (13). The reader’s attention is directed to the fact that hydrolysis of the cyclic

acyl phosphate (XIII) occurs with mainly P-0 rather than C-0 bond fission, a manifestation of the increased sensitivity of phosphorus when incorporated in a five-membered ring to displacement processes. Carboxylate catalysis is encountered in the hydrolysis of diesters derived from salicyl phosphate (76). These reactions proceed through exclusive exocyclic rather than endocyclic displacement although the former ligand in terms of pK, may constitute the poorer of the two leaving groups. The stereospecificity of the reaction apparently stems from a pentacovalent intermediate which cannot pseudorotate freely owing to the presence of two oxyanions in the equatorial position. This mechanism gains support from the actual observation of the cyclic acyl phos-

R = CJi,

, 3-NO&& , CH,

75. S. A. Khan, A. J. Kirby, M. Wakselman, D. P. Homing, and J. M. Lawlor,

JCS,B

p. 1182 (1970).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

223

phate and the high sensitivity of rate to substitution in the leaving group (/I = -1.2) (14). Given the limits of product detection, the barrier to pseudorotation must be a minimal 5 kcal mole-1 in reasonable agreement with calculated estimates stated earlier. Complete esterification of the phosphoryl moiety, as one might anticipate, generates mechanisms involving nucleophilic catalysis by either carboxyl or carboxylate (74, 76, 7 7 ) . The hydrolysis of dibenzylphosphoenolpyruvate proceeds via stepwise loss of benzyl alcohol (90%) and the concomitant formation of minor amounts (10%) of dibenayl phosphate ( 7 4 ) . The pH-rate profiles for release of benzyl alcohol reveal that the hydrolytically reactive species must involve a protonated carboxyl group or its kinetic equivalent. In the presence of hydroxylamine the course of the reaction is diverted to essentially quantitative formation of dibenzylphosphate and pyruvic acid oxime hydroxamate although the rate is effectively unchanged. Mechanism (15) is in accord with these observations. Hydroxylamine is postulated to trap the acyclic acyl

phosphate (XV) whose formation is rapid and reversible relative to benzyl alcohol expulsion. The fact that in aqueous solution only a small fraction of the total product is dibenzylphosphate would be a consequence of the ratio of hydroxylaminolysis to hydrolysis for acyl esters with excellent leaving groups. The reversibility of acyclic acyl phosphate formation in contrast to the diester may result from greater ease of hy76. R. H. Rromilow, S. A. Khan, and A. J. Kirby, JCS, B p. 1091 (1971). 77. R. H. Bromilow, S. A. Khan, and A. J. Kirby, JCS, Perkin Trans. 11, p. 911 (1972).

224

S. J . BENKOVIC AND K. J. SCHRAY

droxyl group addition to a tri- rather than diester. As above a pseudorotation is necessary in order to avoid an equatorial-apical displacement. Triesters derived from salicyl phosphate undergo rapid hydrolysis owing to catalysis by the carboxyl and/or carboxylate group (76, 77). The composition of the products, endo- or exocyclic displacement, depends exclusively on the basicity of the leaving group in accord with the fact that the necessary pseudorotations are allowed and moreover the freeenergy barrier separating them is small relative to decomposition. The hydrolysis of (XVI) yields mainly the product of endocyclic displacement

(96%) whereas the hydrolysis of (XVII) leads to that from exocyclic dis-

placement (98%). The ratio of endo-exocyclic products is unity for an exocyclic leaving group pK, z 8.5, which is within estimates of the basicity of the salicylate oxygen. Linear free-energy relationships involving the rate coefficients for exocyclic and endocyclic displacement as a function of the pK, of the exocyclic leaving group exhibit a high sensitivity, p = -1.4 +- 0.2 and -0.3, respectively. The former implicates a transition state for exocyclic displacement with considerable P-0 bond fission and conforms to the supposed ionic character of the apical bond. The response of the endocyclic rate to changes in exocyclic substitution on phosphorus is consistent with the previous hypothesis that the transition state in triester nucleophilic displacement reactions is coupled and furthermore implies that a considerable percentage of leaving group sensitivity resides in bond formation. One can estimate from these data that the half-life of the suspected pentacovalent intermediate is on the order of sec. The above reactions, in addition to their unique stereochemical as-

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

225

pects, manifest rate coefficients in excess of their bimolecular counterparts, indeed if one can be found. Pertinent cases are collected in Table 111 (32,74, 75, 77, 78). Several features are worthy of attention as follows : 1. Catalysis by carboxyl rather than carboxylate is generally more efficient. 2. The rate coefficients for hydrolysis reactions proceeding via transition states incorporating five- or six-membered rings are-for the present substrates-roughly comparable. 3. Electrostatic repulsion in the intramolecular systems, uncorrected for inequality in electronic effects, is only about loz; thus, after correction for unequal electronic effects a t phosphorus, the effect is negligible. 4. The ratio of kintra/kLnter, an index of effective molarity, is unity for triesters and forty molar for the diester monoanions. At concentrations of acetate equivalent to that for the o-carboxy esters, the rate of the intramolecular reactions are 104-10s more rapid than their bimolecular counterparts. Several of the above conclusions are particularly important for enzymic mechanisms. Observation 3 implies that once the substrate-enzyme complex has been formed, the retarding effect on rates of nonbonded repulsion terms has been largely overcome. The efficient operation of both modes of catalysis 1 and 4 argues that catalysis of phosphoryl transfer need not be restricted to a narrow pH range. Although there are a number of difficulties associated with the dissection of kinetic rate coefficients in intramolecular systems, proximity may be generally assigned ti value of 102-103 in “uncomplicated” five- and six-membered ring systems ( 4 ) . The achievement of pentacovalency may account, therefore, for about 102-103 rate advantage, in satisfactory agreement with our earlier estimate. Although it may appear surprising that ring size is not manifest in these reactions, this finding is in accord with the relief of ring strain accompanying the attainment of pentacovalency a t phosphorus as noted previously. The displacements, either endocyclic or exocyclic, are in essence possible mechanisms for enzymic phosphoryl transfer assuming that the enzyme may provide a means for stabilizing the intermediate pentacovalent species that is furnished by cyclization in the model systems. In a general sense, the exocyclic process is simply an apical-apical displacement or an “in-line mechanism” and may function to phosphprylate a group on the enzyme or a bound acceptor molecule (18). The‘ endocyclic process, which requires a pseudorotation for phosphoryl transfer, 78. J. Steffens, E. Sampson, I. Siewers, and S. J. Benkovic, JACS 95, 936 (1973).

TABLE I11 RATECOEFFICIENTS FOR REPRESENTATIVE SYSTEMSINVOLVING INTRAMOLECULAR CATALYSIS BY CARBOXYL OR CARBOXYLATE Rate coefficients

Substratea

Ref.

7.0

x

in-*

77

2.5

x 10-2

77

1.9

x

10-2

1.8

x

10-3

74

3.4

x

10-4

76

($,,OCHzCeHs

/

CH*=C

\\

0-

\

OCHzCeHs

COOH itOCHzCeHi

0CHFC

/

\

0-

\COOH

0-

0' '\

CIIa-CH

coo-\O/

\

1 . 0 x 10-4

0-

\

a

1 designates point of bond c-leavage.

b

Conditions, p = 1.0, 35-39',

rniii-'

except where noted.

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

227

is an apical-pseudorotation-apicaldisplacement or “adjacent mecha-

D = donor ; A = acceptor

(18)

(19)

nism” and is restricted to neutral or monoanionic pentacovalent species so that the pseudorotation be allowed by the preference rules (19). It is difficult to see an advantage for the latter process, particularly in view of the enzymic structural reorganization demanded. However, there are cases where an intramolecular phosphoryl migration within the substrate molecule may be advantageous, i.e., for activation, and presuming enzyme catalysis would be an example of (19). Depending on the initial dissociation state of the acceptor group, the donor will depart as DOor DOH in either mechanism. It is plausible that presented with a situation where proton transfer from A to D is thermodynamically unfavorable, that this step may become rate determining (74). I n conclusion, it should be obvious that several of the model systems discussed above are directly applicable to enzyme-catalyzed phosphate diester hydrolysis. Although pseudorotation does not play a part in the mechanism of action of ribonuclease (Section VI), there may be other situations where the operation of (18) or (19) for diesters will have a stereochemical advantage (endo- vs. exocyclic departure), although admittedly the initial positioning of substrate on the enzyme could have the same consequences. A relevant case might be the mechanism of action of the pyrophosphokinases where bond formation occurs between the p-phosphoryl group of the triphosphate and the acceptor (78a). Although this discussion has concentrated on carboxyl group catalysis, similar mechanisms apply to other functional groups, particularly hydroxyl (79), but the evidence here is less complete.

B. METALION CATALYSIS The important search for viable models. for the requirement of divalent metal ions in enzymic phosphoryl transfer reactions may also benefit 78a. A highly speculative mechanism of oxidative phosphorylation invokes pseudorotation in the synthesis of ATP from ADP Pi CE. F. Korman and J. McLiclr, Proc. N u t . Acad. Sci. U.8.67, 1130 (197011. 79. D. A. Usher, D. I. Richardson, Jr., and D.G . Oakenfull, JACS 92, 4699 (1970).

+

228

S. J. BENKOVIC AND K. J . SCHRAY

from the application of the previously discussed concepts. The metal ion may have a number of imagined roles: (a) a template for orienting substrates and enzymic catalytic groups, (b) charge neutralization, (c) chelation promoting metaphosphate expulsion, (d) complexation with a pentacovalent intermediate to control the stereochemical course or enhance the rate of the reaction, and (e) a hydroxyl carrier to promote hydrolysis. Several representative model systems will illustrate these points. The transfer of a phosphoryl moiety from phosphorylimidazolium ion to pyridine-2-carbaldoxime anion proceeds through a ternary complex with Znz+serving as the bridging ligand as demanded by the saturation kinetics and the lack of phosphoryl transfer in the absence of the metal ion. A mechanism in accord with these results is presented in (20). The

function of Zn2+is obviously a template one with accompanying charge neutralization in order to ensure the formation of the ternary complex. Its contribution to catalysis of phosphoryl transfer within the complex may be relatively unimportant, particularly if the transition state has considerable metaphosphate character, typical of phosphoramidates (80). Specific chelation to the leaving group in order to facilitate metaphosphate expulsion from a monoester dianion is exemplified by several cases. All possess the common feature of a second ligand in close proximity to the ester oxygen in order to induce productive complexation (81-83). Rate enhancements of 10-lOs for (21) and (22) are directly proportional to the resultant decrease in leaving group pK, owing to the metal ion-oxyanion complex and fit predictions based on the structure reactivity correlation for the hydrolysis of phosphate monoester dianions. I n the case of highly favorable chelation, a displacement process may 80. G . J. Lloyd and B. S. Cooperman, JACS 93, 4883 (1971). 81. S. J. Benkovic and L. Dunikoeki, Jr., JACS 93, 1526 (1971). 82. B. Cooperman, JACS (1973) (in press). 83. Y. Murakami and J. Samamoto, Bull. Chem. SOC.Jap. 44, 1827 (19711, and references therein.

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

229

subtend the metaphosphate mechanism (23) (84). Attempts, however, to

demonstrate a similar catalysis of metaphosphate elimination ,from the terminus of a polyphosphate chain have not been successful [Eq. (24)] (86). Since the incipient oxyanion does not lie within the coordination

sphere of the metal ion, it follows that complexation must be very strong, i.e., large stability constant, and may not be attainable in purely aqueous solution. Although metal ion catalysis of pyrophosphate formation from 7-phenylpropyl triphosphate and inorganic phosphate has been observed, the reaction unfortunately was not sufficiently amenable to characterization to establish its mechanism (86). Model reactions demonstrating the last postulates are less readily available. A possible instance of metal ion interaction with pentacovalent phosphorus occurs in the intramolecular exocylic displacement reaction of phenyl N - (glycy1)-phosphoramidate (25). The observed rate of the reaction is increased 50- to 300-fold in the presence of Mg2+and Zn2+, respectively, and constitutes one of the few examples of significant catalysis by Mg2+ (87). Catalysis of a similar magnitude is encountered with the corresponding lactate and salicylate esters. The phenomenon, there84. S. J. Benkovic and E. M. Miller, Bioinorg. Chem. 1, 107 (1972). 85. B. Cooperman, Biochemistry 8, 5005 (1969). 86. D. I,. Miller and F. H. Westheimer, JACS 88, 1514 (1966). 87. E. J. Sampson, J. Fedor, P. A. Benkovic, and S. J. Benkovic, J . Org. Chem. (1973) (in press).

230

S. J. BENKOVIC AND K. J. SCHRAY

fore, is independent of the chemical nature of the ligand, 0- or N-, and of ring size. A plausible rationale invokes metal ion interaction with the pentacovalent intermediate whose stability relative to an acyclic species is enhanced by inclusion of two of its ligands within a ring. The result of chelation may be a decrease in the pK, of the leaving group combined with stabilization of the intermediate, hypothetically depicted in (XIX) with both effects being catalytic. The catalysis is not attributed to

0

(m) charge neutralization in light of the above discussion concerning intramolecular reactions. An additional illustration involves the increased rate of methyl phosphate hydrolysis obtained by binding the ester to triethylenetetramine Cos+ ion. The acceleration in rate is about 100-fold relative to monoanionic methyl phosphate (88). No catalysis is observed with CoS+ complexes where bidentate coordination is precluded. The crucial question is whether the chemical identity of the reactive species is (XX) or (XXI). If (XX) , then catalysis may be interpreted in terms of hydroxyl

(xx)

(=I)

attack on a monoester monoanion and would be a combined example of (a) and (e). Its magnitude is anticipated from the data displayed 88. F. J. Farrell, W. A . Kjellstrom, and T. G. Spiro, Science 164, 320 (1969).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

231

in Table 11. If (XXI), then the rate of hydrolysis is ca. 1O1O faster than trimethyl phosphate. This, however, seems unlikely, even in the improbable event that bidentate coordination induces strain comparable to that observed for ethylene phosphate. The catalysis of phosphate transfer is also subject to the properties of the medium. The incorporation of the substrate into a micellar phase represents a process analogous to enzymic catalysis. The spontaneous hydrolysis of 2,4- and 2,6-dinitrophenyl phosphate dianions is increased by 25-fold ( a t saturating detergent) upon the incorporation of these esters into the cationic micelle of cetyltrimethylammonium bromide (89). Catalysis is neither observed in the hydrolysis of the corresponding monoanions nor in the reaction between hydroxide ion and the dianions. These results suggest that fragmentation of the dianion into two monoanions is energetically favorable on the micelle surface and/or incorporation of the substrate into a micellar environment simulates an aqueous organic solvent which is known to accelerate dianion hydrolysis rapidly. The lack of micellar assistance to hydroxyl attack has been explained in terms of substrate and nucleophile competing for identical sites which, once occupied, repel the approach of additional reactant molecules. With monoanionic bis-2,4-dinitrophenyl phosphate, the cationic micelle catalyzes hydroxide attack about thirtyfold owing to incorporation of the more hydrophobic ester within the interior of the micelle (90). In summary, significant catalysis by metal ions and/or changes in substrate environment is feasible for either dissociative or associative phosphoryl transfer mechanisms. It is worth emphasizing that the effectiveness of the former does not necessarily depend on the metal ion being positioned as encountered in the normal ion pair. I n fact, the commonly held hypothesis of charge neutralization may act to diminish the efficiency of phosphoryl transfer within an enzyme-substrate complex as illustrated by the data in Table 11. Additionally, the metal ion may affect the stereochemical aspects of the transfer by lowering the energy requirements for pseudorotation processes between dianionic pentacovalent species (91). The latter has as yet not been demonstrated experimentally and, indeed, as argued earlier, is degenerate with alternate modes of substrate binding. 89. C. A. Bunton, E. J. Fendler, L. Sepulveda, and K. Yang, JACS 90, 5512 (1968). 90. G. J. Buist, C. A. Bunton, L. Robinson, L. Sepulveda, and M. Stam, JACS 92, 4072 (1970). 91. A. S. Mildvan. “The Enzymes,” 3rd ed., Vol. 2, p. 480, 1970.

S. J . BENKOVIC AND K. J. SCHRAY

VI. Enzymic Catalytic Mechanisms

An attempt will now be made to delineate probable mechanisms and modes of enzymic catalysis by drawing on the extensive model studies of phosphoryl transfer which have been summarized in the previous sections. Several of the aforementioned mechanism speculations will be augmented. I n Table IV (IS, 2.4, 29, 78, 91a-109) are listed representative compounds of biological interest including those serving either as substrates or functioning as phosphoryl enzyme intermediates. Reaction rates of their monoanions are contrasted to rates observed in selected enzyme reactions involving these phosphoryl compounds. Although hydrolysis is not the enzymic reaction generally taking place, hydrolytic data are uniformly available for these substrates, whereas information on truly analogous nucleophilic displacements are not. These comparisons are intended to give the reader examples of the catalytic effectiveness of various enzymes catalyzing phosphoryl transfer. It is readily seen that there is enormous catalysis brought about by the enzymes contrasted to all cases of phosphoryl transfer from these com91a. T. E. Barman, “The Enzyme Handbook.” Springer-Verlag, Berlin and New York, 1969. 92. C. A. Bunton, D. R. Llewellyn, K. G. Oldham, and C. A . Vernon, JCS, London p. 3588 (1958). 93. D. L. Filmer and D. E. Koshland, BBA 77, 334 (1963). 94. W. J. Ray and E. J. Peck, “The Enzymes,” 3rd ed., Vol. 6, p. 407, 1972. 95. C. A. Bunton and H. Chaimovich, JACS 88, 4082 (1966). 96. Ch. Degani and M. Halmaan, JACS 88, 4075 (1966). 97. V. A. Najjar, “The Enzymes,” 2nd ed., Vol. 6, p. 161, 1962. 98. L. Raijman, S. Grisolia, and H. Edelhoch, JBC 235, 2340 (1960). 99. S. J. Benkovic and K. J. Schray, Biochemistry 7, 4097 (1968). 100. A. Tietz and S. Ochoa, “Methods in Enzymology,” Vol. 5, p. 365, 1962. 101. K. A. Holbrook and L. Ouellet, Can. J. Chem. 35, 1496 (1957). 102. H. S. Penefsky and R. C. Warner, JBC 240, 4694 (1965). 103. M. E. Pullman, H. S. Penefsky, A. Ditta, and E. Racker, JBC 235, 3322 (1960). 104. D. Samuel and B. L. Silver, JCS, London p. 289 (1963). 105. L. Anderson and G. R. Jolles, ABB 70, 121 (1957). 106. B. Edlund, L. Rask, P. Olson, 0. Walinder, 0. Zetterquist, and 1,. Engstrom, Eur. J. Biochem. 9, 451 (1969); R. H. Yue, R . L. Ratliff, and S. A. Kuby, Biochemistry 6, 2923 (1967). 107. L. Noda, S. Kuby, and N. A. Lardy, “Methods in Enzymology,” Vol. 2, p. 605, 1955. 108. H. M. Eppenberger, D. M. Dawson, and N. 0. Kaplan, JBC 242, 204 (1967). 109. H. Follmann, H. J. Wieker, and H. Witzel, Ew. J. Biochem. 1, 243 (1967).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

233

TABLE IV COMPARISON OF HYDROLYTIC RATE COEFFICIENTS WITH ENZYME-CATALYZED TRANSFER Esters

Monoanion hydrolysis min-I ca. 25"

Enzyme-catalyzed transfer min-1 a 6 X lo4 (93,94)

a-Glucose-1-pb

3X

Glucose-6-Pb

4

x

10-8 (95,961

1 . 9 x 104 (94, 97)

Acetyl-P

2

x

10-3 (94)

1 . 7 x 103 (98)

PEP

1

x

10-6

ATP, ADPc

3 x 10-6 (101)

SerineP*

4

x

10-8

Phosphohistidined

0.6

x

10-4 (2.9)

Phosphocreat>ineb

2

x

10-2

Nucleotide diested analog (cis-tetrahydrofuran-4-013-phenyl phosphate)

(9.2)

(phosphoglucomutase) (phosphoglucomutase) (acylphosphatase)

(99)

(104)

(1s)

0 . 6 X lo-' (78)

1 . 0 x 103 (100)

(pyruvate kinase) 3 . 2 x 104 (im,ios) (adenosine triphosphatase) 1 . 9 x 104 (106) (phosphoglucomutase) 1 . 6 x 104 (106) (nucleoside diphosphate kinase) 7 . 2 x 104 (107,108) (creatine kinase)' 1 . 4 X lo6 (109) (ribonuclease)

a Values assume homogeneity of enzyme preparation and utilize the specific activities and molecular weight of these references. For compilation see Barman (91a). b Calculated from E,, of reference cited. c Primarily ADPI- and ATPS-; E. g 25 kcal/mole. Assumed E, 30 kcal/mole. e At 38". Assumed E, g 25 kcal/rnole.

pounds in the absence of enzyme. The question then is simply: What catalytic factors might be utilized to achieve these rates?

A. THEMETAPHOSPHATE MECHANISM As described in Section II,A, all classes of phosphate monoester monoanions or dianions hydrolyze via rates predictable from the pK, of the leaving group with the exception of the corresponding pyrophosphate species which is slightly less reactive than anticipated. Representative biological esters, e.g., acetyl phosphate dianion, the species ADP3- and ATP4- hydrolyze with expulsion of a negatively charged ligand owing to the latter's low pK,. Similarly, the dianion of glucose 6-phosphate, like salicyl phosphate, is hydrolytically labile because of the availability

234

S.

J. BENKOVIC AND K. J. SCHRAY

of a proton from the neighboring acidic hydroxyl group (95, 96). This suggests a major mechanism by which enzymic facilitated phosphoryl transfer may occur, i.e., general acid catalysis to enhance the leaving characteristics of the ligand. Virtually all enzymic reactions occur a t pH values where the completely ionized species predominate for the phosphate monoester substrates or di- or triphosphates. Even for the case of phosphate monoester dianions which hydrolyze more rapidly than their monoanions (pK, < 7), the P-0 cleavage reaction is far more rapid for the monoanion if decomposition of the zwitterionic species is considered separ+ ately. Thus, the rate coefficient for P-0 fission of RXHP0:- ranges from to lo9 min-l for various leaving groups a t 25" as noted previously. It is, however, difficult to envision a simple process for increasing the concentration of the ewitterionic species of a monoester or the unsymmetrical di- or trianionic polyphosphate tautomer by increasing the basicity of the incipient departing ligand without a compensating decrease in its reactivity. A similar argument views the attendant difficulties of increasing the former's concentration by increasing the acidity of the enzymic functional group above that of hydronium ion. Indeed, actual preequilibrium protonation of the leaving ligand probably is not necessary, as seen above for intramolecular systems. If one analogously assumes that the interaction between an acidic enzyme functional group and the leaving ligand generates an assemblage whose pK, for proton dissociation is 6 6 pK, units greater than the free ligand, then observed hydrolytic rates of 10-3-102 min-' may be attained for the esters listed in Table IV. Perturbations of pK, of this order of magnitude have been observed (110). Thus, protonation by an acidic residue concomitant with P-X bond cleavage would a priori constitute the single most effective contribution to catalysis. A related means for enhancement of the leaving group tendency involves metal ion catalysis. This generally may be viewed as a consequence of the increased stability toward protonation of the metal ion ligand complex relative to the free ligand, the difference between the two being a crude index of the overall efficiency of this process. For dianions this dependency is ca. 10-fold per pK, illustrating the high catalytic potential of this mode. Large rate accelerations, 105-10G,attributable to such catalysis in dianion hydrolysis have been observed, although seldom, as of yet, with metal ions commonly required by phosphoryl transfer enzymes. It is obvious that the magnitude of such catalysis will diminish markedly as the ligand is removed from direct coordina110. P. A. Frey, F. C. Kokesh, and F. H. Westheimer, JACS 93, 7266 (1971).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

235

tion with the metal ion. Insofar as metal ion complexation at the a,,& phosphoryl moieties of a triphosphate can approximate protonation, then catalysis of metaphosphate loss from this complex also should be observed. The stability constants of 1 :1 metal-ADP complexes are of the order of lo2 for Ca2+,MgZ+,and MnZ+;thus, even with the assumption that only the desired complex is formed, catalytic effects would not be large ( I l l ) . The stability of such complexes, however, may be increased in the enzymic environment. Since dissociative decomposition of either monoester mono- or dianions must proceed through transition states involving charge separation, large acceleratory effects may be anticipated for these reactions conducted in less polar solvents. At present it is difficult to assess the magnitude of possible catalysis caused by substrate location in hydrophobic regions of low polarity a t the active site. Estimates derived from experiments in mixed solvents are not applicable owing to specific solvent effects, but the micellar studies above indicate increased rate factors of a t least 100-fold and possibly larger. One may therefore reasonably argue that combinations of these modes of catalysis by the enzyme should suffice to attain the listed rate coefficients for the dissociative metaphosphate mechanism. Finally, it is clear that there is no requirement or role for general base catalysis in this mechanism. The P-X bond cleavage is rate determining, the reaction of X-H with metaphosphate is rapid, and solvent could readily absorb the excess proton. Since most nucleophilic displacements on acyclic phosphate monoesters have highly uncoupled transition states, they too may be properly considered within this classification. Thus, the observation of a phosphoryl-enzyme intermediate is neither evidence for nor against a metaphosphate-type transition state on either side of the intermediate. However, the occurrence of this intermediate additionally may signify (a) a stereochemical difficulty in transferring the phosphoryl moiety directly between donor and acceptor molecules or (b) an actual associative mechanism.

B. BIMOLECULAR OR ASSOCIATIVE MECHANISMS Several methods may be envisaged to enable the relatively nonelectrophilic mono- or dianions to undergo nucleophilic attack where the bond-forming and breaking processes in the transition state effectively are coupled or an actual pentacovalent species is formed on the reaction pathway. The simplest is protonation of the phosphoryl oxygen (8). 111. M. M. Taqui Khan and A. E. Martell, JAG'S 89, 5585 (1967).

236

S. J. BENKOVIC AND K. J. SCHRAY

Both intuitively and experimentally protonation is effectively esterification. A second means may be metal ion complexation of the phosphoryl moiety. However, since the reactivity of triesters toward nucleophiles only approximates that of monoester dianions, there seems to be little catalytic advantage to utilize these modes independent of others. Attack by bound anionic nucleophiles within the confines of a ternary complex may likewise be only marginally accelerated. An obvious situation which incorporates within the substrate itself the requisite neighboring group for the operation of a pentacovalent mechanism is the hydrolysis of RNA. Recently, Usher et al. (112-114) have examined the stereochemistry of both the cyclieation and ring opening steps of the ribonuclease reaction to obtain evidence for such possible pentacovalent intermediates. Of the two plausible directions of nucleophilic addition to phosphorus-the “in-line” mechanism of nucleophilic approach 180” from the leaving atom, and the “adjacent” mechanism of approach 90” from the leaving atom-the latter requires the existence of a pentacovalent intermediate which must pseudorotate in order to expel the departing ligand from an apical position. The former does not distinguish between existence of a pentacovalent species and simple bimolecular displacement. The stereochemistry was probed with enzymic transformations of the separated diastereomers of uridine2’,3’-cyclic phosphorothioate, e.g., (XXII) , whose geometry is retained

0

\

/ O

S ,H/Hp\o

(XXII)

in a cyclic reaction sequence if an in-line mechanism is operative for either step. The in-line postulate is in accord with their results so that the question of a pentacovalent intermediate remains unresolved. This finding should not be viewed as suggesting by default the simple bimolecular mechanism. 112. D. A. Usher, E. S. Erenrich, and F. Eckstein, Proc. N a t . Acnd. Sci. U . S . 69, 115 (1972). 113. D. A . Usher, D. I. Richardson, and F. Eckstein, Nature (London) 228, 663 (1970). 114. D. A. Usher, Proc. N a t . Acad. Sci. U.S. 62, 661 (1969).

6.

CHEMICAL BASIS OF BIOLOGICAL PHOSPHORYL TRANSFER

237

A mechanism involving a pentacovalent intermediate need not be limited to diester substrates with suitably juxtaposed functional groups. Various mechanisms of constraint may be envisioned whereby the geometry of the pentacovalent phosphorus may be forced on the substrate: (1) the attacking nucleophile may be positioned as to mimic its being a member of a five-membered ring with either the leaving group and/or a phosphoryl oxygen and (2) the phosphate oxygens and/or leaving group may be forced into such a configuration. Both of these would be expected to facilitate nucleophilic attack by factors of about 106 if they approximate closely a five-membered ring. This same process can, of course, occur for E-P. The coupling between enzyme, donor, and acceptor may be seen as a derivative of the strain theory of catalysis (115). In mode 2 “strain” is manifest in a higher potential energy reactant ground state; in mode 1 the product after ligand loss is of higher potential energy-in relationship to the unbound donor-acceptor. It is clear, however, that the stabilization of the pentacovalent species arises from a minimization of unfavorable nonbonded interactions so that modes 1 and 2 represent limiting means for attaining its intermediacy relative to merely a transition state lifetime. These mechanisms require several binding groups a t the active site or possibly a metal ion to accomplish the pseudocyclization. Moreover, the rather rigid stereochemical requirements for the apical addition and departure of ligands from pentacovalent phosphorus dictate specific orientations for donor-acceptor binding in the absence of pseudorotations. As discussed earlier, there may be a requirement for protonation of the departing ligand contingent on the protonation state of the nucleophile. Furthermore, rate differentials between water and hydroxide attack on cyclic or acyclic phosphorus are of the order of lolo. Thus, enzymic catalysis by proper stereochemical placement of acid-base residues may be anticipated. Two factors may act to reduce this effect: (1) many of the nucleophiles in question possess low pK, values, i.e., are mainly dissociated, and (2) the enzymic functional groups available are of limited acidbase strength. Obviously, the anticipated rate acceleration from the combined catalytic modes would again scale the rate coefficients for the nonenzymic reactions into the enzymic realm. In conclusion, there is little ground on which to select or favor one of the possible mechanisms. Regardless, a general acid catalyst is to be expected a t the active site. If an associative mechanism were operative, additional groups capable of serving as rigid binding sites may be 115. W. P. Jencks, “Cat.alysis in Chemistry and Enzymology,” p. 282. McGrawHill, New York, 1969.

238

S. J. BENKOVIC AND K. J. SCHRAY

expected in positions consistent with the geometry of pentacovalent phosphorus. Indeed, an experimental distinction between the two mechanisms in the case of monoesters may prove difficult since the transition states for both in most situations will involve considerable bond cleavage between phosphorus and the departing ligand ; i.e., metaphosphate formation or decomposition of the pentacovalent intermediate is rate determining. These arguments, deliberately general, should be applicable to a variety of enzyme-catalyzed phosphoryl transfer reactions for which specific examples are discussed in preceding chapters.

Phosphofructokinme DAVID P . BLOXHAM

0

HENRY A . LARDY

I . Introduction . . . . . . . . . . . 11. Purification . . . . . . . . . . . I11. Assay of Phosphofructokinase Activity . . . . . IV. Catalytic Properties . . . . . . . . . A . Phosphoryl Acceptor Specificity . . . . . B. Phosphoryl Donor Specificity . . . . . C. Cation Requirement . . . . . . . D . Kinetic Studies of Enzyme Mechanism . . . E . Isotope Exchange Studies . . . . . . V . Structural Properties . . . . . . . . . A . Molecular Weight . . . . . . . . B . Subunit Structure of Rabbit Muscle P F K . . C . Subunit Structure of Clostridium pasteurianum P F K D. Subunit Structure of Eschenehiu coli P F K . . E . Subunit Structure of Rabbit Erythrocyte P F K . F. Isoenzymes of PFK . . . . . . . G . Reversible Inactivation of PFK by Dilution . . H . Phosphorylation of PFK? . . . . . . VI . Regulatory Properties of P F K . . . . . . VII . Role of Specific Groups in Enzymic Activity . . . A . Thiol Groups . . . . . . . . . B. Role of Histidine . . . . . . . . C . Other Functional Groups . . . . . . VIII . The Role of PFK in the Control of Glycolysis . . . A . The Pasteur Effect . . . . . . . . B . Control of Pyridine Nucleotide Oscillations . . C . Hormonal Control of Glycolysis . . . .

239

. .

. .

. . . .

. . . .

. .

. . . .

. . . . . .

. .

.

.

. .

. . . . . .

. . . .

. . . . . .

.

.

. . . .

. .

240 241 243 244 244 245 247 248 252 253 253 254 256 256 257 257 259 260 261 269 269 272 272 274 274 276 277

240

DAVID P. BLOXHAM AND HENRY A. LARDI’

1. Introduction

Phosphofructokinase (ATP :~-fructose-6-P-1-phosphotransferase, EC 2.7.1.11; hereafter PFK) catalyzes the transfer of the terminal phosphate of ATP to the C-1 hydroxyl of fructose 6-phosphate (F6P), as shown in Eq. ( l ) , to produce fructose 1,g-diphosphate (FDP). The re-

HO

HO

action catalyzed by this enzyme represents the first unique step in glycolysis; hence, it is not surprising ( 1 ) that the enzyme is profoundly regulated by various metabolites in a manner that controls rates of glycolysis in accord with the cells’ need for energy or glycolytic intermediates (8-4a). 1. R. A. Yates and A. B. Pardee, JBC 221, 757 (1956). 2. D. E. Atkinson, Annu. Rev. Biochem. 35, 85 (1966). 3. E. R. Stadtman, Advan. Enzymol. 28, 41 (1966). 4. 0. H. Lowry and J. V. Passoneau, JBC 241, 2268 (1966) 4n. T. E. Mansour, Cum. Top. Cell. Reg. 5, 1 (1972).

5. K.-H. Ling, F. Marcus, and H. A. Lardy, JBC 240, 1893 (1965). 6. A. Parnieggiani and E. G. Krebs, BBRC 19, 89 (1965). 7. A. Pnrmegginni, J. H. Luft, D. S. Love, and E. G. Krebs, JBC 241, 4625 (1966). 8. V. Paetlcnu and H. A. Lardy, JBC 242, 2035 (1967). 9. R. B. Layzer, L. P. Rowland, and W. J. Bank, JBC 244, 3823 (1969). 10. H . Frenkel, ABB 125, 166 (1968). 11. T. E. Mansour, “Methods in Enzymology,” Vol. 9, p. 430, 1966. 12. T. E. Mansour, N. Wakid, and H. M. Sprouse, JBC 241, 1512 (1966). 13. T. E. Mansour and C. E. Ahlfors, JBC 243, 2523 (1968). 14. M. Y. Lorenson and T. E. Mansour, JBC 244, 6420 (1969). 15. R. G. Kemp, JBC 246, 245 (1971). 16. D. J. H. Brock, BJ 113, 235 (1969). 17. N. Kono and K. Uyeda, BBRC 42, 1095 (1971). 18. L. M. Y. Lee, ABB 148, 607 (1972). 19. S. Tnrui, N. Kono, and K. Uyeda, JBC 247, 1138 (1972). 20. A. Sols and M. L. Salas, “Methods in Enzymology,” Vol. 9, p. 436, 1966. 21. T. J. Lindell and E. Stellwagen, JBC 243, 907 (1968). 22. W. .4tzpodien and H. Bode, Eur. J. Biochem. 12, 126 (1970).

7.

PHOSPHOFRUCTOKINASE

24 1

Phosphofructokinase is now available in highly purified form from a number of sources which provides an opportunity to investigate both the catalytic and regulatory mechanisms of the enzyme in molecular detail. Because of this enzyme's molecular complexity, its multifaceted regulation and its importance in controlling rates of glycolysis in normal and neoplastic tissue, it presents a great challenge to biochemists.

II. Puriflcation

Table I summarizes a few properties of the PFK's that have been purified to homogeneity (4-28). Phosphofructokinase is considered a cytosolic enzyme and is found in the soluble fraction of most tissues. However, when PFK was isolated from sheep, cow, and pig hearts, the homogenate showed little activity (12, 2 9 ) . If the inactive particulate fraction, obtained from these homogenates by centrifuging a t 24,000 g for 30 min, was incubated for 2 min a t 37" with ATP and MgSO,, high PFK activity appeared in the soluble fraction. The location of P F K in the particulate fraction is probably an artifact of the morphological changes occurring on transportation of the hearts from the abattoir to the laboratory, since in a homogenate of freshly excised guinea pig hearts the enzyme is located exclusively in the soluble fraction (29, SO). This aggregation with particulate matter and reactivation by ATP and Mg2+is reminiscent of the earlier finding ( 6 ) that P F K activity in phosphate extracts of rabbit muscle is lost during storage and can be regenerated by incubation with MgATP. The loss of activity is also prevented by 0.03 M F-, which simultaneously prevents aggregation to higher molecular weight forms (5). This behavior resembles that of enzymes that are regulated by phosphorylation-dephosphorylation processes ($1); however, there is no evidence that the regulation of P F K is effected in this way (see Section V,H) . A common feature of the enzyme that is used in purification is its stability to heat. Phosphofructokinase from many sources withstands 23. 24. 25. 26. 27. 28. 29. 30. 31.

H. Wilgus, J. R. Pringle, and E. Stellwagen, BBRC 44, 89 (1971). C. C. Griffin, B. N. Houck, and L. Brand, BBRC 27, 287 (1967). D. Blangy, FEBS Lett. 2, 109 (1968). D. Blangy, Biochimie 53, 135 (1971). K. Uyeda and S. Kurooka, JBC 245, 3315 (1970). T. Sumi and M. Ui, BBA 268, 354 (1972). T. E. Mansour, Advan. Enzyme Regul. 8, 37 (1970). T. E. Mansour, JBC 238, 2285 (1963). H. Holzer and W. Duntze, Annu. R e v . Biochem. 40, 345 (1971).

N b P

N

TABLE I PURIFICATION AND PROPERTIES OF PFK

source Skeletal muscle (rabbitp Skeletal muscle (human) Heart muscle (ox) Heart muscle (sheep)O Liver (rabbit) Liver (sheep) Liver (chicken) Erythrocyte (human) Erythrocyte (rabbit) Brain (sheep) Yeast (baker’s) Escherichia coli Clostridium pasteuriunuma Ascites tumor (mouse) a

Specific activity (unit/mg)

Apparent purification

Minimum MW for full activity

120-160 99.5 93 157

145 169 9,300 70 2,600 480

3 . 8 X lo6 (13.1 S) (13.7-14.8 S) (15.2 S)

48

18.5 114 34 139 18 116 190 160 150

Indicates that the enzyme has been crystallized.

-

745 20,400 238 527 890 444 150

-

-

(13.9 S) (8.8-23.3 S) 5 x 105 5 . 9 x 105 (16.3 s) 1.42 X lo5 ( 7 . 8 S) 1.44 x 105 (7.8 s) 3 x 105

References 5-8 9 10 11-14 15 16 17 9,18 19

4 20-23 24-26 27 28

P

2 tl ? W

E : z

z 9

3! 24 *

7. PHOSPHOFRUCTOKINASE

243

heating to 40"-60" for 30 min without loss of activity. The most extreme case is the enzyme from Flavobacterium thermophilum where the catalytic activity survives 80" for 2 hr (32,33). The enzyme from chicken liver is inactivated by decreasing temperature (60 and 80% loss in 2 hr at 8" and 0", respectively) which must be recognized in purification (17); high protein concentration and polyvalent anions protect against inactivation, whereas monovalent anions enhance inactivation. The inactivation a t lower temperatures results from dissociation of the enzyme since the enzyme has a sedimentation coefficient of 13.9s at 25" which is reduced to 5.4 S a t 4" (17). Phosphofructokinase has been successfully crystallized from several sources. The original method for crystallization, developed by Parmeggiani and Krebs (6) for rabbit muscle PFK, involves increasing the concentration of ammonium sulfate to 40% saturation in the presence of ATP. For the most part, P F K will crystallize only in the presence of ATP. Recently, Uyeda and Kurooka (97)have shown that the enzyme from Clostridium pasteurianum can be crystallized in the absence of ATP; however, in this case, crystals form much more slowly.

111. Assay of Phosphofructokinase Activity

Essentially, there are two methods for assaying P F K activity. I n the first, the formation of product is coupled through auxiliary enzymes to a reaction involving oxidation or reduction of diphosphopyridine nucleotide which can be followed spectrophotometrically or fluorometrically. Alternatively, the reaction can be monitored in a p H stat, for at pH 8.5, Eq. (1) produces 1 mole of H+ per mole of product (@), The choice of assay depends upon the particular requirements of the experiment. The most common assay involves linking FDP production through aldolase and triosephosphate isomerase to D P N H oxidation by a-glycerophosphate dehydrogenase. This reaction has a stoichiometry of 2 moles of DPNH oxidized per mole of FDP produced. The reaction often shows a lag phase during which the rate increases until a linear rate is achieved. This effect may result from FDP accumulation which causes enhancement of PFK activity. It can be minimized by assaying a t low 32. M. Yoshida, T. Oshima, and K. Imahori, BBRC 43, 36 (1971). 33. M. Yoshida, Biochemistry 11, 1087 (1972). 34. J. E. Dyson and E. A . Noltmann, Anal. Biochem. 11, 362 (1965).

244

DAVID P. BLOXHAM AND HENRY A. LARDY

concentrations of P F K relative to that of the auxiliary enzyme ( 4 ) . Alternatively, the initial rate may be obtained from plots of time versus log rate, which allows the rate a t zero time to be determined by extrapolation (35). A requirement of this assay is that the enzyme should be completely free of FDPase activity. When purification of P F K is undertaken from liver, where the FDPase activity is much higher than the activity of PFK, this is not the case (16, 36). I n these conditions, it is necessary to assay the production of ADP which can be linked to D P N H oxidation via pyruvate kinase and lactate dehydrogenase.

IV. Catalytic Properties

A. PHOSPHORYL ACCEPTOR SPECIFICITY Phosphofructokinase is fairly specific for F6P, phosphorylates D-tagatose 6-P a t about half the rate with F6P (37’), and uses D-fructose l-P (38),D-glucose l-P (39),and sedoheptulose 7-P (37) only slowly a t reasonable concentrations. L-Sorbose l-P, L-sorbose 6-P, and D-ribulose Benkovic and Mildvan 5-P are not detectably phosphorylated (39~). (4.0) have modified their earlier conclusion concerning the preferred anomeric conformation of the substrate (41). P-D-FGPis rapidly used by PFK, and it has not yet been established whether the a anomer is utilized directly or only after mutarotation. Uyeda (38) has shown that rabbit muscle P F K catalyzed the phosphorylation of fructose l-P a t 5% of the rate of F6P. The relative capacity of the enzyme to phosphorylate fructose l-P and F6P was similar through purification, chromatography, and inactivation. Fructose l-P was found to be a competitive inhibitor of the phosphorylation of F6P indicating that both sugars compete for the same active site. Space-filling models indicate that a-D-fructose l-P more closely resembles P-D-FGP than does the p-D-fructose l-P anorner. It is likely that the a-fructose l-P fits on the enzyme with the phosphate group normally located and the sugar moiety “upside down.” Using this reason35. C. I. Pogson and P. J. Randle, BJ 100, 683 (1966). 36. A. H. Underwood and E. A. Newsholme, BJ 95, 868 (1965). 37. E. L. Totten and H. A. Lardy, JBC 181, 701 (1949). 38. K. Uyeda, JBC 247, 1692 (1972). 39. P.Eyer, H.W. Hofer, E. Krystek, and D. Pette, Eur. J . Biochem. 20, 153 (1971). 39a. H.A. Lardy, “The Enzymes,” 2nd ed., Vol. 6,p. 67, 1962. 40. S. J. Benkovic and A. S. Mildvan, personal communication. 41. K. J. Schray, S. J. Benkovic, P. A. Benkovic, I. A. Rose, and A. S. Mildvan, Fed. Proc., Fed. Amer. Soc. Exp. Biol. 31, 419 (1972).

7.

PHOSPHOFRUCTOKINASE

245

ing, D-fructose l-P may be considered as an analog of p-D-fructose 6-P and L-sorbose l-P as an analog of the (Y anomer. Since L-sorbose l-P is not a substrate, it would appear that a-D-fructose 6-P is also inactive. An interesting contrast exists for the bacterium Aerobacter aerogenes where fructose metabolism proceeds through the formation of fructose l-P by the enzyme PEP: D-fructose-l-phosphotransferase (4.2) rather than through the formation of F6P. This organism possesses a unique form of PFK that uses fructose l-P as the preferential substrate and catalyzes phosphorylation a t C-6 ( 4 3 ) . This enzyme is distinct from the enzyme that phosphorylates F6P, and F6P is a competitive inhibitor of the phosphorylation of fructose 1-P. Using [ Y - ~ ~ P ] A T and P glucose l-P as a substrate, Eyer et al. (39) demonstrated that rabbit muscle P F K catalyzed the formation of a compound that was chromatographically identical to glucose 1,6-diP. The phosphorylation proceeded a t Q.776 of the rate of phosphorylation of F6P. Glucose 1,6-diP is an essential catalytic component of the phosphoglucomutase reaction, and it is possible that the P F K mediated phosphorylation of glucose 1-P could constitute an important route for its synthesis. Another enzyme, isolated from rabbit muscle and yeast extracts, catalyzes the ATP-dependent phosphorylation of glucose l-P (44, 46). This enzyme is distinct from PFK since i t does not catalyze the phosphorylation of F6P or fructose l-P ( 4 5 ) .

B. PHOSPHORYL DONOR SPECIFICITY I n contrast to the high degree of specificity for sugar phosphates, P F K can use a wide variety of nucleoside triphosphates as phosphoryl donors in the catalytic reaction. Table I1 shows the apparent K,,, values of nucleoside triphosphates for the enzymes from yeast, E . coli, and muscle (20, 46-48). Muscle P F K can also use 2-amino-9-/3-~-ribofuranosylpurine-5’-triphosphate (49), 6-mercapto-9-/3-~-ribofuranosy~purine-5’triphosphate (491, and 1,N6-etheno-ATP ( 5 0 ) .For yeast PFK, changing 42. T. E. Hanson and R. L. Anderson, Proc. Nut. Acad. Sci. U . S. 61, 269 (1968). 43. V. Sapico and R. L. Anderson, JBC 244, 6280 (1969). 44. A. C. Paladini, R. Caputto, L. F. Leloir, R. E. Trucco, and C. E. Cardini, ABB 23, 55 (1949). 45. L. F. Leloir and R. E. Trucco, “Methods in Enzymology,” Vol. 1, p. 354, 1955. 46. K.-H. Ling and H. A. Lardy, JACS 76, 2842 (1954). 47. K. Uyeda and E. Racker, JBC 240, 4682 (1965). 48. D. Blangy, H. Buc, and J. Monod, J M B 31, 13 (1968). 49. D. P. Bloxham, unpublished observations. 50. J. A. Siecrist, J. R. Barrio, and N. J. Leonard, Science 175, 646 (1972).

APPARENT

Rabbit muscle Yeast E . wli a

30 20 60

TABLE I1 MICHAELIS CONSTANTS FOR NIJCLEOSIDE TRIPHOSPH.4TES

a

100 1200

70 200 0

a

400 2000

33 800 2000

These nucleoside triphosphates are substrates, but comparable K , values are not available.

-

-

-

-

80

3500

4G47 90

48

fd

k

3c

E

7.

PHOSPHOFRUCTOKINASE

247

the nucleoside triphosphate alters the p H optimum of the enzyme (bf ) . With ATP, the enzyme has maximum activity a t p H 7.8, whereas with ITP, the enzyme shows two pH maxima at 7.6 and 8.2. Not all enzymes show such a wide nucleotide specificity. Thus, Dennis and Coultate have shown that P F K from brussels sprouts uses purine nucleoside triphosphates (ATP, GTP, and ITP) preferentially; pyrimidine nucleoside triphosphates are poor phosphoryl donors (61). As will be discussed in greater detail later in this review, increasing the concentration of ATP above the catalytic optimum causes an inhibition of PFK activity. The structural requirements for this inhibitory phenomenon appear to be somewhat more stringent than for catalytic activity. Muscle PFK is inhibited by ATP, CTP, and UTP but is not inhibited by ITP despite the fact that this nucleotide serves as a substrate (47).Yeast P F K appears to show even more specificity since, so far, only ATP has been shown to be inhibitory; G T P and ITP do not inhibit this enzyme (81,52, 53). Furthermore, ITP participates in the catalytic reaction but does not inhibit E. coli PFK (64). C . CATION REQUIREMENT All phosphoryl transfer reactions involving ATP require a divalent cation-ATP complex as the active substrate. The Mgz+complex is usually the most effective. Brain PFK uses Mg?+,Mnz+,and Co2+with apparent K , (6 mM ATP) values of 1.2, 0.6, and 1.8 mM, respectively (56). The relationship between ATP and Mgz+ plays an important role in enzymic activity. At high concentrations, divalent cations can be inhibitory (K,Mg2+,4 mM; KICa2+,0.37 mM) for brain P F K ; however, there is evidence to show that free Mgz+is required for catalytic activity (56).This was first Aoted with the muscle enzyme at pH 7, where activity of the enzyme was negligible unless the concentration of free Mg2+ exceeded that of ATP (8, 67). I n experiments with yeast PFK, Mavis and Stellwagen (66)calculated the concentrations of free Mgz+ and of MgITP present a t varying total concentrations of both Mg2+and ITP. The catalytic activity was clearly 51. D. T. Dennis and T. P. Coultate, BBA 146, 129 (1967). 52. E. Vinuela, M. L. Salas, and A. Sols, BBRC 12, 140 (1963). 53. A. Ramaiah, J. A. Hathaway, and D. E. Atkinson, JBC 239, 3619 (1964). 54. D. E. Atkinson and G . M. Walton, JBC 240, 757 (1985). 55. J. A . Muntz, ABB 42, 435 (1953). 56. R. D. Mavis and E. Stellwagen, JBC 245, 674 (1970). 57. H. A. Lardy and R. E. Parks, in “Enzymes: Units of Biological Structure and Function” (0. H. Gaebler, ed.), p. 584. Academic Press, New York, 1956.

248

DAVID P. BLOXHAM AND HENRY A. LARDY

dependent on the concentration of both free Mg2+ and MgITP. In addition to a catalytic role for free Mg2+,this cation may, in some cases, enhance the rate of the P F K reaction by decreasing the concentration of free ATP which is a more potent inhibitor than is MgATP (4, 8, 24, S9a, 57, 58). However, there is evidence that MgATP rather than free ATP is the effective inhibitor of yeast PFK (56). As pointed out by Lowenstein ( 5 9 ) ,nonenzymic transphosphorylations are often enhanced by monovalent cations. They may act either by substituting for divalent cations or by neutralizing the excess negative charge remaining on the MgATP complex. Phosphofructokinase conforms to this general pattern since the enzyme from many sources can be activated by either K+ or NH,+ ions (4, 8, 22, 55, 56, 60-62) ; NH,+ stimulated the activity of muscle PFK a t very low concentrations but became slightly inhibitory a t higher concentrations, and K+ had a lower affinity than NH,+ but produced a higher maximum velocity (8). Yeast PFK is also activated by both NH,+ and K+ and apparently there are two enzyme-cation complexes for each cation ( 6 6 ) .The preferred activating cation is K+ since (a) it is effective in physiologically occurring concentrations ; (b) it shows a normal activity-concentration relationship; and (c) in solutions containing Pi, it does not form insoluble complexes as ammonium ion does with Mg2+and Pi.

D. KINETICSTUDIES OF ENZYME MECHANISM Studies of the kinetic mechanism of PFK are complicated by the fact that the catalytic activity in the forward direction is a function of a t least four species, namely, MgATP, F6P, free Mgz+ and K+. The influence of Mg'+ and K+ can be minimized by using these in excess and treating the reaction as a two-substrate system. The substrates and products may exert regulatory effects as well as catalytic effects. For studies of mechanism, regulatory phenomena can be reduced by making kinetic measurements at a pH where regulatory effects are negligible. This approach has been widely used for mammalian PFK's where the enzyme generally loses its susceptibility to regulation as the pH becomes more alkaline (>7.6). Alternatively, kinetic studies could be made on PFK preparations that are not regulated. Such enzymes are known, but 58. 59. 60. 61. 62.

D. T. Dennis and T. P. Coultate, BBRC 25, 187 (1967). J. M. Lowenstein, BJ 75, 269 (1960). S. L. Abrahams and E. S. Younathan, JBC 246, 2464 (1971). A. H. Underwood and E. A. Newsholme, BJ 104, 296 (1967) D. D. Hoskins and D. T. Stephens, BBA 191, 292 (1969).

7.

PHOSPHOFRUCTOKINASE

249

unfortunately their kinetic properties have not been evaluated extensively. For two-substrate systems, measuring initial velocities in the presence of several fixed concentrations of one substrate while the concentration of the second is varied yields reciprocal plots, the slopes and intercepts of which can be used to predict the enzyme mechanism (63). Plots of this type for P F K from rabbit muscle (64), ox heart ( 6 5 ) ,yeast (52), Dictyosteliuin discoideum ( 6 6 ) ,human skeletal muscle ( 9 ) , human erythrocytes (9), calf lens (67), and Flavobacterium thermophilum (33) produced a series of apparently parallel lines. Sheep heart PFK when studied in the reverse direction gave a similar result (68). Parallel lines are often indicative of a ping-pong mechanism in which the first substrate reacts with the enzyme to produce a free modified enzyme which can then react with the second subdrate yielding the final product. The demonstration that the reciprocal plots produce apparent parallel lines is not conclusive evidence for a ping-pong mechanism. The initial rate equation for a sequential mechanism involving the formation of a ternary complex between enzyme, MgATP, and F6P is of the form

shown in Eq. (2) where K i M g A T P is the dissociation constant of MgATP from the enzyme and K M g A T p and KFcp are the respective Michaelis constants for MgATP and F6P. However, if the constant term of the denominator [containing K i M g A T p in Eq. ( 2 ) ] is much smaller than the other terms, the rate equation reduces to that for a ping-pong mechanism ( 6 3 ) . Therefore, when the variable substrate is employed a t concentrations much greater than its Km, apparently parallel lines will always be observed. In this laboratory, it has been shown (69, 7 0 ) that for rabbit muscle PFK, the value of K i M g A T p (5 p M ) is smaller than KMg:ATP (20 p M ) and KFcP(21 p M ) . I n agreement with this, a t low substrate concentrations, a series of intersecting lines were obtained for the reciprocal plots using rabbit muscle PFK (69,7 0 ) . Intersecting lines were also obtained for PFK from brain ( 4 ) , Lactobacillus casei (71), W. W. Cleland, “The Enzymes,” 3rd ed., Vol. 2, p. 1, 1970. K. Uyeda, JBC 245, 2268 (1970). E. C. Hulme and K. F. Tipton, BJ 122, 181 (1971). P. Baumann and B. E . Wright, Biochemistry 7, 3653 (1968). M. F. Lou and J . H. Kinoshita, B B A 141, 547 (1967). M. Y. Lorenson and T. E. Mansour, JBC 243, 4677 (1968). R. L. Hanson, Fed. Proc., Fed. Amer. Soc. Exp. Biol. 29, 408 (1970). R. L. Hanson, F. B. Rudolph, and H. A. Lardy, JBC (1973) (submitted for publication). 71. H. W. Doelle, B B A 258, 404 (1972). 63. 64. 65. 66. 67. 68. 69. 70.

250

DAVID P. BLOXHAM A N D H E N R Y A. LARDY

TABLE I11 PRODUCT INHIBITION PATTERNS FROM DIFFERENT MECHANISMS FOR PFK Substrate" ~

Mechanism

Product inhibitor

MgATP

F6P

Ping-pong

FDP MgADP

C NC

NC C

Random addition

FDP MgADP

NC C

C NC

Ordered sequential*

FDP MgADP

NC C

NC NC

,. Abbreviations: C, competitive; and NC, noncompetitive.

* MgATP bin& first; MgADP leaves last.

Lactobacillus plantarwn (71), and Aerobacter aerogenes (43) using fructose 1-P as a substrate. Uyeda (38) reevaluated his experiments on rabbit muscle PFK using fructose 1-P as a substrate and also found that the reciprocal plots were intersecting. The presently available information strongly suggests that the reaction mechanism for PFK involves the formation of a ternary complex and is not ping-pong. A useful tool in the study of reaction mechanisms is the use of product inhibition patterns ( 6 3 ) . The predicted inhibition patterns for three possible mechanisms are shown in Table 111. Hanson (69),using F6P, and Uyeda (38),using fructose 1-P as a substrate, found that for rabbit muscle PFK the product inhibition pattern corresponded to an ordered sequential mechanism. Kee and Griffin (79)have found that the pattern corresponds to a random addition mechanism. The difference in these two results is in the nature of the inhibition by FDP with respect to F6P. A tentative explanation of this difference can be made. Scheme (I) shows a diagrammatic representation of the hypothetical orientation of the substrate and products on the enzyme surface. This scheme is analogous to that of Reynard et a2. (73) for the binding of substrates to pyruvate kinase. If the representation shown in Scheme (I) is correct, then FDP overlaps both the ATP and F6P binding sites. At low ATP concentration when ATP sites are free, FDP could influence binding at both ATP and 72. A. Kee and C. C. Griffin, ABB 149, 361 (1972). 73. A. M. Repard, L. F. Hass, D . D . Jacobsen, and P. D. Boyer, JBC 236, 2277 (1961).

7.

251

PHOSPHOFRUCTOKINASE

I

I

I

I

I

I

I

I

I

I

I l-,F6P

Site

I

I I

I

7 -. i -

I

ATP Site-

I

I

I-FDP

Site

I I

I

I

I

-

ADP Site-

I -

I

SCHEME I. Diagrammatic representation of the orientation of substrates and nmdiirta nf

PPK

F6P sites and the inhibition should be noncompetitive with respect to F6P. However, a t higher ATP concentrations when all ATP sites are occupied, F D P may influence the binding of only F6P and the inhibition should be competitive with respect to F6P. In agreement with this suggestion, Kee and Griffin (79)used a high ATP concentration (50 rJM) to obtain competitive inhibition; whereas Hanson (69, 70) used a low ATP concentration (20 &) to obtain noncompetitive inhibition. The actual order of addition of substrates was examined further by Hanson e t al. (70) using the substrate analog, arabinose 5-P. It was found to be a competitive inhibitor of both F6P and FDP in the forward and reverse reactions, respectively, whereas it was noncompetitive with both ATP and ADP. This result combined with the competitive inhibition by MgADP with respect to MgATP suggests that the reaction sequence

I

252

DAVID P. BLOXHAM AND HENRY A. LARDY

is random rather than sequential. Further confirmation comes from the use of guanosine 5’-diP as an alternative substrate inhibitor. If ATP production from ADP is measured in the reverse reaction, the addition of guanosine 5’-diP lowers ATP synthesis in a manner which is competitive with respect to ADP. Guanosine 5’-diP was found to be noncompetitive relative to FDP which is consistent with a random mechanism. If the reaction mechanism were ordered, double reciprocal plots should have yielded nonlinear lines (74). Instead, the lines were linear (70).

E. ISOTOPE EXCHANGE STUDIES A reasonable reaction sequence for a ping-pong mechanism for P F K is shown in Eq. ( 3 ) . This scheme predicts that it should be possible to Mgmp

MgATP

E

E * MgATP

F6P

(E * P)

FDP

(EP F6P)

E

demonstrate an isotope exchange between ATP and ADP and between F D P and F6P. In the case of E . coli PFK, Blangy (96)did not detect any significant isotope exchange for either substrate pair. Ox heart PFK (66)and rabbit muscle P F K (64, 69, 70) both catalyzed ADPATP exchange at approximately 1% of the forward reaction rate. This was shown to be a genuine result since omission of either Mg2+or the enzyme abolished the exchange (66). Both enzymes also catalyzed a very slow exchange between F6P and FDP. For the heart enzyme, the exchange was dependent on MgATP which is inconsistent with a pingpong mechanism (66). Although these exchange reactions appear to be real, they are too low to be consistelit with a ping-pong mechanism for PFK. Possibly they represent some form of side reaction catalyzed by the enzyme. Hexokinase (yeast and brain), which does not involve pingpong mechanism, catalyzes a similar slow ADP-ATP exchange which does not appear to occur at the normal catalytic site (75-78). 74. 75. 76. 77. 78.

F. B. Rudolph and H. J. Fromm, Biochemistry 9, 4660 (1970). A. Kaji and S. P. Colowick, JBC N O , 4454 (1965). H. J. Fromm and J. Ning, BBRC 32, 672 (1968). F. Solomon and I. A. Rose, ABB 147, 349 (1971). D. L. Purich and H. J. Fromm, ABB 149, 307 (1972).

7.

PHOSPHOFRUCTOKINASE

253

V. Structural Properties

A. MOLECULAR WEIGHT (7th) The values of the minimum molecular weight of P F K from a number

of sources were presented in Table I. The molecular weight for most of these preparations is in the range 3 X lo5 to 6 X lo5. The enzymes from Clostridium pusteurianum (27) and E . coli (25) are much smaller and have molecular weights of 1.4 X lo” (7.8s). Two features appear to be common to all preparations examined so far: First, there appear t o be multiple forms of the active enzyme, and, second, the enzyme appears to have a complex subunit structure. I n the Ultracentrifuge, rabbit muscle PFK gave three peaks corresponding to s20,wvalues of 13.8 S, 20.9 S, and 31 S (6). This pattern was obtained when the protein concentration was varied in the range of 5-15 mg/ml. Centrifugation on a sucrose density gradient also revealed that the enzymic activity of this same preparation was heterogeneous. When the protein concentration was decreased, the enzyme revealed only a single peak. The minimum molecular weight of the smallest fully active species of rabbit muscle PFK, assayed in the forward reaction, has been estimated as 380,000 (13.1S) (8). The heterogeneous pattern is not the result of enzyme impurity but results from the existence of the enzyme in several aggregation states (80). Support for this concept is provided by the demonstration that when ultracentrifugation was performed in 2 M urea a single symmetrical peak at 13.7s was obtained. When the urea was removed, the heterogeneous pattern was again obtained (5). Heterogeneity in the ultracentrifuge depends upon the buffer used. In 50 mM glycerophosphate containing 2 mM EDTA, p H 7.2, the protein sediments as a single peak, with sedimentation coefficients between 27 S and 36 S, depending on protein concentration (7). However, in 100 mM potassium phosphate, p H 8, 0.2 mM EDTA and 0.2 mM FDP, the three-component pattern is obtained ( 7 ) . The tendency to aggregate is more apparent in purified muscle P F K that has not come into contact with a thiol reducing agent (molecular weight, 1.6 X lo6) (7, 8 ) . The molecular weight immediately reduces to 3.8 X lo5 on exposure to di78a. We should like to urge a halt to the current epidemic fallacy of reporting molecular weights in daltons. This can even lead to reporting the “ethoxyformylation of nearly 4 moles of histidine per 105 daltons” (79, p. 5533) of PFK I Avogadro, requiescat in pace. 79. B. Setlow and T. E. Mansour, JBC 245, 5524 (1970). 80. C . Frieden, Annu. Rev. Bbchem. 40, 653 (1971).

254

DAVID P. BLOXHAM AND HENRY A. LARDY

thiothreitol (8). The tendency of PFK to aggregate may be related to an interaction of thiol groups since p-hydroxymercuribenzoate decreased the molecular weight of muscle PFK from 36 S to 13.1 S (7). An excellent demonstration of the reversible association-dissociation phenomenon has been provided for human erythrocyte PFK (81). When the purified enzyme was chromatographed on Sepharose 4E! it eluted as five separate peaks, all possessing activity, which had molecular weights that were integral multiples of 220,000. If a single peak was isolated and subjected to a second chromatography then it gave rise to five peaks again, clearly demonstrating that PFK exists as an equilibrium mixture of the various aggregation states. It seems probable that the high molecular weight aggregates of P F K are an artifact of the high protein concentrations that can be obtained with the purified enzyme. This is clearly shown for rabbit erythrocyte PFK where the purified enzyme has a molecular weight of 5 X lo6 (80 S), whereas in a fresh hemolysate the molecular weight is 5 X lo6 (19).

B. SUBUNITSTRUCTURE OF RABBITMUSCLE PFK Catalytically active rabbit muscle PFK of molecular weight 380,000 (13.1 S) is easily dissociated into a form with a sedimentation coefficient of 7 5 by decreasing the protein concentration at pH 6.7 or acidification to pH 5.8 in the presence of 0.8 M urea (8). This dissociation is reversible since the enzyme can be made to reaggregate under appropriate conditions. Dissociation results in loss of 95% of the enzymic activity, but a residual activity of at least 1-276 appears to be retained by the protein of lower molecular weight (8). Reaggregation of the enzyme results in the return of normal enzymic activity. On exposure to 4 m M sodium dodecyl sulfate at pH 11, the enzyme dissociates to a form with a molecular weight of 93,000 (8). Kemp and Krebs (82) have shown that 90,000 daltons of the enzyme are able to bind one molecule of either F6P, AMP, cyclic AMP, or ADP, and suggested that the protomer of P F K (“the identical subunits associated with an oligomeric protein”) is the component with a molecular weight of 90,OOO. 81. K.-W. Wenzel, G. Zimmerman, J. Gauer, W. Diezel, G. St. Liebe, and E. Hoffmann. FEBS Lett. 19, 285 (1972). 82. R. G. Kemp and E. G. Krebs. Biochemistry 6, 423 (1967).

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The earlier work of Paetkau et al. (83) indicated that PFK could be degraded to smaller subunits by high molarity guanidine or urea solutions. I n either 5.5 M guanidine hydrochloride or 8 M urea, the molecular weight decreased to 47,000 (2.3s). A further apparent decrease was reported (83)to be achieved by raising the concentration of guanidine hydrochloride to 6 M or higher. However, in more recent work in this laboratory (84) and by Leonard and Walker ( 8 5 ) ) molecular weights below 50,000 have not been observed. The possibility remains that dissociation to the 24,000 molecular weight species requires some unique condition that we are now unable to repeat or, more likely, that the guanidine used in the earlier work degraded the enzyme as has recently been reported by Fosmire and Timasheff (86) for lactate dehydrogenase. The studies of Reed et aZ. ( 8 4 ) ,using sedimentation equilibrium ultracentrifugation and gel chromatography in 6.5 M guanidine hydrochloride showed that the minimum molecular weight was 67,000 -I- 5,000 which compares reasonably to the value of 76,000 k 5,000 obtained by Leonard and Walker (86). If the molecular weight of the subunit is about 67,000, the active form of the enzyme (MW 380,000) is probably a hexamer and the acid- or dilution-dissociated form of the enzyme corresponds to a dimer with a molecular weight of 134,000. Using the formula developed by Paetkau and Lardy ( 8 ) , where molecular weight = 5.4 X lo3 X ( s ~ " X, ~ 1013)1.rrn, it can be predicted that the observed s ~ of ~this, species ( 7 s ) would give a molecular weight of 136,000. There are still discrepancies which must be sorted out before this concept of the enzymes' structure can be accepted. First, acrylamide gel electrophoresis of P F K in sodium dodecyl sulfate gives an homogenous protein with an apparent molecular weight of 90,OOO ( 8 4 ) . Second, tryptic digestion of PFK yields 50 peptides (19,83). Since there are 50 lysine and arginine residues per 47,000 daltons of enzyme, this suggests that the minimum molecular weight is 47,000. Possibly this discrepancy could result from the presence of repeated peptide sequences. At present it appears that this problem will not be resolved until the primary sequence of PFK is elucidated. The amino acid analyses of muscle PFK performed in three different laboratories (7,19,8s) are in good agreement apart from small discrepancies in the content of threonine, serine, proline, methionine, and aspartic acid. In all cases, the amino acids recovered do not account for 83. V. H. Paetkau, E. S. Younathan, and H. A. Lardy, JMB 33, 721 (1968). 84. 3. Reed, D. P. Bloxham, and H. A. Lardy, unpublished observations. 85. K. R. Leonard and I. 0.Walker, Eur. J . Bwchem. 26, 442 (1972). 86. G. J. Fosmire and S. N. Timasheff, Biochemistry 11, 2455 (1973).

~

256

DAVID P. BLOXHAM A N D H E N R Y A. LARDY

the total weight of P F K ; thus, it would appear that other components contribute to the total structure. At present, little information is available on the possible carbohydrate or lipid content of the enzyme. Paetkau et al. (83) were unable to detect any free N-terminal or C-terminal amino acids by conventional chemical or enzymic methods.

C. SUBUNITSTRUCTURE OF Clostridium pasteuriunum P F K Clostridium pasteuriunum P F K has been purified to the stage of crystallization and was homogeneous during equilibrium centrifugation (87). The minimum molecular weight of the active enzyme is 144,000 (7.8S) . Exposing to 7 M guanidine hydrochloride, 4-8 M urea or reaction of the protein with maleic anhydride resulted in the formation of a species with a molecular weight of 35,000.This species was judged to be homogeneous by acrylamide gel electrophoresis of the maleylated P F K or by disc gel electrophoresis of the denatured enzyme in 8 M urea. Amino acid analysis showed that the amino acids constitute 92% of the dry weight of the protein again suggesting that the protein contains some additional structural component. The enzyme contains approximately 34 arginine and lysine residues per 35,000 daltons. Tryptic digestion resulted in the formation of 33-38 peptides indicating that the subunits of molecular weight 35,000 are identical. Spraying the peptide map with Sakaguchi reagent demonstrated that there were 15 argininecontaining peptides which is in reasonable agreement with the observed value of 18 arginine residues per 35,000daltons. The evidence indicates that P F K from C . pusteuriunuin is a tetramer composed of four identical subunits. This structure is inherently different from that of muscle P F K which may be responsible for the different regulatory properties of these two enzymes (see Section VI). D. SUBUNIT STRUCTURE OF Escherichia coli P F K The structure of PFK from E . coli is very similar to that from C. pasteuriunum. I n 0.1 M tris-HC1 buffer, pH 8.2,the enzyme sediments as a single symmetrical peak with an s ? ~value , ~ of 7.8 S over the protein concentration range 0.5-5 mg/ml ( 2 5 ) . Sedimentation equilibrium indicated a molecular weight of 142,000.I n 6 M guanidine hydrochloride the molecular weight decreased to 36,500 which is the smallest detectable subunit. Disc gel electrophoresis in 0.1% sodium dodecyl sulfate revealed two proteins with molecular weights of 68,000 and 35,000.Presumably the form with a molecular weight of 68,000 corre-

7.

PHOSPHOFRUCTOKIN ASE

257

sponds to a dimeric form of the enzyme. The available evidence indicates that E . coli PFK is a tetramer. No information is available a t present on the structural similarity of the subunits.

E. SUBUNITSTRUCTURE OF RABBIT ERYTHROCYTE PFK Purified rabbit erythrocyte PFK (19) in 50 mM tris-phosphate, p H 8, 1 mM, EDTA, and 20 mM dithiothreitol tends to sediment as a large aggregate with a molecular weight of 5 X lo6 (80s). However, in a fresh hemolysate, enzymic activity sediments with a molecular weight of 5 X lo5 when analyzed by centrifugation on a sucrose density gradient. This is the smallest molecular weight of the active species that has been detected. Exposing the enzyme to 7 M guanidine hydrochloride and 100 mM 2-mercaptoethanol for 24 hr at 4" resulted in the formation of an homogeneous protein with a molecular weight of 53,000, which corresponds to the smallest detectable subunit. Comparative analysis of rabbit erythrocyte and muscle P F K revealed that the amino acid compositions differed by more than could be accounted for by experimental error, particularly for isoleucine, serine, and threonine. Rabbit erythrocyte PFK contains 46 lysine and arginine residues per 50,000 daltons. Tryptic digestion resulted in the formation of approximately 50 peptides indicating that the subunits of erythrocyte PFK are probably identical. When the peptide maps of muscle and erythrocyte P F K were compared there were significant differences, again emphasizing the structural dissimilarity of the two proteins. This aspect will be discussed further in the section on isoenzymes.

F. ISOENZYMES OF P F K The initial indication of the possible existence of isoenzymes of P F K came from the description of an inherited muscular disease in humans that is characterized by a decrease in muscle P F K activity (87, 88). In patients with this disease, the muscle P F K activity reduced to zero, erythrocyte P F K levels were half of normal, and the white blood cell PFK level was unaffected (88). Since this disease is a genetic defect, it follows that the enzymes from these sources are coded differently. The 87. S. Tarui, G. Okuno. Y. Ikura, T. Tanaka, M. Suda, and M. Nishikawa, BBRC 19, 517 (1965). 88. K . B. Layzer, L. P. Rowland, and H. M. Ranney, Arch. Neurol. (Chicago) 17, 512 (1967).

258

DAVID P. BLOXHAM AND HENRY A. LARDY

properties of the enzymes from various human tissues were studied by chromatography on DEAE-cellulose and by their reaction with human muscle P F K antibody (9, 89, 90).Muscle, thyroid, and brain P F K were all eluted as single peaks a t 0.21M buffer (tris-phosphate) and all gave nearly complete precipitation with antibody. Erythrocyte P F K eluted a t a different position on the column (0.32 M buffer), and its sensitivity to antibody precipitation was markedly reduced. White blood cell and platelet PFK both eluted as two distinct peaks at 0.21 and 0.32 M buffer. Only 15% of the white blobd cell PFK was precipitated by muscle antibody, and platelet PFK did not precipitate. Collectively these results emphasize that there are structural variations in human PFK’s from different tissue sources. Rat muscle and liver PFK’s have been clearly separated by chromatography on DEAE-cellulose (91). Kemp (16) compared the regulatory properties of rabbit PFK from muscle and liver and found the liver enzyme was less inhibited by ATP; less sensitive to activation by AMP, ADP, and cyclic AMP; less inhibited by citrate; and more sensitive to 2,3-diphosphoglycerate inhibition. He concluded from these results that liver PFK was less suited for anaerobic energy production than muscle PFK. Rabbit muscle and erythrocyte PFK are different in their reaction toward muscle PFK antibody (19). Erythrocyte P F K was less sensitive to inhibition by the antibody and required much higher concentrations of antibody for complete precipitation than did muscle PFK. The essential concept of isoeneymes is that they are produced by the differential arrangement of dissimilar subunits. The protomers of acid dissociated muscle and liver P F K recombine a t neutral pH to form three electrophoretically distinct isoeneymes in addition to the two parent enzymes (92, 93). This result is consistent with the concept that muscle and liver PFK’s are different proteins but that both are tetramers. This would lead to the formation of three hybrids, namely, M3L, L3M, and M,Lz. There is evidence to show that multiple isoenzymes can exist in a single tissue (93-96). 89. S. Tarui, N. Kono, T. Nasu, and M. Nishikawa, BBRC 34, 77 (1969). 90. R. B. Layzer and M. M. Conway, BBRC 40, 1259 (1970). 91. C. B. Taylor and M. Bew, BJ 119, 797 (1970). 92. R. G. Kemp and M. Y. Tsai, Fed. Proc., Fed. Amer. SOC. Exp. Biol. 31, 499 (1972). 93. M. Y . Tsai and R. G . Kemp, ABB 150, 407 (1972). 94. 0. H. Lowry and J. V. Passoneau, Naunyn-Schmiedebergs Arch. Exp. Pathol. Pharmakol. 248, 185 (1964). 95. W. M. Poon and T. Wood, BJ 110, 792 (1968). 96. K. P. Maier, BBA 250, 75 (1971).

7.

259

PHOSPHOFRUCTOKINASE

G. REVERSIBLE INACTIVATION OF PFK

BY

DILUTION

Exposing PFK to mildly acidic conditions causes a loss of enzymic activity which is accompanied by a fall in molecular weight to about half of the value of the fully active form (8, 12, 13, 97-100).This dissociation is reversible. The property that dissociation is accompanied by a loss of activity has been used extensively to investigate the conditions required for dissociation and aggregation of the enzyme. Phosphofructokinase from rabbit muscle, (8, loo), heart (98),and Brevibacterium Ziqwjasciens (101) are all inactivated by dilution. At p H 8, muscle enzyme is relatively insensitive to inactivation by dilution, even a t protein eoncentrations of 0.1 pg/ml (8). As the p H is lowered the concentration a t which the enzyme becomes inactivated increases progressively. The rate of loss of activity is relatively rapid with a half-time of the order of minutes. Enzymic activity can be partially restored by making the p H more alkaline (i.e., pH 8 ) . The rate of reactivation is relatively slow and can be accelerated by the addition of components of the catalytic reaction. Reactivation of sheep heart P F K is accelerated by ATP, ADP, F6P, FDP, and cyclic AMP (98). In the case of P F K from B. Ziquejasciens (IOI),only MgATP will prevent inactivation by dilution. This effect is enhanced by the presence of NaF. The function of ATP in reactivating dissociated and other inactive forms of P F K is of great interest. Under some circumstances either ATP or FDP prevents the decrease in both molecular weight and enzymic activity brought about by decreasing the pH to 6.7 (8). The reactivation process wets investigated in greater detail by Alpers et al. (102) who found that 0.3 mM ATP causes 67% reactivation in 1 min and complete reactivation at 10 min. The reactivation rate becomes maximum a t 0.5 mM ATP. The action of ATP appears to involve an initiation process since, after exposing the enzyme to ATP for 15 sec, the ATP can be diluted out and activation still continues at the rate that would be expected with the original ATP concentration. If, after exposing to 97. M. F. Utter, Fed. Proc., Fed. Amer. SOC. E z p . Biol. 6, 299 (1947). 98. T. E. Mansour, JBC 240, 2165 (1965). 99. H. W. Hofer and D. Pette, Hoppe-Seyler’s 2. Phy&l. Chem. 349, 1105 (1968). 100. C. Frieden. “The Regulation of Enzyme Activity and Allosteric Interactions,” p. 59. Univemite Forlaget, Oslo, 1968. 101. M. Ide, M. Tokushige, and 0. Hayaishi, ABB 146, 361 (1971). 102. J. €3. Alpers, H. Paulus, and G. A. Bazylewicz, Proc. Nut. Acud. Sci. U.S.68, 2937 (1971).

260

DAVID P. BLOXHAM AND HENRY

A.

LARDT

pH 6, the enzyme was returned to pH 8 and allowed to stand for 1 min before ATP was added, only 30% of enzymic activity was restored compared to that obtained when ATP was added a t the same time as the pH was readjusted, suggesting that the enzyme can misfold a t pH 8 unless ATP is present. These results with the “acid dissociated” enzyme contrast with those observed with enzyme that has not been exposed to low pH. Parmeggiani et al. (7) found that a t all concentrations of muscle PFK, ATP caused a marked lowering of the sedimentation coefficient. For sheep heart P F K , ATP favored the formation of the dissociated form of the enzyme (7.5 S) whereas FDP favored the aggregated form (14.5 S) ( 1 3 ) . The effect of ATP in dissociating PFK may be related to its regulatory properties since Uyeda (64) has shown that skeletal muscle P F K has an s20,w of 13.1 S in the presence of MgITP (possesses only catalytic ac, in~ the presence of MgATP. tivity) but has an s ~ of ~8.1 S

H. PHOSPHORYLATION OF PFK? A number of enzymes involved in glucose metabolism such as phosphorylase, glycogen synthetase, and pyruvate dehydrogenase are regulated by a phosphorylation-dephosphorylation mechanism ( 3 1 ) .At present there is no conclusive evidence that such a mechanism applies to PFK in general; however, the properties of yeast and liver fluke P F K are reminiscent of the behavior of proteins that are regulated by phosphorylation. From both sources it is possible to isolate the enzyme in two forms. For liver fluke (29, 103, l o g ) , one form of the enzyme is active, whereas the other is inactive. The inactive form of the enzyme can be activated by incubation with a particulate enzyme fraction, ATP, Mg2+,and cyclic AMP. This activation has an absolute requirement for cyclic AMP. Physical properties of the two forms of the enzyme show that the in, ~whereas the active form has an s20,w active form has an s ~ of ~ 5.5S, of 12.8s. The incubation procedure for activation of the enzyme results in the formation of a thermostable fraction that can activate the enzyme in the presence of cyclic AMP. Analyses indicated the heat stable fraction contained considerable quantities of ATP, ADP, AMP, and Pi. A synthetic mixture containing these components as well as cyclic AMP and Mg‘f also activated the enzyme. The activation process can be 103. D. B. Stone and T. E. Mansour, Mol. Pharmacol. 3, 161 (1967). 104. D. B. Stone and T.E. Mansour, Mol. Phnrmrtcol. 3, 177 (1967).

7.

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261

reversed by dialysis which suggests that activation does not involve formation of a stable chemical bond. For yeast PFK, one form of the enzyme is sensitive to ATP inhibition (PFK,), whereas the other form is desensitized to ATP inhibition (PFKd) (106-107); PFK, can be converted to PFKd by incubation with a desensitizing protein, NaF, RSgATP, and cyclic AMP. The yeast desensitizing protein has been purified 50-fold (10'7). Furthermore, PFKd can be subjected to ammonium sulfate fractionation and still retain the PFKd characteristic which suggests a stable structural transformation (107).Afting et al. (108)have recently suggested that the desensitizing process may not involve a chemical transformation. They demonstrated that a mixture containing ADP, F6P, NH,', Mg2+, and NaF could stimulate tlie conversion of PFK, to P F K , in the absence of the desensitizing protein. The desensitizing protein may alter the composition of the nucleotides added to the incubation medium such that they prevent the enzyme from being inhibited by ATP. I n agreement with this proposal, the ultrafiltrate from the incubation of desensitizing enzyme with PFK and cofactors actually promotes the conversion of PFK, to PFKO. In conclusion, it appears that these phenomena represent changes in the relative concentrations of effectors rather than chemical modification of the enzyme. VI. Regulatory Properties of PFK

The major features of the regulation of P F K are as follows: 1. Inhibition of the enzyme by high concentrations of ATP. Generally, ATP decreases the affinity for the second substrate, F6P. 2. Inhibition by citrate in the presence of inhibitory concentrations of ATP. 3. Counteraction of inhibition by AMP, cyclic AMP, and FDP. 4. I n the presence of inhibitory concentrations of ATP, increasing the concentration of F6P has a cooperative effect on the activity of the enzyme. Thus, a plot of initial velocity versus F6P concentration is sigmoidal and Hill plots have slopes greater than unity. 105. E. Vinuela, M. 1,. Salas, M. Salas, and A. Sols, BBRC 15, 243 (1964). 106. J. M. Gancedo, W. Atzpodien, and H . Holzer, FEBS Lett. 5, 199 (1969). 107. W. Atzpodien, J. M. Gancedo, V. Hagmaier, and H. Holzer, Eur. J . Biochem. 12, 6 (1970). 108. E.-G. Afting, D. Ruppert, V. Hagmaier, and H. Holzer, ABB 143, 587 (1971).

TABLE I V

THEACTIONOF EFFECTORS ON PFK

Source

Shape of saturation curve for F6P

Muscle Brain Heart Liver Human erythrocyte Kidney cortex Adipose tissue Lens Sperm Jejunal mucosa Ascites tumor

Sigmoidal Sigmoidal Sigmoidal Sioidal Sigmoidal

Pea seed

Double hyperbola' Sigmoidal Sigmoidal

Carrots Brussels sprout leafh Corn

Hyperbolicb Sigmoidal Sigmoidal

FROM

VARIOUS SOURCES

Eff ectoro ATP

I I I I I I I I I I

Citrate

AMP

Mammalian I A I A I A I A I I A I A Weak1 WeakA I A

-

Ad

I*

Ie

A

10

I

Plant Weak I

I I I

I I I

Weak I I I

cyclic AMP

ADP

A A A A

A A A A

A

A A

A A Weak A A Ad

FDP

Pi

A A A

Reference

47, 57,109,110

4, 111 9, 10, 13, 2.9, 30, 36

15, 16, 36

A

A Weak A A Ad A

9 61

112 67 62

iis,114 115

None

I

A

116-119

None

I I I

A A A'

58 51 120

-

5 N . crassa A . crystallopoietes D. dkwideum C. pasteurianum F. themzophilum E. wli L. easei L. plantarum Yeast

Sigmoidal Hyperbolic Hyperbolic Sigmoidalb Hyperbolic’ Sigmoidalb Hyperbolic Hyperbolic Sigmoidal

I None None None None I None None I

Microorganisms Weak I None None None None None None Weak A None None None A None None I None None I I I A1 I None

I None Ij A

A A None I WeakA

IJ None None None - None A I None None

2 121 122 66 27 32,SS 24, 48, 54 71

71 52,53,12S

Abbreviations: I, inhibitor; and A, activator. Kinetics of enzyme are normal Michaeli-Menten with respect to F6P. This experiment was performed a t pH 8 and the inhibition by ATP was independent of pH or F6P concentration. c FDP does not reverse citrate inhibition of sperm PFK; however, it does activate the enzyme in the presence of ATP. d The maximum activation of this enzyme occurs when all three activators are present simultaneously. e Inhibition of enzyme by ATP and citrate shows a lag in onset of inhibition of about 2 min. J For this enzyme the curve becomes sigmoidal in the presence of inhibitory concentrations of PEP. 0 The most effective inhibitor of the enzyme is PEP. A plot of inhibition vs. PEP concentration is sigmoidal and has a slope greater than one in the Hill plot, suggesting a cooperative interaction of PEP with the enzyme. h Degree of regulation of activity appears to be greatest with PFK from young leaves. In the presence of low ATP levels, Pi is an inhibitor. Activation by Pi is observed at inhibitory ATP concentrations. jBoth of these products were competitive inhibitors with respect to their paired substrate, i.e., FDP with F6P and ADP with ATP. b The sigmoidal kinetics of this enzyme are not pH dependent. 1 Apparently only the enzyme from brewer’s yeast is activated by AMP. Baker’s yeast is not activated by AMP. a

6

t; in

%i

w

m

r:

c1

3 E

z

zB

264

DAVID P. BLOXHAM AND HENRY A. LARDY

The four features mentioned above represent the principal characteristics of P F K from many sources. Their influence on the catalytic activity of P F K is such that changes in concentrations of the effectors occurring under physiological conditions turn P F K on and off in accord with the physiological need for glycolysis. Other regulatory factors will be discussed in lesser detail. Table IV presents the regulatory features of P F K from a large number of sources and illustrates that these properties are variable (109-123). Many of the enzymes show sigmoidal rate curves for F 6 P ; however, the enzymes from lens (67),pea seed (116-119), Flavobacterium thermophilum (32, 3 S ) , Arthrobacter crystallopoietes (122), Dictyostelium discoideum (66), Lactobacillus casei (71), and Lactobacillus plantarum (71)give hyperbolic rate curves; the latter four enzymes are not inhibited by ATP. For P F K from pea seed and F . thermophilum cooperativity of F6P binding can be produced in the presence of the alternative inhibitor, PEP (32, 33, 116-118). Arthrobacter crystallopoietes (122) and D . discoideum (66) use intracellular protein or lipid as an energy source in preference to glucose. The fact that microorganisms, which do not use glucose as a primary energy source, possess unregulated forms of P F K emphasizes the requirement for P F K regulation in the control of glucose metabolism. The regulatory properties of PFK show certain phylogenetic traits. The most obvious of these is that ADP is an activator of mammalian enzymes, whereas it is an inhibitor for plant enzymes. There are numerous activators for mammalian PFK, whereas P, is the only phosphate compound capable of activating plant PFK. The regulated enzymes from microorganisms appear to show characteristics of both plant and mammalian enzymes. The inhibitory action of ATP was first reported for rabbit muscle 109. J. V. Passoneau and 0. H. Lowry, BBRC 7, 10 (1962). 110. A. Parmeggiani and R. H. Bowman, BBRC 12, 268 (1963). 111. J. V. Passoneau and 0. H. Lowry, Advan. Enzyme Regiil. 2, 265 (1964). 112. R. M. Denton and P. J. Randlc, BJ 100, 420 (1966). 113. W. Ho and J. W. Anderson, BBA 227, 354 (1971). 114. G. A. Tejwani and A. Ramaiah, BJ 125, 507 (1971). 115. R. W u , JBC 241, 4680 (1966). 116. G. J. Kelly and J. F. Turner, BBRC 30, 195 (1968). 117. G. J. Kelly and J. F. Turner, BJ 115, 481 (1969). 118. G. J. Kelly and J. F. Turner, BBA 208, 360 (1970). 119. G. J. Kelly and J. F. Turner, BBA 242, 559 (1971). 120. L. A. Garrard and T. E. Humphreys, Phytochemistry 7, 1949 (1968). 121. M. U. Tsao and T. I . Madley, BBA 258, 99 (1972). 122. J. Ferdinandus and J. B. Clark, BJ 113, 735 (1969). 123. M. L. Salas, E. Vinuela, M. Salas, and A. Sols, BBRC 19, 371 (1965).

7.

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265

PFK ( 6 7 ) . Inhibition by an excess of substrate is not unusual, but when that substrate is one whose concentration varies under different physiological conditions, the possibility of a functional regulatory mechanism becomes apparent. In the case of PFK the regulation is made more acute by the fact that metabolic degradation products of ATP specifically reverse the inhibition by ATP. Increasing the concentration of ATP above that necessary for the optimum catalytic activity causes a progressive decrease in P F K activity. The inhibitory action of ATP is reduced as the concentration of F6P is increased (13, $6, 104, 109). This illustrates that the ability to demonstrate the regulatory effects of either substrate is a function of the relative concentration of the other substrate. The pH of the medium is an important factor in demonstrating inhibition by ATP (18.4, 126). Uyeda and Racker (47) showed that for rabbit muscle PFK, ATP is a powerful inhibitor ( K , = 2.5 mM) a t pH 7.1, inhibits only a t very high concentrations ( > 6 mM) a t pH 7.6-8.5, and does not inhibit a t pH 9. The inhibition was far greater when the incubation was performed in imidazole buffer, pH 7.1, than in phosphate buffer, pH 7.1, reflecting the activating properties of Pi. The enzyme from brain ( 4 ) , heart ( I S ) , and diaphragm (124) shows similar properties. Yeast PFK shows the reverse phenomenon in that it is more sensitive to inhibition a t alkaline pH 8 (21, 126). Thus, as ATP concentration is increased the pH optimum shifts to lower pH values where ATP is noninhibitory ( 2 1 ) .The enzyme from E . coli (48) is also inhibited a t alkaline pH (pH 8.5). The structural requirements for inhibition by nucleoside triphosphates were discussed earlier (see Sections IV,B and IV,C) . The second major inhibitor of P F K is citrate, which satisfies the original concept of alloderic effectors in that it shows no structural resemblance to the substrates or products of the catalytic reaction. The inhibitory action of citrate was originally discovered by Garland e t al. (127) using rat heart PFK. The inhibition was highly specific since no inhibition was seen with cis-aconitate, L,-isocitrate, a-ketoglutarate, succinate, fumarate, malate, tricarballylic acid, CoASH, or acetyl-CoASH, This contrasts with rat brain P F K (128) where citrate (0.03 mM), cis-aconitate (0.1 mM) , isocitrate (0.2 mM) , malate (0.6 mM), succinate 124. M . Ui, BBA 124, 310 (1966). 125. B. Trivedi and W. H. Danforth, JBC 241, 4110 (1966). 126. G. Kopperschlager, R. Freyer, W. Diesel, and E. Hoffmann, FEBS Lett. 1, 137 (1968). 127. P. B. Garland, P. J. Randle, and E. A. Newsholme, Nature (London) 200, 169 (1963). 128. J. V. Passoneau and 0. H. Lowry, BBRC 13, 372 (1963).

266

DAVID P. BLOXHAM AND HENRY A. LARDY

(1.5 m u ) , and a-ketoglutarate (2.5 mM) were all inhibitors with the K I values shown in parentheses. Other enzymes show intermediate properties with regard to specificity. Corn (120) and yeast (123) P F K are inhibited by citrate and isocitrate but are not affected by a-ketoglutarate. The enzyme from kidney cortex is inhibited by both malate and succinate as well as citrate but is not affected by other citric acid cycle intermediates ( 6 1 ) .In all cases examined so far citrate is the most potent of the citric acid cycle intermediates in inhibiting PFK. The action of citrate shows a complex relationship to the ATP concentration depending upon the enzyme source. For rat heart PFK, Pogson and Randle (35) showed that the inhibitory action of citrate was the same a t either inhibitory or noninhibitory concentrations of ATP. I n both ATP concentration ranges, citrate caused an increase in the K , for F6P without altering V,,,,. For PFK from yeast ( I d s ) , rat liver ( 3 6 ) ,kidney cortex ( G I ) , primate sperm ( 6 2 ) ,rat brain (128), and corn scutellum (120), the inhibition by citrate produced a similar increase in the K , for F6P; however, the mechanism of inhibition is different since it is enhanced by increasing the concentration of ATP. Thus, for rat brain PFK activity measured a t 0.01 mM ATP, citrate is virtually without effect, but a t 0.12 mM ATP increasing the concentration of citrate produces almost complete inhibition (128). Other inhibitors may play a role in the control of the enzyme. The list of effective inhibitors includes P E P (15, 47, 32, 116-118, 129), ADP in plants (51, 59, 120), phosphocreatine ( l a g ) , 3-phosphoglycerate (15, 118, l a g ) , 2-phosphoglycerate (15, 118, l a g ) , 2,3-diphosphoglycerate (15, l a g ) , 6-phosphogluconate ( l a g ) , and DPNH (130). The inhibitory action of D P N H may be nonspecific since it also inhibits aldolase, which is one of the auxiliary enzymes used in the assay of enzymic activity (130). I n the presence of inhibitory concentrations of either ATP or citrate, it is generally possible to activate the enzyme by adding agents such as AMP, cyclic AMP, ADP (for mammalian enzymes), Pi, and FDP. Increasing concentration of activators generally causes a decrease in the K , for F6P and a decrease in the cooperativity of F6P binding to the enzyme (see Table IV for references). In summary, inhibitors shift the rate curve for F6P to the sigmoidal form whereas activators change the rate curve to the hyperbolic form. A true change in the curve from sigmoidal to hyperbolic must result in a decrease in the cooperativity of F6P binding. For PFK from numerous sources this 129. J. Krzanowski and F. M. Matschinsky, BBRC 34, 816 (1969). 130. E.A. Newsholme, P. H. Sugden, and L. H. Opie, BJ 119, 787 (1970).

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is the case (13, 48, 54,109). Yeast P F K presents an exception to this rule since activators and inhibitors have no influence on the cooperativity of F6P binding as determined from the slope of the Hill plot (131). The one remaining problem for discussion involves the underlying, mechanism responsible for the sigmoidal rate curve for F6P. Enzymes with sigmoidal rate curves can be of profound significance to metabolic control since, as pointed out by Koshland (IS%!),an enzyme showing hyperbolic kinetics requires a large change (81-fold) in concentration of substrate to increase the rate from 10 to 90% of full activity, whereas, for an enzyme showing sigmoidal kinetics, only a small change (3-6fold) is required. A number of theories have been advanced to explain this phenomenon including the symmetrical theory of Monod et al. (IS$) and the sequential mechanism of Koshland et al. (132, 134). We do not wish to discuss the relative merits of these theories but rather to draw attention to some of the factors that may be responsible for sigmoidal rate curves. The first aspect to be considered is the involvement of ATP in this phenomenon. It is apparent that five of the enzymes that are not inhibited by ATP all show hyperbolic rate curves for F6P (32, 33, 66, 71, 122). If PFK’s that are susceptible to inhibition by ATP are assayed a t a pH where ATP inhibition is negligible (Le., p H 8 for muscle and heart P F K ) , then the rate curve for F6P is hyperbolic. Kopperschlager et al. ( 1 2 6 ) ,using yeast PFK, found that over the concentration of ATP used, the interaction coefficient for F6P binding was constant a t about 2.25. However, if the noninhibitory substrate, ITP, was substituted for ATP the value of the interaction coefficient fell to one, showing the F 6 P binding was not cooperative. This result has been confirmed by Lindell and Stellwagen (21).This pattern did not hold for another noninhibitory nucleoside triphosphate, UTP, where the value of the interaction coefficient for F6P was two (126).The results with activators also tend to confirm that sigmoidal kinetics for F6P are the result of ATP binding at the inhibitory site since activators that prevent the inhibition by ATP generally lead to a decrease in the interaction coefficient of F6P binding. Certain experimental evidence, however, suggests that other factors may be involved in the production of sigmoidal rate curves. Thus, the enzyme from C. pasteurianum (27) shows a sigmoidal rate curve for F6P despite the fact that ATP does not inhibit the enzyme. Furthermore, as pointed out earlier, Atkinson e t al. (131) have shown that the CO131. D.E.Atkinson, J. A. Hathaway, and E. C. Smith, BBRC 18, 1 (1965). 132. D. E. Koshland, Advan. Enzyme Regul. 6, 291 (1968). 133. J. Monod, J. Wyman, and J. P. Changeux, J M B 12, 88 (1965). 134. D. E. Koshland, G . Nemethy, and D. Filmer, Biochemistry 5, 365 (1966).

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operativity of F6P binding to yeast PFK is unaffected by the presence of increasing concentrations of inhibitors or activators. Recent evidence has pointed to an influence of protein concentration in the cooperativity of substrate binding and has suggested that in some cases sigmoidal kinetics may represent an experimental artifact. In order to measure reaction rates by conventional spectroscopy i t is generally necessary to make high dilutions of the enzyme to bring the rate to a level which can be measured with the available cquipment. Phosphofructokinase is highly susceptible to inactivation by dilution, and this inactivation is reversed by components of the catalytic reaction (see Section V,G). It follows that when the enzyme is diluted in the absence of substrate, it may be inactivated. Addition of F6P to the enzyme might increase enzymic activity as a result of its effect in promoting reassociation of the enzyme. This phenomenon could again explain cooperativity of F6P binding since F6P should increase the number of active species of enzyme. It follows that if sigmoidal kinetics are the result of dilution, then increasing the protein concentration should decrease the sigmoidal nature of the rate curve for F6P. Ramaiah and Tejwani (135),using rabbit liver PFK, have shown that the rate curve was a function of the dilution of the enzyme. This concept has been extended by study of the reaction of rabbit muscle PFK with its antibody (136, 137).The formation of enzymically active precipitable complexes between specific antibodies and P F K is a function of enzyme concentration. The precipitating effect of antibodies is optimal a t high P F K concentrations; a t low concentrations the amount of antibody required to produce precipitation is increased. These results can be explained on the basis that P F K is an equilibrium mixture of active (associated) and inactive (dissociated) units and that the antibody reacts only with the associated form of the enzyme. The ability of the antibody to form a complex with the associated form of P F K may promote aggregation (activation) of the enzyme by removing the associated form from the equilibrium. The rate curve for PFK as a function of F6P concentration is hyperbolic in the presence of antibody in contrast to the sigmoidal rate curve in the absence of antibody. If the action of antibody is to promote aggregation of the enzyme a t dilute protein concentrations, F6P would not be able to exert any additional effect in this direction which would explain the loss of the cooperativity of F6P binding. 135. A. Ramaiah and G. Tejwani, BBRC 39, 1149 (1970). 136. M. Donnicke, H. W. Hofer, and D. Pette, FEBS Lett. 20, 184 (1972). 137. M. Donnicke, H. W. Hofer, and D. Pette, FEBS Lett. 20, 187 (1972).

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The dilution-inactivation theory to explain the sigmoidal kinetics of PFK is not an isolated ease. Thus, 5’-adenosinemonophosphate aminohydrolase, at pH 7, shows sigmoidal kinetics a t a protein concentration of 0.5 pg/ml, which become hyperbolic as the protein concentration is increased to 50 pg/ml (139, D-Lactate dehydrogenase from Aerobacter aerogenes also exhibits a similar phenomenon (139). The dilution-inactivation theory docs not provide an explanation of thc inhibitory action of ATP. If this theory were applied to the inhibitory phenomenon, ATP would be predicted to enhance the dissociation of the enzyme. This is not the ease since ATP has exactly the reverse effect and protects against inactivation by dilution (8, 98, 100-102).However, it must be pointed out that Hulme and Tipton (14O), using beef heart PFK, have shown that the inhibitory action of ATP is decreased as the concentration of P F K is increased. Possibly this indicates that the ability of ATP to bind to the inhibitory site is enhanced as the tendency to dissociate is increased. At present, no definitive explanation is available to rationalize the action of the various inhibitors and activators of PFK. Kinetic and binding studies in the presence of various effectors do indicate that there are multiple binding sites for ATP (4, 14, 29, 82, 141, 142). For heart PFK, i t is reported (14, 29) that there are two catalytic sites and two inhibitory sites per 100,000 daltons. For muscle PFK, there is one active site per 90,000 daltons and two additional ATP binding sites ( 8 2 ) .

VII. Role of Specific Groups in Enzymic Activity

A, THIOLGROUPS Engelhardt and Sakov (143) recognized a t an early stage that PFK could be inactivated by oxidation with agents such as ferricyanide, HzOz, I?, quinones, and alloxan. The activity of PFK from rabbit muscle is markedly increased by raising the ratio of reduced to oxidized glutathione (8).The regulatory role of thiol groups is also demonstrated by the fact that thc enzymic activity is decreased by treatment with 138. 139. 140. 141. 142. 143.

R. M. Hcmphill, C. I,. Zielkc, and C. H. Sueltcr, JBC 246, 7237 (1971). R. V. Sawula and I. Suzuki, BBRC 40, 1096 (1970). E. C. Hulme and K. F. Tipton, FEBS Lett. 12, 197 (1971). B. Setlow and T. E. Mansour, Biochemistry 11, 1478 (1972). D. Garfinkel, JBC 241, 286 (1906). V. A. Enpelhnrdt, and N. E. Sakov. Biokhimiyn 8, 9 (1943).

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DAVID P. BLOXHAM AND HENRY A. LARDY

thiol-reactive agents such as iodoacetic acid, iodoacetamide, and p mercuribenzoate (144-1 4.6’). Whether the thiol groups are directly involved in the catalytic reaction is open to doubt since treatment with agents such as iodoacetamide seldom leads to complete inactivation of the enzyme (144). One feature of this inactivation by thiol-reactive agents is that it is possible to protect the enzyme by incubation with appropriate substrates or products of the PFK reaction. This property has been used to investigate possible structural changes caused by binding of substrates, products or regulators. There are 16-18 thiol groups per 90,000 daltons for rabbit muscle PFK, and in denaturing solvents (6 M urea, 4 M guanidine, 3 mM sodium dodecyl sulfate, all of these groups reacted with 5,5’-dithiobis (2-nitrobenzoic acid) (DTNB) ( 1 4 , 147). Since there are 17-18 half-cystines per 90,000 daltons (7, 19, 83) disulfide bonds seem not to be part of the enzyme structure. A similar conclusion was reached for heart PFK (146). On the basis of the rate of reactivity of thiols with DTNB, i t is possible to classify five possible groups (144, 147, 148) as follows: 1. Thiols reactive virtually instantaneously with DTNB. Younathan e t al. ( 1 4 ) placed a value of 6 moles of thiol per 90,OOOg of PFK, whereas Kemp (147-150) fixed a value of one for this figure. This could be related to the fact that Younathan e t al. took extensive precautions

to maintain the enzyme in the fully reduced form whereas Kemp did not. This group is protected completely by MgATP. 2. Two thiols somewhat less reactive and protected by F6P and adenine nucleotides. 3. One thiol that reacts at 0.4 the rate of the second group. 4. Five thiols. The reactivity of thiols in this group and in group 3 are pH dependent and protected by FDP. 5. The remaining thiols which react only when the enzymes’ tertiary structure is destroyed. The loss of enzymic activity can be correlated with the progressive alkylation of the enzyme with an agent such as iodoacetamide (144). The reaction of about 6 thiols per 90,000 daltons causes a loss of 60% of E. S. Younathan, V. Paetkau, and H. A. Lardy, JBC 243, 1603 (1968). H. C. Froede, G. Geraci, and T. E. Mansour, JBC 243, 6021 (1968). H. W. Hofer, Hoppe-SeyZer’s 2. Physiol. Chem. 351, 649 (1970). R. G. Kemp and P. B. Forest, Biochemistry 7 , 2596 (1968). M. M. Mathias and R. G. Kemp, Biochemistry 11, 578 (1972). R. G. Kemp, Biochemistry 8, 3162 (1969). 150. R. G. Kemp, Biochemistry 8, 4490 (1969).

144. 145. 146. 147. 148. 149.

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the enzymic activity and further alkylation produces little additional change. Both MgATP and F6P protect the enzyme from inactivation by iodoacetamide ( 1 4 ) . The apparently unique reactivity of the thiol in class 1 has been studied extensively by Kemp and his co-workers (147, 14.9, 150).This thiol group has a reactivity 2 X lo4 times greater than the reactivity of the thiol groups in denatured PFK, suggesting that this thiol is activated by virtue of its location in the native enzyme. The reactivity of this group to DTNB is decreased by MgATP. Kemp (149) has suggested that the binding of MgATP to P F K places the enzyme in a new conformation in which this thiol is no longer exposed. Chapman et al. (151)have presented evidence that thiol modification with either DTNB or p-mercuribenzoate leads to a change only in catalytic activity of the enzyme and not in regulatory properties (i.e., still inhibited by ATP and activated by AMP). The major effect of reaction of DTNB with the class 1 thiol is to cause a decrease in V,,, indicating that this actiX-Ray vated thiol may be related to the active site of the enzyme (150). irradiation of PFK also caused a progressive loss of thiol groups and enzymic activity (162).The regulatory activity as measured by ATP inhibition and AMP activation was far less sensitive to radiation and could be abolished only by very high doses of radiation. This emphasizes the relationship of catalytic activity to enzyme thiol groups. Recently, Mathias and Kemp (148) have studied the reactivity of the class 1 thiol with [14C]flu~r~dinitr~benzene (FDNB) in an attempt to analyze the influence of ligand binding on PFK conformation; FDNB reacts much more slowly with the class 1 thiol than does DTNB, making it suitable for rate studies. Adenosine triphosphate partially protected the enzyme from FDNB, whereas MgATP gave complete protection; MgGTP, MgCTP, and MgITP all provided protection. I n contrast to the protecting effect of MgATP, F6P enhanced the reactivity of the thiol with FDNB. The concentration of F6P giving a half-maximal increase in reaction rate was 50 p*M. Cyclic AMP, AMP, and ADP also stimulated the reaction with FDNB. This result indicates that these different agents may produce different conformational changes a t the active site of the enzyme. Whether the thiol group is involved in MgATP binding was studied by investigating the rate of thiolysis of the aryl thioether bond. If the arylated enzyme is incubated with excess dithiothreitol, the aryl group is lost from the enzyme. Addition of 1 m M MgATP slows the rate of thiolysis. Since MgATP alters the reactivity of the thioether, it 151. A. Chapman, T. Sanner, and A. Pihl, Eur. J . Biochem. 7, 588 (1969). 152. A. Chapman, T. Sanner, and A. Pihl, BBA 178, 74 (1969).

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DAVID P . BLOXHAM A N D H E N R Y A. LARDY

cap be argued that the free thiol is not required for binding of klgATP but rather the formation of the complex between the enzyme and MgATP alters the conformation of the protein so that the thiol group is placed in a modified environment.

B. ROLEOF HISTIDINE

If heart P F K is exposed to white light in tlic prcscncc of nictliylenc blue the enzyme is oxidized to a new species which is no longer sensitive to inhibition by ATP a t pH 7 (14,29, 153). The Hill coefficient of the enzyme was 3.13 prior to oxidation and decreased to 1.1 after oxidation (153).Despite this change in the regulatory properties of the enzyme, the catalytic activity was not appreciably influenced when assayed a t pH 8.2. These changes were correlated with alterations in the binding properties of the enzyme (14,2 9 ) . At saturation, the normal enzyme binds 3.6 moles of ATP, 2 moles of citrate, and 1.75 moles of F6P (specific binding as opposed to nonspecific binding of 11.8 moles of F6P) per 100,OOOg of PFK. After photooxidation, the enzyme binds 2 moles of ATP, 2 moles of F6P, and no citrate per 100,000 g. From this it seems reasonable to deduce that there are two catalytic and two inhibitory sites per 100,000 daltons of enzyme. The loss of binding sites for ATP and citrate on photooxidation is consistent with the loss of the inhibitory properties of these two ligands. Photooxidation resulted in the loss of half of the cystcines, and of 3 histidincs per 100,000 daltons of enzyme (79). I n order to determine whether either histidine or cysteinc was specifically involved in the inhibitory phenomenon, the effect of ethoxyformic anhydride, which apparently reacts specifically with histidine, was studied (79). At pH 6.8, the cthoxyformylatcd (3.5 molcs/100,000 g) enzyme was no longer inhibited by ATP and had lost cooperativity of F6P binding. The enzyme also showed decreased sensitivity to citrate inhibition. Cyclic AMP or AMP did not activate the enzyme in accord with the loss of inhibition by ATP. These results seem to provide a solid basis for involving histidine in the regulatory binding site of PFK.

C. OTHERFUNCTIONAL GROUPS Phosphofructokinasc from both muscle and heart can be inactivated by treatment with agents that react with amino groups (maleic anhy153. C. E. Alilfors and

T.E. Mansour, JBC 244, 1247

(1969).

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dride, succinic anhydride, and pyridoxal phosphate) (164, 155). Unlike the inactivation by sulfhydryl reagents (ie., iodoacetamide) which produces a maximum inhibition of 70%, the inhibition by amino reactive agents is virtually complete. The formation of maleyl and succinyl derivatives of muscle P F K caused dissociation of the enzyme to a form with a molecular weight of 80,000 (154). Muscle PFK was more resistent to dissociation by pyridoxal phosphate but did dissociate whcn 8 moles of pyridoxal phosphate were bound per 100,OOOg. On dissociation a 7 s form of the enzyme (molecular weight ca. 140,000) was produced compared to the 80,000 molecular weight species formed with maleic anhydride or succinyl anhydride. Heart PFK that had reacted with pyridoxal phosphate was fairly resistant to dissociation a t 20" but dissociated to a 7 S species a t 0" (155). The tendency of maleic anhydride and succinic anhydride to disrupt the quaternary structure of P F K indicates that these agents could be inhibiting by a nonspecific mechanism. However, the inhibition by pyridoxal phosphate which has a lesser tendency to disrupt quaternary structure may indicate a role of amino groups in the catalytic process. Heart PF K is completely inactivated whcn only 4 moles of pyridoxal phosphate are bound in a Scliiff's base linkage to 100,OOOg of the enzyme (155). This inhibition is readily reversed by dialysis; however, if the Schiff's base linkage is stabilized by reduction with NaBH,, the inhibition of the eiizynx is irreversible. One interesting feature of the inhibition by amino-reactive agents is the protective action of various ligands ; MgATP was the best protecting agent for thiol groups (144, 147-150). However, in the case of the reaction of amino groups, it provided only poor protection. Fructose 6-phosphatc and FDP were the best protective agents indicating that amino groups could be related to the F6P binding site (164, 155). Chapman e t al. (151) have proposed a role for a tyrosine residue a t the inhibitory binding site of ATP. Reaction of muscle P F K with low concentrations of the tyrosine-selective reagent, N-acetylimidazole, at 0" produced only a minimal change in the catalytic properties of the enzyme whcn assayed a t pH 8. However, the enzyme showed a marked change in its regulatory properties which was reflected in a loss of inhibition by ATP and an increase in the affinity for F6P. When the alkylation reaction was pcrformcd in the presence of ATP, there was no change in the regulatory propertics of PFK. 154. K. Uycdn, Biochemistry 8, 2366 (1969). 155. B. Setlow nnd T.E. Mansour, BBA 258, 106 (1972).

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DAVID P. BLOXHAM AND HENRY A. LARDY

So far, the role of various functional groups in the action of PFK has been based on the actions of agents that react with a general class of chemical groups. The most successful studies of groups involved in the active site of enzymes has come from the use of alkylating agents that are related to the enzyme’s substrate. Recently, Brunswick and Cooperman (166) have described the synthesis of 0-2’- (ethyl 2-diazomalonyl) cyclic AMP. This is a photoaffinity label of P F K which can be covalently bound to the enzyme under appropriate conditions, diminishing thc ability of cyclic AMP to activate the enzyme. Derivatives of this type may be of great value in specifically labeling the active and regulatory sites of the enzyme. Another useful approach is the demonstration that limited tryptic digestion of yeast PFK results in a progressive loss of sensitivity to ATP inhibition (157). It would be interesting if this effect could be related to the loss of specific regions from the protein.

VIII. The Role of PFK in the Control of Glycolysis

The analysis of the contribution of P F K to the control of glycolysis is divided into three sections: Pasteur effect, control of pyridine nucleotidc oscillations, and hormonal regulation of glycolysis. The control of PFK activity is of great importance to these phenomena and it is possible to extrapolate the in vitro results with metabolic effectors to produce reasonable control mechanisms for P F K that are in accord with in vivo observations.

EFFECT A. THE PASTEUR Facultative organisms subjected to anaerobic conditions enhance their rate of glycolysis to compensate for the lack of energy production by oxidative phosphorylation. This is known as the Pasteur effect. Cori (158) originally proposed the possibility that the regulation of PFK act,ivity was involved in the control of glycolysis. For perfused heart 156. D. J. Brunswick and B. S. Cooperman, Proc. Not. Acnd. Sci. U . S. 68, 1801 (1971). 157. M. L.Salas, J. Salas, and A. Sols, BBRC 31, 461 (1968). 158. C. F. Cori, “Respiratory Enzymes,” p. 175. Univ. of Wisconsin Press, Mad-

ison, 1941.

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(169,160), brain (161), yeast (123,162), kidney cortex (163, 164), Novikoff hepatoma (163), and adenocarcinoma (163) there is clear-cut evidence for a metabolic crossover point a t the level of P F K since the onset of anaerobic conditions is associated with a decreased concentration of the intermediates prior to P F K (i.e., F6P, glucose 6-P, and intracellular glucose) and an increase in subsequent metabolites, particularly FDP. In all of these cells, with the exception of brain and yeast, these, changes in P F K activity can be related to nucleot.ide concentrations. Thus, in most cases under anaerobic conditions, there is an increase in the concentrations of AMP and ADP paralleled by a decrease in ATP concentration. Since AMP and A D P arc activators whereas ATP is an inhibitor of P F K activity in vitro, it seems reasonable to propose that the altered concentration of these effectors would cooperate to increase the activity of PFK. For brain the only activators that incrcasc significantly in anaerobiosis are Pi and FDP (161). A fall in phosphocreatine concentration was also detected which could be related to local changes in nucleotide concentration. I n short-term experiments phosphocreatine could maintain ATP levels by phosphorylation of ADP. However, as the phosphocreatine concentration diminished, the energy potential, [ATP]/[ADP] * [Pi], dccreascd progressively with time. Phosphocreatine is also a potent inhibitor of brain PFK, so the increase in glycolysis could be related to its disappearance (189). Yeast cells are a major exception to the regulation by adenine nucleotide concentrations since Lyncn et al. (162) have shown that the level of ATP remains essentially constant under aerobic and anaerobic conditions. It appears that the control of yeast P F K activity is mediated by citrate which is an inhibitor (123). Thus, in yeast the increase in P F K activity under anaerobiosis is associated with a decrease in citrate concentration. This could produce a de-inhibition of the enzyme t o explain the increase in fermentation. Tissues such as the rat jejunal mucosa do not show the Pasteur effect 159. E. A. Ncwsholmc and P. J. Randle, BJ 80, 655 (1961). 160. J. R. Williamson, JBC 241, 5026 (1966). 161. 0. H. Lowry, J. V. Passoneau, F. X. Hasselbergcr, and D. W. Schultz, JBC 239, 18 (1964). 162. F. Lynen, G. Hartmann, K. F. Netter, and A. Schuegraf, Regul. Cell Metab., Ciba Found. Sump., 1968 p. 256 (1959). 163. R . Wu, BBRC 14, 79 (1964). 164. A. H. Underwood and E. A . Newsholme, BJ 104, 300 (1967).

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despite the fact that PFK isolated from this tissue is regulated by adenine nucleotides. Tejwani and Ramaiah (114) suggested that the activity of PFK may be high under aerobic conditions as a result of a failure of ATP to inhibit the enzyme for the following reasons:

+

1. The “energy charge” (ATP + $$ ADP)/(AMP ADP + ATP), and presumably the energy potential, are relatively low in jejunal mucose . 2. The concentration of F6P is likely to be high since the jejunum is the major site of absorption of glucose and fructose. 3. Concentration of NH,’ may also be activating as a result of metabolism of nitrogenous compounds by intestinal flora.

B. CONTROL OF PYRIDINE NUCLEOTIDE OSCILLATIONS Oscillations of DPNH concentrations can be shown in intact cells and cell-free systems (references 165 and 166 provide a comprehensive reference source). These oscillations can be produced by the addition of several sugars, and their nature depends upon the conditions employed. A reciprocal oscillation of the concentrations of F6P and FDP as well as PEP and pyruvate can be shown to accompany the oscillation of DPNH, indicating that PFK and pyruvate kinase represent major control points. Phosphofructokinase plays a key role in the establishment of oscillations since F6P is the last sugar in the glycolytic pathway that can produce oscillations ; the addition of FDP increases glycolysis but does not produce oscillations. Phosphofructokinase and pyruvate kinase are closely related to the control of ATP levels, suggesting that adenine nucleotide control may be involved in the production of oscillations of DPNH concentrations. Studies of the oscillations of ATP and ADP concentrations show that ADP follows DPNH whereas ATP follows DPN. The oscillations of F6P and FDP are slightly out of phase (retarded) with the oscillations of ATP and ADP. These changes can be accounted for by the observed t30-90% change in PFK activity occurring during a single oscillatory cycle (166). By inhibiting PFK, the increasing concentration of ATP results in a decreased concentration of FDP and an increase of F6P. As fermentation slows, ATP concentrations decrease and ADP and F6P increase to the point where PFK is again activated. 165. B. Hess, A. Boiteux, and J. Kruger, Advan. Enzyme Regul. 7, 149 (1988). 166. B. Hess and A. Boiteux, Annu. Rev. Biochem. 40, 237 (1971).

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C. HORMONAL CONTROL OF GLYCOLYSIS I n animals there is a requirement for a continuous supply of glucose for erythrocytes and for cells of the nervous system. This requirement has necessitated the evolution of systems to regulate the utilization of glucose. Muscle represents a large proportion of the body weight; thus, the control of glucose utilization by muscle is of prime importance to the regulation of blood glucose. A metabolic crossover point at the level of PF K has been demonstrated in a variety of situations. Starvation, alloxan diabetes (equivalent to insulin withdrawal), or exposure to fatty acids leads to a decrease in glycolysis which is associated with a rise in glucose 6-P and F6P concentrations and a fall in FDP concentration. This effect can be demonstrated in the perfused heart (110, 159, 167, 168), skeletal muscle (169), and kidney cortex slices (164) and clearly indicates that the decrease in glycolysis is related to a decrease in PFK activity. In the intact animal, one of the most obvious changes resulting from an altered circulating insulin level is a change in plasma fatty acid concentrations which plays an important role in the control of glycolysis (168). The inhibition of glycolysis by fatty acids is not related to changes in adenine nucleotide concentrations but is mediated by effects on citrate concentration (110, 127, 164, 168). Thus, adding fatty acid to the perfused normal r a t heart elevated citrate concentrations (110, 168). Hearts from alloxan diabetic rats had a higher content of citrate than those from normal rats and insulin administration to alloxan diabetic rats resulted in a decrease in cardiac citrate concentration (168). It was proposed by Randle et al. (168) that the increase in citrate concentration is sufficient to result in an inhibition of PFK and a concomitant decrease in glycolysis whereas a fall in citrate concentration will produce the opposite effect. Studies with fluoroacetate show that citrate can produce feedback inhibition in vivo (170). Parmeggiani and Bowman (110) have shown that under anaerobic conditions the ability of octanoate to raise the level of citrate is reduced and the fatty acid has no influence on the rate of glycolysis. 167. E. A. Newsholme, P. J. Randle, and K . L. Manchester, Nature (London) 193, 270 (1962). 168. P. J. Randle, P. B. Garland, C. N. Hales, E. A. Newsholme, R . M. Denton, and C. I. Pogson, Recent Progr. Horm. Res. 22, 1 (1966). 169. D. M. Kipnis and C. F. Cori, JBC 235, 3070 (1960). 170. J. R . Williamson, E. A. Jones, and G . F. Azzone, BBRC 17, 696 (1964).

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Inhibition of P F K and the resultant accumulation of F6P and glucose 6-P in cells serve to decrease glucose utilization because hexokinase is inhibited by its product, glucose 6-P (171). One further factor is the possible role that cyclic AMP may play in the control of glycolysis by virtue of its action as an activator of P F K (4, 104, 172). Catecholamines stimulate glycolysis in muscle which is accompanied by an increase in tissue cyclic AMP levels (173).The stimulation of glycolysis is probably the result of the ability of cyclic AMP to stimulate glycogenolysis (activation of phosphorylase ; inhibition of glycogen synthetase) (31, 173). Cyclic AMP is no more effective than AMP in activating muscle PFK, and the concentrations of the cyclic nucleotide in muscle probably never reaches that required to overcome inhibition of PFK by ATP ( 4 , 94, 104, 178). Postulation of a direct effect of cyclic AMP to activate PFK has come from studies on adipose tissue. In this tissue, stimulation of glycolysis by epinephrine (174) is accompanied by a large increase in citrate levels (176) which is a condition generally associated with a decrease in glycolysis. The failure of citrate to inhibit glycolysis under these conditions is probably related to the rise in cyclic AMP levels. Thus, Denton and Randle (112) have shown that of all the nucleotide activators of PFK, cyclic AMP was the most potent in reversing the inhibition by citrate. On the basis of the concentrations of cyclic AMP in adipose tissue reported by Butcher et al. ( l 7 6 ) ,it seems possible that cyclic AMP could activate PFK.

171. R. K. Crane and A. Sols, “Methods in Enzymology,” Vol. 1, p. 277, 1955. 172. T. E. Mansour, Pharmacol. Rev. 18, 173 (1966). 173. G. A. Robison, R. W. Butcher, and E. W. Sutherland. “Cyclic AMP,” p. 159. Academic Press, New York, 1971. 174. J. P. Flatt and E. P. Ball, JBC 239, 675 (1964). 175. R. M. Denton, R. E. Yorke, and P. J. Randle, BJ 100, 407 (1966). 176. R . W. Butrher, R. J. Ho, H. C. Meng, and E. W. Sutherland, JBC 240, 4515 (1965).

Adeny late Kinase L . NODA I . Biological Aspects . . . . . A . Introduction and Distribution B . Genetics and Disease . . C. Function . . . . . . I1. Molecular Properties . . . . A . Preparat.ion and Purity . . B . Composition . . . . . C . Reactive Groups . . . . D . Physical Properties . . . I11. Catalytic Properties . . . . . A . Metal Requirement . . . B. NucleotideSpecificity . . . C . Assay . . . . . . D . Equilibrium Constants . . E . Mechanism . . . . .

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279 279 282 285 288 288 291 293 295 297 297 298 300 302 302

.

I Biological Aspects

A. INTRODUCTION AND DISTRIBUTION Elsewhere in this volume enzymes are treated with varying specificity of the type XTP + YMP F? X D P Y D P. I n this chapter. attention is focused on those enzymes acting on adenine nucleotides as substrate:

+

ATP

+ AMP

2ADP

(1)

The Enyzme Commission designation is ATP: AMP phosphotransferase (EC 2.7.4.3) with trivial name adenylate kinase . Although the common 279

280

L. NODA

trivial name myokinase has long been used to apply to the enzyme isolated from muscle, it does seem advantageous to use the one name adenylate kinase for all the enzymes catalyzing the same reaction, regardless of their source. Adenylate kinase from rabbit muscle was originally purified to a considerable extent by Colowick and Kalckar in 1943 ( 1 ) by taking advantage of its remarkable acid and heat stability. It was M. Johnson of the University of Wisconsin who first suggested “phosphate dismutation” as descriptive of the reaction catalyzed by the enzyme ( 2 ) . In 1948, the Russian investigator Kotel’nikova (3) identified the enzyme in liver and erythrocytes and suggested the name ADPphosphomutase. The name “ATP-AMP transphosphorylase” ( 4 ) has also been used, and the name “adenylate kinase” was suggested by Colowick (5). Several reviews that include this class of enzymes have appeared (5-1 1) . The enzyme is plentiful in tissues where the turnover of energy from adenine nucleotides is great, such as muscle and mitochondria. The enzyme from rabbit muscle was isolated and crystallized ( 4 ) , and physical properties (12) and kinetics were studied (13). Other purification procedures and properties of the enzyme from the same source have been reported (14, 15). I n 1960, Chiga and Plaut (16) purified the enzyme from the soluble portion of swine liver and studied its properties. The enzyme has been isolated from purified bovine mitochondria and extensively studied (17, 18). Adenylate kinase has been purified from rat liver and properties determined (19, 20). S. P. Colowick and H. M. Kalckar, JBC 148, 117 (1943). H. Kalckar, JBC 148, 127 (1943). A. V. Kotel’nikova, Dokl. Akad. Nauk SSSR 59, 527 (1948). L. Noda and S. A. Kuby, JBC 226, 541 (1957). S. P. Colowick, “The Enzymes,” 1st ed., Val. 2, Part 1, p. 114, 1951. 0. Hoffmann-Ostenhof, Advan. Enzymol. 14, 219 (1953). 7 . H. M. Kalckar and H. Klenow, Annu. Rev. Biochem. 23, 527 (1954). 8. B. Axclrod, Annu. Rev. Biochem. 24, 45 (1955). 9. 0. Hoffmann-Ostenhof, Annu. Rev. Biochem. 29, 73 (1960). 10. L. Noda, “The Enzymes,” 2nd ed., Val. 6, p. 139. 1962. 11. R. K. Crane, Compr. Biochem. 15, 200 (1964). 12. L. Noda and S. A. Kuby, JBC 226, 551 (1957). 13. L. Noda, JBC 232, 237 (1958). 14. 0. H. Callaghan, BJ 67, 651 (1957). 15. 0. H. Callaghan and G. Weber, BJ 73, 473 (1959). 16. M. Chiga and G. W. E. Plaut, JBC 235, 3260 (1960). 17. F. S. Markland and C. L. Wadkins, JBC 241, 4124 (1966). 18. F. S.Markland and C. L. Wadkins, JBC 241, 4136 (1966). 19. V. Sapico, G. Litwack, and W. E. Criss, BBA 258, 436 (1972). 20. W. E. Criss, V. Sapico, and G. Litwack, JBC 245, 6346 (1970). 1. 2. 3. 4. 5. 6.

8.

ADENYLATE KINASE

28 1

The ubiquity of adenylate kinase is shown by the variety of materials in which it is found. Adenylate kinase was shown to be present in nuclei isolated from thymus and also in nuclei from mammary adenocarcinoma cells (bl), in lemon leaves (2b), in sweet and sour lemons (23),in wheat (W4), and in red blood cells ( 2 6 ) . It was purified from cockroach muscle mitochondria (26). Adenylate kinase has been purified from bovine eye lens (27), and a baker’s yeast enzyme has been studied at length (28-30). In the slime mold, Physuruin polycephalum, Chin and Bernstein (31) studied the variation of adenine nucleotide pools with stage of growth and found that AMP concentration did not vary with ADP and ATP, possibly indicating that adenylate kinase was compartmentalized or otherwise not functioning a t all stages of growth. The enzyme has been purified from Escherichiu coli, and the activity in cell-free extracts of Thiobacillus denitrificuns was compared with the activity in E . wli (32). Spudich and Kornberg (33) purified adenylate kinase from spores and vegetative cells of Bacillus subtilis and found the enzyme from the two sources indistinguishable with respect to gel electrophoresis, DEAE-cellulose chromatography, and specificity toward substrates. In the skin of neonatal rats adenylate kinase is associated with particles ( 3 4 ) .Todd et al. reported the distribution of the enzyme in various postmortem human tissues ( 3 5 ) , and Thuma e t al. (36) crystallized the enzyme from human muscle. Adenylate kinase is distributed unequally in certain regions of the cell. While Markland and Wadkins (17’) accomplished a notable purification of the enzyme from liver by first isolating mitochondria, Brdiczka et ul. 21. L. A. Miller and A. Goldfeder, Exp. Cell Res. 23, 311 (1961). 22. D. Van Noort and A. Wallace, Plant Physiol. 36, 368 (1961). 23. A. M. Abou-zamzam and A. Wallace, J. Amer. SOC.Hort. Sci. 95, 199 (1970). 24. J.-L. Bomsel and A. Pradet. Physiol. Veg. 5, 223 (1967). 25. P. Cerletti and G. DeRitis, Clin. Chim. Actu 7, 402 (1962). 26. R. R. Mills and D. G. Cochran, Comp. Biochem. Physiol. 18, 37 (1966). 27. J. Klethi and P. Mandel, Bull. SOC.Chim. Biol. 50, 69 (1968). 28. C . 4 . Chiu, S. Su, and P. J. Russell, BBA 132, 361 (1967). 29. S. Su and P. J. Russell, BBA 132, 370 (1967). 30. S. Su and P. J . Russell, JBC 243, 3826 (1968). 31. B. Chin and I. A. Bernstein, J . Bacteriol. 96, 330 (1968). 32. B. D. Patterson, B. Taylor, T. J. Bowen, and F. C. Happold, Abstr. Int. Congr. Biochem., 6th, 1964 Vol. IV, p. 327 (1964). 33. J. A. Spudicli and A. Kornberg, J . Bncteriol. 98, 69 (1969). 34. T. Rosett, I. Matsuo, A . Bailey, D. Smith, T. McDonald, and K. Brown, BBA 222, 5 (1970). 35. J. K. Todd, J. L. Bell, and D. N. Baron, BJ 90, 7P (1964). 36. E. Thuma, R. H. Schirmer, and I. Schirmer, BBA 268, 81 (1972).

282

L. NODA

(37) demonstrated two years later in 1968 that the mitochondrial enzyme in rat liver is localized between the outer and inner membrane of mitochondria. By careful use of digitonin or by freezing, mitochondria were prepared with broken outer membrane but with the inner membrane and its contents intact. In such preparations the adenylate kinase was shown to be released while the mitochondrial matrix enzymes, glutamate dehydrogenase and p-hydroxy-CoA dehydrogenase, were not released. Criss (38) clarified some of the conflicting reports of cellular distribution of the enzyme by showing that, of the four isozymes from rat liver identified by electrophoresis, nuclei contained the minor isozyme I and a small amount of isozyme 111, the mitochondria contained isozyme I11 which was about 68% of the total cellular adenylate kinase activity ; the microsomal fraction contained very little enzymic activity and the cytosol contained most of isozyme I1 which represented only about 20% of the total cellular adenylate kinase activity. Rat liver isozymes I and IV were minor components and identified only on the basis of isoelectric focusing in which the low ionic strength favors protein-protein interactions. It seems to this writer that the existence of these as distinct isozymes should be held as tentative pending isolation and demonstration of other properties different from those of isozymes I1 and 111. Depending on the use of hypotonic or isotonic medium in fractionating cellular particles, workers have been led to believe that adenylate kinase was in the cytoplasm or in the mitochondria. I n yeast, Chiu et al. (28) found very little activity in microsomal and mitochondrial fractions, while 98% of the total cellular enzymic activity was found in the supernatant. They did not state the tonicity of the medium used in the extraction.

B. GENETICSAND DISEASE The presence of adenylate kinase in red blood cells makes the enzyme readily accessible for genetic analysis in human populations. The work of Fildes and Harris (39) showed that three distinct isozyme patterns of individuals are evident after starch gel electrophoresis of red cell lysates. Electrophoretically separated bands of enzyme were visualized by use of hexokinase, glucose-6-phosphate dehydrogenase, phenazine methosulfate, and thiazolyl blue. Three adenylate kinase phenotypes 37. D. Brdiczka, D. Pette, G . Erunner, and F. Miller, Ew. J. Biochem. 5, 294 (1968). 38. W. E. Criss, JBC 245, 6352 (1970). 39. R. A. Fildes and H. Harris, Nature (London) 209, 261 (1966).

8.

ADENYLATE KINASE

283

were identified and designated AK,, AK,-,, and AK, on the basis of electrophoretic gel patterns. In the British population 90% of the people had A K , isoayme pattern, 10% had AK,-,, while the AK, pattern was rare. Other rare human isoayme AK,-, and AK,-, (40) have been reported. The isoenayme AK,-, is common in old-world monkeys. Analysis of the inheritance patterns of the adenylate patterns of the adenylate kinase phenotypes by Fildes and Harris suggested that the adenylate kinase phenotypes result from autosomal alleles AK‘ and AK,; that is, the A K , phenotype results from the homoaygous (AK’, A K ‘ ) genotype, the AK,-, phenotype is the heteroaygote (AK’, A K , ) , while the rare AK, phenotype is the result of the (AKe,A K P )homoaygote. The study by Berg (41) of 96 Norwegian families with 455 children confirm the inheritance patterns suggested by Fildes and Harris in that there is no evidence for sex linkage and the evidence indicates that the AK’ and AK, alleles are on an autosomal locus. The red blood cells of the AK,-, phenotype were found to have 20-50% higher enzymic activity than the red cells of an individual with AK,-, phenotype (48, 43). The significance of the difference in enzymic activity between the two phenotypes is not known. Attempts have been made to correlate the distribution of the different adenylate kinase phenotypes with the movements of peoples (39, 44). Brock (&), in a study of adenylate kinase isozymes from different human tissues, concluded that AK, or AK,-, isozyme patterns identified in the red blood cells by Fildes and Harris correspond to the adenylate kinase isoayme pattern identified in adult and fetal muscle, liver, kidney, brain, spleen, lung, and leukocytes although the intensity of individual bands from the various tissues varied. On the other hand, on the basis of S H reactivity and antibody inhibition studies, Khoo and Russell (46) concluded from data on rabbit and human isoaymes from various tissues that there is a minimum of two sets of isozymes within an individual. The adenylate kinases from muscle, erythrocytes, and brain were similar and form one set of isoaymes while that from liver, kidney, spleen, 40. E. R . Giblett, “Genetic Markers in Human Blood,” p. 512. Blackwell, Oxford, 1969. 41. K. Berg, Hum. Hered. 19, 239 (1969). 42. A. W. Eriksson, J. Fellman, M. Kirjarinta, M.-R. Eskola, S. Singh, H.-G. Benkmann, H. N. Goedde, .4. E. Mourant, D. Tills, and W. Lehmann, Humangenetik 12, 123 (1971). 43. S. Rapley and H. Harris, Ann. Hum. Genet. 33, ,361 (1970). 44. S. Rapley, E. Robson, H. Harris, and S. Smith, Ann. Hum. Genet. 31, 237 (1968). 45. D. J. Brock, Biochem. Genet. 4, 617 (1970). 46. J. C. Khoo and P. J. Russell, BBA 268, 98 (1972).

284

L. NODA

and heart were found to be similar and form another set of isoaymes. The lack of agreement on the question of isoaymes from different tissues may result from the use of different criteria to identify the isoenaymes; also, in the former work, which was based on electrophoresis for identification, the results may be complicated by hemoglobin interactions, as the author pointed out. Attempts have been made to use the enzyme as a diagnostic tool for certain diseases without sufficient success to warrant general clinical application. Lehmann et al. (47) found that adenylate kinase showed faster elimination from heart muscle than creatine kinase and proposed its use for early diagnosis of myocardial infarction. But Kaffarnick and Klaus (48) tested for adenylate kinase in the serum of patients with acute hepatitis, liver metastases and myocardial infarction and found the test less sensitive than the usual clinical tests and of no diagnostic importance. Adenylate kinase activity was tested in normal mice and mice with genetically induced muscular dystrophy and no significant difference was found by Kaldor and Gitlin (49).One complicating factor is that red blood cells are relatively rich in adenylate kinase. Schirmer and Thuma (60) studied the relative reactivities of the -SH groups of adenylate kinase from human muscle and liver of normal and dystrophic (progressive muscular dystrophy, Duchenne type) individuals. Normal human muscle adenylate kinase was completely inhibited by Ellman’s reagent while adenylate kinase from normal liver was not inhibited. Muscle adenylate kinase from dystrophic individuals was inhibited only 40-5076. Thus, the adenylate kinase from dystrophic muscle with respect to inhibition by Ellman’s reagent appeared to become more like the liver isoayme. However, it was also found that in two specimens of dystrophic muscle in which more than 80% of the muscle fibers had been replaced by fat and connective tissue that the extractable adenylate kinase remaining was completely inactivated by Ellman’s reagent just like adenylate kinase from normal muscle. The authors were led to speculate that isozyme of the liver type is present in dystrophic muscle or that in some way the -SH groups of adenylate kinase of normal muscle was altered to resemble the -SH group of dystrophic muscle. Isolation and study of the enzymes from normal and dystrophic muscle as proposed may shed light on the disease process. 47. F. G. Lehmann, K. W. Schneidrr, and H. Menge, Enzymol. Biol. Clin. 6, 36 (1966). 48. H. Kaffarnick and D. Klaus, Artzneim.-Folach. 16, 361 (1962); Biol. Abstr. 45, 58662 (1964). 49. G. Kaldor and J. Gitlin, Proc. Soc. Ezp. BWZ. Med. 113, 802 (1963). 50. R . H. Schirmer and E. Thuma, BBA u18, 92 (1972).

8.

285

ADENYLATE KINASE

C. FUNCTION Whether in rapidly metabolizing cells like those of liver, or in cells of muscle that converts chemical energy into mechanical energy, the role of adenylate kinase is believed to be to facilitate the storage and use of the high energy of the adenine nucleotides. Analyses of tissues with respect to AMP, ADP, and ATP show a remarkable constancy under various conditions that is maintained by the activity of this adeninespecific “high-energy-phosphate” transferring enzyme. In systems utilizing ATP with formation of ADP, the adenylate kinase effectively makes available the energy residing in the P-phosphate group of the original ATP molecule as shown in the reverse direction of Eq. (1). A further significance of the role of adenylate kinase in such systems may be the removal of ADP which may be an inhibitor of the enzyme system. In the regeneration of ATP from AMP arising from the above action of adenylate kinase or formed by hydrolysis of the pyrophosphate bond in the energy utilizing step of the system, adenylate kinase makes AMP an active component of the energy exchange system. This may be uniquely important at least in oxidative phosphorylation if it be accepted as claimed by Colli and Pullman (51) that only ADP is the primary phosphoryl acceptor. Atkinson proposed the quantity, energy charge (52), defined by the [ADP] + [AMP]) as a measterm ( [ATP] + 0.5[ADP])/( [ATP] ure of chemical energy available from the adenine nucleotides of the system, Thus, if all the adenine nucleotide in the system were in the form of ATP, the energy charge would be one; if all in the form of AMP, energy charge would be zero. The quantity is useful in the study of dynamic energy systems where the availability of energy as well as the concentration of the individual adenine nucleotides may be important. In leaves of plants, Bomsel and Pradet (53) measured the amounts of adenine nucleotides and found the energy charge to be remarkably stable even under rapid turnover of adenine nucleotides induced by changing physical conditions. These observations were interpreted as a result of the effectiveness of adenylate kinase in maintaining equilibrium. In retrospect, it comes as no surprise that in a variety of enzyme systems the concentration of the individual adenine nucleotides may exert a metabolic control. For example, some enzymes of citric acid cycle and of glycolysis are inhibited by high ATP (energy charge ap-

+

51. W. Colli and M . E. Pullman, JBC 244, 135 (1969). 52. D. E. Atkinson, Biochemistry 7, 4030 (1968). 53. J.-L. Bomsel and A. Pradet. BBA 182, 230 (1968).

286

L. NODA

proaching one) or stimulated by low AMP (energy charge toward zero) or by either condition such as phosphofructokinase and pyruvate dehydrogenase (64, 56), pyruvate kinase (56) and NAD isocitric dehydrogenase and citrate synthetase (57'). Still other enzymes are stimulated by high ATP (energy charge approaching one) or inhibited by high AMP (energy charge approaching zero) or by either condition such as phosphoribosyl ATP synthetase and aspartokinase (58), fructose-1,6-diphosphatase (59) and the citrate cleavage enzyme (60). Thus, the energy charge of the adenylate system can provide the cell with metabolic control. It has been suggested that free magnesium ion concentration, which is greatly influenced by the relative amounts of AMP, ADP, and ATP, might serve as a feedback signal for control of energy metabolism. Rose (61) calculated the adenylate kinase equilibrium constant for a variety of conditions and correlated values with that measured in erythrocytes. Blair (62) measured the constant in vitro and calculated values for the various species in the adenine nucleotide system for a range of energy charge values. Qualitatively, at least in an isolated system, it is clear that as ATP is hydrolyzed to ADP or AMP the free magnesium ion concentration must rise since ADP and AMP form much weaker complexes with magnesium than does ATP. Thus, the variation in free magnesium ion or one of its complexes with adenine nucleotide might conceivably serve as means of regulating energy metabolism. Adenylate kinase could be an important factor in control by the above means by (1) its role in stabilizing the relative amounts of adenine nucleotide, (2) its ubiquitous distribution, (3) possible flux in and out of mitochondria (see below), and (4) cellular control of the total adenylate kinase activity. Adelman et al. (63) found that the activity of adenylate kinase in rat liver was influenced by diet as well as by hormones. On fasting 48 hr, the level of adenylate kinase increased from 135 to 374 units/g liver, and on refeeding a high glucose diet the enzymic activity in 16 hr 54. I,. C. Shen, L. Fall, G . M. Walton, and D. E. Atkinson, Biochemistry 7, 4041 (1968). 55. E. R. Schwartz, I,. 0. Old, and L. J. Reed, BBRC 31, 495 (1968). 56. P. Maeba and B. D. Sanwal, JBC 243, 448 (1968). Symp. 27, 60 (1968). 57. D. E. Atkinson, Biochem. SOC. 58. L. Klungsoyr, J. G . Hageman, L. Fall, and D. E. Atkinson, Biochemistry 7, 4035 (1968). 59. K. Sat0 and S. Tsuiki, BBA 199, 130 (1968). 60. D. E. Atkinson and G . M. Walton, JBC 242, 3239 (1967). 61. I. A. Rose, Proc. Natl. Acad. Sci. U . S. 61, 1@79 (1968). 62. J. McD. Blair, Eur. J . Biochem. 13, 384 (1970). 63. R . C. Adelman, C.-H. Lo, and S. Weinhouse, JBC 243, 2538 (1968).

8.

ADENYLATE KINASE

287

was found to be about 40 units/g. With fed alloxan diabetic rats the adenylate kinase activity was about a t the high level of fasting and was reduced to normal levels by administering insulin. The authors pointed out the similarity to changes in activities under the same conditions for phosphoenolpyruvate kinase, glucose-6-phosphatase, and fructose-1,6diphosphatase, and they also pointed out that these enzymes catalyze essential steps in gluconeogenesis. Adenylate kinase appears to be involved directly or indirectly in the control of gluconeogenesis. The amount of adenylate kinase in the liver of fetal rats was low and increased rapidly after birth to the age of 10-12 days ( 6 4 ) . This was observed for both liver adenylate kinase I1 and liver adenylate kinase 111. This increase in adenylate kinase in liver after birth was interpreted to parallel the change from dependence of the fetus on glucose obtained primarily across the placental membrane before birth to dependence on the enzymes of gluconeogenesis and amino acid metabolism, which in turn depend on the energy charge of the cell and thus on the activity of adenylate kinase. Twelve days after birth the concentrations of the two liver isozymes I1 and I11 are fairly constant except for the influence of diet and hormonal control on isozyme I11 noted above. In a related study, Hommes e t al. (65) reported that adenylate kinase in the cytosol and mitochondria of rat liver increased about 10-fold during the period 8 days before birth to adulthood. Kendrick-Jones and Perry (66) measured the activity of a number of enzymes of rabbit skeletal muscle during late fetal and early postnatal development. Adenylate kinase activity of rabbit leg muscle increased rapidly during the period 5-15 days after birth, while in diaphragm muscle the rapid increase to the adult enzymic activity level took place just prior to and a t about the time of birth. When young rabbits were encouraged to become active a t an earlier age than normal the muscle adenylate kinase activity correspondingly increased above the normal controls. The authors suggested that the physical activity pattern of the muscle is an important factor in determining the time a t which the activities of adenylate kinase and several enzymes tested increase rapidly to adult levels. As pointed out above, rat liver adenylate kinase I11 is localized in the space between the outer and inner membrane of mitochondria. Criss (67) has made the interesting observation that phosphate as well as alkaline pH promote flux for adenylate kinase to the outside of the 64. 65. 66. 67.

R. Filler and W. E. Criss, BJ 122, 553 (1971). F. A. Hommes, A. R. Richters, and A. Beere, BBA 230, 327 (1971). J . KendrickJones and S. V. Perry, BJ 103, 207 (1967). W. E. Criss, J . Biochem. (Tokyo) 70, 273 (1971).

288

L. NODA

rhitochondria and that, while nucleotides did not prevent, the addition of divalent metals did prevent the phosphate-induced release. The release of enzyme slightly preceded or accompanied mitochondrial swelling. The mitochondrial uptake of released adenylate kinase under conditions of initial, but not complete, swelling was reversible to an extent by the addition of ADP (1 mM), MgCl, (3 mM), and glutamate (2 mM). Like other findings of the release of a variety of enzymes from mitochondria under physiological conditions, the reported flux of adenylate kinase across the outer mitochondrial membrane is of great interest to the understanding of the intimate mechanisms of energy metabolism in the cell. In the energy metabolism of muscle, no primary and essential role of adenylate kinase in the contractile mechanism has been demonstrated although in preparations of myosin the last traces of adenylate kinase activity is very difficult to remove. Earlier data (68) appeared to indicate adenylate kinase as a relaxing factor for muscle fibers in &TO, but later work (69) showed that there were other protein factors involved in the relaxing mechanism. Adenylate kinase has not been directly implicated in the generally accepted sliding filament mechanism of muscle contraction. The importance of adenylate kinase in biological systems is in its involvement in the maintenance of equilibrium among the adenine nucleotides, thereby functioning in the maintenance of energy charge. Since adenylate kinase is localized in cell compartments, is itself subject to control, and effects adenine nucleotide concentrations which serve in turn to exert metabolic control, this enzyme is important to the energy economy of living systems.

II. Molecular Properties

A. PREPARATION AND PURITY The purification procedures for the various adenylate kinase from the wide variety of sources studied show considerable similarities in the steps used as shown in Table I. I n all cases listed, with the exception of bovine liver adenylate kinase prepared from isolated mitochondria, advantage is taken of the acid stability of the enzyme. The acid treatment was usually carried out by the addition of acid at ice bath tempera68. J . R. Bendall, Nature (London) 173, 518 (1954). 69. H. Kumagai, S. Ehashi, and F. Takeda, Noticre (London) 178, 166 (1955).

E

2

TABLE I PROCEDURES USED IN PURIFICATION OF ADENYLATE KINASES Purification

Source and Ref.

Units/mg

Rabbit muscle (70) Human muscle (36) Porcine muscle (71) Rat liver I1 (19) Rat liver I11 (20) Bovine liver mito. (17)

2200 1920 1810 60 1000 1062

130 X 100 X 923 X 136 X 1570 X 700 X

Yeast (28)

1900

1600 X autolyzate

ext. ext. ext. cytosol homog. mito. ext.

Cellulose Gel Acid Ammonium chroma- chromastep sulfate tography tography

+ + +

+ + +

PhosphoPhosphoPhospho-

+

+ +

DEAE, CMTEAE

+ +

+ +

+

+

Crystallization

+ + +

Other ZnAcz precipitation ZnAcz precipitation Electrofocusing Electrofocusing IRCSO; Ca-phosphate gel Acetone fractionation

290

L. NODA

tures to about pH 3 followed by raising the p H to neutrality to precipitate the denatured inert proteins. It may be that this acid step could be used to advantage in the preparation of the enzyme from liver mitochondria as well. Ammonium sulfate precipitation to concentrate the enzyme or for fractionation was used in all cases with the exception of rat liver adenylate kinases I1 and 111. With kinases I1 and I11 ultrafiltration was used as a technique for concentrating dilute solutions of the enzyme. It may be that the reported poor stability of liver enzyme I1 might be ameliorated if the total protein concentrations were kept high and if storage in higher salt concentrations (such as resulting from ammonium sulfate fractionations) were the general rule. Substituted cellulose column chromatography was used in all cases except in the preparation of the rat liver enzymes in which isoelectric focusing was used. Phosphocellulose was used preponderantly, and this might possibly be a reflection of the fact that adenylate kinase is an enzyme for substrates that contain phosphate groups. Gel chromatography was used to advantage in less than half of the preparations listed and perhaps could have been helpful in others if tried. I n the muscle preparations, zinc ion precipitation of activity a t alkaline pH was useful in reducing the volume after a step such as acid treatment in which protein concentration was greatly reduced, but purity was not appreciably increased by the zinc step. In the muscle preparations a more efficient procedure has been to drop the pH and/or to dilute the solution to attain absorption of enzyme on P-cellulose and then to follow with gradient elution. In general, adenylate kinases seem to be too unstable under conditions of organic solvent fractionation and the procedure has found limited application. The preparations listed in Table I were tested for homogeneity by two or more procedures involving sedimentation, electrophoresis, chromatography, or antibody techniques and found to be essentially pure (estimated to be 85% in the least pure of the preparations listed). The specific activities for enzyme preparations from muscle of various species and from yeast are about 2000 units/mg, while rat liver I11 and bovine liver mitochondria1 enzymes have specific activities in the range of 1000 units/mg. A second rat liver enzyme (rat liver adenylate kinase 11) is reported to have the drastically low value of 60 units/mg and also to be so unstable as to limit further purification. Since high protein purity was found for the liver adenylate kinase I1 preparation, a possible explanation for the very low specific activity may be minor alteration in the enzyme molecule during preparation so that activity is lost without affecting the criteria of protein purity by the techniques of sedimentation and dodecyl sulfate-urea gel electrophoresis. The preparative procedures for nearly all the adenylate kinases take

8.

ADENYLATE KINASE

291

advantage of the remarkable acid stability of the enzyme followed by salt fractionation, ion exchange chromatography (or electrophoresis), and in some cases, gel chromatography. The preparations listed in Table I (17,19, 90,28, 36, 70, 7 1 ) ,with the possible exception of liver adenylate kinase 11, are essentially pure and can be obtained in reasonable quantities.

B. COMPOSITION The adenylate kinases are globular, low molecular weight proteins composed of only the usual amino acids. There are no bound cofactors. Table I1 summarizes the amino acid compositions. Amino acids present in relatively low amounts are proline, methionine, tyrosine, phenylalanine, histidine, and half-cystine. Of the adenylate kinases analyzed, trytophan is reported to be absent with the exception of that isolated from liver. The bovine liver mitochondrial enzyme has 2 tryptophan residues ( l 7 ) , while in the report of the enzyme from rat liver (20) no mention is made in connection with amino acid analysis employing the usual hydrolysis by HC1 in which tryptophan is lost, whether this residue is present or not. It may be possible that tryptophan is a distinguishing amino acid present in adenylate kinases of the liver type (liver, kidney, spleen, and heart) but not in adenylate kinases of the muscle type (skeletal muscle, erythrocytes, and brain). Rat liver adenylate kinase I11 and bovine mitochondrial adenylate kinase resemble each other with respect to specific activity and amino acid composition. The total number of residues is 216 and 197, respectively. The reported apparent distinction of the rat liver adenylate kinase I11 to be observed to form polymers may be a consequence of differing concentrations at which sedimentation was measured. The conclusion seems warranted that rat liver adenylate kinase I11 and bovine mitochondrial adenylate kinase are the same enzyme from different sources. Histidine is present in low amounts as shown in Table I1 (17, 90, 36, 71, 79, 7 9 a ) , and in view of the demonstration (73) that a t least one residue is critically involved in the catalytic mechanism of the rabbit 70. I,. F. Kress, V. H. Bono, Jr., and L. Noda, JBC 241, 2293 (1966). 71. I. Schirmer, R. H. Schirmer, G. E. Schulz, and E. Thuma, FEBS Lett. 10. 333 (1970). 72. T. A. Mahowald, E. A. Noltmann, and S. A. Kuby, JBC 237, 1138 (1962). 72a. H. Schirmer et al., unpublished data (1970). 72b. L. Soda, unpublished data (1971). 72c. K . Ando and L. Noda, unpublished data (1972). 73. R. H. Srhirmer, I. Schirmer, and L. Noda, BBA 207, 165 (1970).

h3 (0

h3

TABLE I1 AMINO ACID COMPOSITION OF ADENYLATE KINASES

Amino acid Aspartic Threonine Serine Glutamic Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lysine Histidine Arginine Tryptophan Half-cystine Amide-ammonia Total residues a

HUmaIl (Ref. 36) 21,500 MW

P~ 21,400 MW

13 13 12 26 7 18 10 15 4 8 17

7 5 20 4 13 0 2 12 194

From Schirmer et aE. (72a) and Noda (72b).

13 14 11

25 6 19 8 16 6 8 18 7 5 20 2 11 0

2 191

Rabbit (Ref. 7.2) 21,300 MW 13 12 10 25 6 18 12 15 5

8 18 7 5 20 3 12 0 2 10 191

Squidb 27,500 MW

Rat liver I11 (Ref. 20) 23,000MW

Bovine liver (Ref. 17) 21,500 MW

23 14 13 32 10 19 16 20 6 10 24 7 11 24 3 8 0 8

21 14 9 24 8 16 23 15 4 15 21 6 7 17 3 9 4

20 13 15 17 13 5 9 20 5 7 15 4 10 2 4

248

216

197

18

9 11

r

* From Ando and Noda ( 7 2 ~ ) .

U b-

8.

ADENYLATE KINASE

293

myokinase, the variation from 2 to 4 residues among the various adenylate kinases is of interest. An even number of half-cystines has been found for all enzymes. In the muscle-type enzymes these exist apparently for the most part as free -SH whereas the enzymes isolated from rat and bovine liver are not sensitive to -SH reagents. This, together with the presence of tryptophan in the liver enzymes, may be distinguishing amino acid compositions that differentiate between the muscle and the liver types of adenylate kinases. The determination of amino acid sequence is being undertaken on several adenylate kinases (71, 7 4 ) .With the rabbit muscle adenylate kinase, Olson and Kuby reported in 1964 that the carboxyl terminal residues were -(Ala, Val)-Lys-Leu and that the carboxyl-leucine residue was not essential for activity (74). C. REACTIVE GROUPS I n their early paper on adenylate kinase (myokinase), Colowick and Kalckar (1) stated that the enzyme “probably contains free -SH groups which are readily oxidized.” The rabbit muscle adenylate kinase has two free titratable -SH groups and the addition of -SH compounds prior to or during the assay procedure generally gave increased activity. Mahowald et al. (76) found, on the basis of inhibition by -SH reagents under certain defined conditions, a one-to-one relationship between the two -SH groups and loss of activity; i.e., each -SH appeared to be independently essential for activity. Kress et al. (70) found that reaction of p-hydroxymercuribenzoate and several of its structural analogs with -SH groups result in the formation of adenylate kinase-mercurial complexes possessing activities ranging up to half of the initial activity. The K , of the adenylate kinase-p-hydroxymercuribenzoate complex does not differ significantly from that of intact adenylate kinase, indicating that the -SH groups are not involved in substrate binding. Thus, contrary to the inclination to interpret loss of enzymic activity on reaction with an -SH reagent as demonstrating direct participation of the -SH group in the reaction mechanism, caution is required. The -SH group may be involved in maintaining a conformation state or facilitating subtle conformation changes necessary to the catalytic process. Price (76a) observed that the rates of reaction of 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole with the two -SH groups of porcine muscle 74. 0. E. Olson and S. A. Kuhy, JBC 239, 460 (1964). 75. T. A. Mahowald, E. A. Noltmann, and S. A. Kuby, JBC 237, 1535 (1962). 75a. N. C. Price, Fed. Proc.. Fed. Amer. Soc. Exp. Biol. 31, 601 Abs. (1972).

294

L. NODA

adenylate kinase differed by a factor of about 35-fold and further that the presence of adenine nucleotides markedly decreased the rate of reaction of the one “fast” -SH while the reaction of the “slow” group was effected to a much lesser degree. Reaction of the fast -SH with the reagent resulted in complete inactivation. Spectrophotometrically, it was observed that in the intact but not denatured enzyme molecule the reagent group migrated to an -NH, group of lysine that must be suitably close to the fast -SH group of the native enzyme. If the interpretation be allowed that reaction with the bulky reagent by the fast S H of porcine adenylate kinase leads to inactivation by steric interference or slight conformational changes, Price’s observation can be interpreted to be in harmony with the proposal made on the basis of results with rabbit adenylate kinase that the -SH groups are not directly involved in binding of substrates or in the catalytic process itself. The view that -SH groups may serve a secondary rather than a primary role in the catalytic mechanism gains added support from the variability with respect to -SH groups of the adenylate kinases isolated from various sources. Adenylate kinases from human, porcine, and rabbit muscle have free -SH groups which are required for maximal activity. The adenylate kinase from bovine mitochondria which contains 4 half-cystine residues had no free -SH groups (17) but was inactivated by treatment with -S-S- reducing agents and could be subsequently activated by air-oxidation. The yeast adenylate kinase does not have free sulfhydryl groups. Assuming that the basic catalytic mechanism is the same for all adenylate kinases, and in view of the presence and absence of free -SH groups together with the divergent requirements for the intact sulfhydryl group, one must agree with the conclusion made above that -SH groups do not participate directly in the enzymic process. Two to three methionines of the five methionine groups of rabbit muscle adenylate kinase have been modified by reaction with iodoacetic acid (76) (the -SH groups previously blocked and later unblocked). The loss of activity was shown to result from formation of the carboxymethylsulfonium derivative of methionine, but a conclusion could not be drawn whether derivatization directly affected the catalytic mechanism or had modified enzyme conformation. The imidazole group of histidine is involved in the catalytic center of many enzymes and has been shown to be a t the active center of ribonuclease; it is postulated to be involved in the mechanism of a number of phosphate transferring enzymes (77). One or more of the 76. L. F. Kress and L. Noda, JBC 242, 658 (1967). 77. D. C. Watts and B. R. Rabin, BJ 85, 507 (1962).

8.

ADENYLATE KINASE

295

cationic groups postulated by Crane (11) to be on the enzyme and interacting with phosphate oxygen might be an imidazole group. Schirmer et al. (73) photooxidized rabbit adenylate kinase in the presence of methylene blue and found that the single rapidly photooxidized histidine group of three present was not essential for activity. Destruction by photooxidation of one of the two less reactive histidine groups resulted in loss of activity. They observed that adding adenosine to the photooxidation mixture resulted in more rapid loss of activity while the presence of any adenine nucleotide or of MgATP, each a t 5 mM, was equally effective in protecting against loss of activity. One possible interpretation of the results is that an imidazole group may be interacting with a phosphate group of the adenine nucleotide. Cohn et al. (77a) have reported NMR spectral observations made a t 220 MHz with porcine adenylate kinase. They found that the C-2 proton resonance of one of the histidine groups is shifted down field when ATP binds to the enzyme. This unequivocally and directly demonstrates the involvement of a histidine in the catalytic mechanism. They further observed that as Mn'+ is added to the enzyme-ATP complex, not only does the histidine C-2 proton peak but also a second adjacent peak broadens. They tentatively assigned the second peak to an €-amino group of lysine. While the presence of imidaaole at the active site of adenylate kinase may be accepted as demonstrated and a secondary role based on chemical evidence for -SH group(s) seems tentatively acceptable, any presumed involvement at the active site of c-NH, of lysine or S-atom(s) from methionine(s) at this time is speculative. Of the tools presently available, NMR and X-ray diffraction studies (71) permit direct probing of structure and function, and with the work of several laboratories underway we can expect elucidation of the chemical mechanism. The NMR studies in Dr. Mildred Cohn's laboratory are continuing, and from the Heidelberg laboratory of Dr. Ken Holmes the 6-A resolution for the structure of crystalline porcine adenylate kinase has been reported (78). D. PHYSICAL PROPERTIES

In Table I11 are listed the molecular weights of adenylate kinases. Most enzymes have a molecular weight near 21,500 with the excep77a. M. Cohn, J. S. Leigh, Jr., and G . H. Reed, Cold Spring Harbor S y m p . Quant. BioZ. 36, 533 (1972). 78. W. Kabsch, R . H. Schirmer, and G . E. Schulz, Proc. Znt. Congr. Biophys. 4th M. 2, 37 (Abs. of contributed papers) (1972).

TABLE I11 SOME PHYSIC.4L PROPERTIES O F h E N Y L . 4 T E

x Source Muscle Human Pig Rabbit Liver Bovine mitochondria Rat I1 Rat I11 Other Bovine eye lens Yeast

x

KINASES

PI

S20.W

D200.w

Partial specific vol.

flfo

A 0.1% 280 nm

9.95 10.25 10.0"

0.74 0.74 0.73

1.15 1.15 1.15

0.667b 0.53@ 0.53b

36

6.1

2.3a 2.3" 2.3d 2.49 3.02 1.23

10.3 5.7

0.73 0.65

1.11 1.3

0.856'

17

@j,fJoo

3.52

4.8

0.74

1.1

21,000 41 ,000"

2.96'

7.1i

MW 21,500 21,300 21,000 21,500 49,000 23,0000 46,000

7.5

1013 0

10'

Ref.

ri 12

19 19, .20

8.0

r9 2%

~

Not corrected to water. At 279 mm. c At 277 mm. At. 25". At 25" and 0.77% protein. Calculated from Table IV data in Ref. (17). 0 A t concentrations of about 3 mg/ml and higher dimers and trimers are found. Assume partial specific volume 0.74. Not corrected for 0.10 M K-phosphate pH 6.7 plus 0.10 M KCl. 0

r

8.

297

ADENYLATE KINASE

tion of yeast and the liver enzymes. Rat liver mitochondria1 enzyme 111 was found to be a monomer a t low protein concentrations (less than 3 mg/ml in phosphate buffer) and to exist as dimers and trimers a t higher concentrations of the protein. The monomer is active as demonstrated by the fact that assays were a t very dilute concentrations. The unstable and rarer rat liver enzyme I1 was found to have a molecular weight of 46,000-49,OoO under conditions in which rat liver enzyme I11 was found to exist largely as the dimer. Of the stable enzymes prepared in relatively large amounts, the yeast enzyme has the highest molecular weight, 41,000 (neglecting the dimer and trimer of liver enzyme I11 which have molecular weights of 46,000 and 68,000). The remaining five enzymes listed have molecular weights of about 21,500. It seems possible that 21 ,O00-23,000is the normal molecular weight of adenylate kinases and that like the liver enzymes the yeast enzyme might possibly be a dimer. This conjecture is somewhat supported by high substrate binding ratios calculated on the basis of a molecular weight of 41,000 [Table I11 in Su and Russell ( 2 9 ) l . Crystals of adenylate kinase have been prepared from ammonium sulfate solutions of enzyme purified from rabbit, pig, and human muscle. Some physicochemical properties, in addition to molecular weights, are collected in Table I11 (12, 17, 19, 20, 28, 36, 71, 7 9 ) . The isoelectric point of rabbit muscle enzyme is pH 6.1 while the liver enzymes have isoelectric points of about 7.5. The muscle enzymes, none of which is reported to contain tryptophan, have an absorbancy for a solution of 0.1% a t 280 nm of about 0.6, while the bovine liver enzyme has a value of 0.856 in which the higher value reflects the presence of the tryptophan.

111. Catalytic Properties

A. METALREQUIREMENT Adenylate kinase catalyzes the transfer of phosphate between MgATP and AMP and for the reverse reaction between MgADP and ADP as shown by the equation

+ AMP

+ ADP

(2) A bivalent metal is required for adenylate kinase activity. Lowenstein reported in 1958 (80) that nonenzymic transphosphorylation between MgATP

MgADP

79. J. Klethi, Ezp. Eye Res. 7, 449 (1968). 80. J. Lowenstein, BJ 70, 222 (1958).

298

L. NODA

ATP and inorganic phosphate were catalyzed by bivalent metals of which MnZ+,Ca2+, and CdZ+were the most effective. The pH optimum with MnC1, was about pH 9, and optimum ATP-to-metal ratio was found to be in the range of 0.6-1.0. For rabbit myokinase, the order of reactivity of the ions (81) has been found to be Mg2+2 Ca2+> Mn2+> Ba2+,and the earlier report of Noda (13) of the failure of Ca2+to meet the rabbit muscle adenylate kinase metal requirement was in error. For the enzyme from yeast (29) the order is Mgz+> Ca2+> MnZ+> Ba2+, and for the bovine liver adenylate kinase Mg2+> MnZ+> Ca’+, Coz+ (18). While there are individual differences it is seen that Mg2+ gives the highest activity followed by Ca2+and Mn2+.Barium and cobalt ion in some cases give a small fraction of the activity observed with the most active metal ion. As will be discussed later in more detail, the metal ion is combined with the nucleotide di- or triphosphate to form a complex. Kinetic, NMR, and binding studies establish that there are two substrate sites per enzyme active site-a site for binding a nucleotide monophosphate or a metal-free diphosphate and another site for binding metal-nucleotide triphosphate or a metal bound diphosphate. B. NUCLEOTIDE SPECIFICITY Some data concerning the specificity of the adenylate kinases are summarized in Table IV. The adenine nucleotides have lower K , values and are the natural substrates. Base substitutions as well as substitution of hydrogen for hydroxyl in the 2‘-ribose moiety lead in general to decreased activity. For the reaction [Eq. (2)] of rabbit muscle adenylate kinase studied in the presence of Mn2+the decreasing order of reactivity for various nucleotide triphosphates were ATP, 2’-dATP, CTP, GTP, UTP, and ITP ( 8 1 ) ; for the yeast enzyme with MgZ+the decreasing order observed (29) was ATP, 2’-dATP, GTP, and ITP. The K , values (13-15, 18-20, 26, 29, 30, 82, 83) (Table IV) for rat liver enzymes I1 and I11 (19) are similar, while the K , values for each nucleotide are considerably larger for the bovine liver mitochondria1 enzyme. The rat liver enzymes reportedly used only 5’-AMP as phosphate acceptor of a variety of purine and pyrimidine nucleotides tested. Some activity was observed when dAMP was substituted for AMP with some of the enzymes, but in general it appears that the specificity of the enzymic AMP site is much more rigorous than the nucleoside tri81. W. J. O’Sullivan and L. Noda, JBC 243, 1424 (1968). 82. W. J. Bowen and T. D. Kerwin, ABB 64, 278 (1956). 83. L. V. Eggleston and R. Hems, BJ 52, 156 (1952).

TABLE IV KINETICCONSTANTS FOR ADENYLATE KINASES Rabbit (Ref. 13)

ATP ADP AMP dATP dADP dAMP dGTP GTP ITP UTP CTP

Bovine liver mitochondria (Ref. 18)

KAM)

Po

g

Yeast (Refs. 29, SO)

Rst liver I11 (Refs. 19, 90)

Rat liver I1 (Ref. 19)

Cockroach mitochondria (Ref. 26)

0.33

0.3,a 0.08b

1.8

0.054

0.43

0.39

0.31

0.33 0.25

1.6a 0.5a

2.7

1.8

0.27 0.058 0.4 2.0 0.4

0.18 0.12 0.83

0.3 0.073 1.8

1.5 0.32

NSc

NS NS

0.04b

0.2P 0.5b 0.8b 0.4b

0.1 0.31

NS NS NS NS NS

NS NS

NS NS

Vmaxd

Reverse (form ATP) Forward (use ATP)

28,000

27,OW

29,600 25,300

0.69,g 0.44h

0.80

25,000

10,700 16,000

KWPJ

Callaghan and Weber (16). In the presence of Mnz+(81). c NS, not a substrate. d (moles substrate)(moles enzyme)-l(min)-l. Callaghan (14). Kapp = [ A T P ~ ~ I ~ [ A M P ~ ~ I I [ A D P ~ ~ I I - ~ . g At 0.001 M Mg"+ (82). h Callaghan and Weber (16) and Eggleston and Hems (83). a

b

'

0.236

0.44

8;4

El E z

i

300

L. NODA

phosphate site. Thus, it seems as a general rule for the adenylate kinases that while substitutions of the base moiety of nucleoside triphosphate may lead to some activity, substitutions of the base moiety of the nucleoside monophosphate leads to greatly decreased or no activity. This more acute specificity of the AMP site has been pointed out in private conversation in connection with NMR studies by M. Cohn and her associates. Su and Russell (29) observed with the yeast enzyme that the substitution of AMP by another purine mononucleotide caused a greater decrease in the rate of reactions than a corresponding substitution of another trinucleotide for ATP. Specifically, they observed that ITP could serve as a substrate in the presence of AMP but that I M P with ATP gave no reaction. Very recently, Secrist et al. (84) synthesized fluorescent analogs of the adenine nucleotides modified through the 6-amino group to give a third ring attached to the purine moiety ( 1,N6-ethenoadenosine nucleotide) . They found the ATP analog a substrate for adenylate kinase as well as hexokinase, phosphofructokinase, and pyruvate kinase. Further, with muscle adenylate kinase, the fluorescent AMP analog was not a substrate, and the authors concluded that, a t least with respect to the 6 position of the purine ring, the AMP site is more specific than the ATP site. The difference in specificity of the two substrate sites leads to the possibilities (1) that the two substrate sites are distinct and specific-one site for AMP or ADP and another site for MgATP or MgADP; or (2) that since the mechanism is random Bi-Bi, as discussed below, the binding of the first substrate determines the configurational specificity of the second site. The first alternative seems adequate to fit the data and because of its simplicity is far more attractive; i.e., the two substrate binding sites of adenylate kinase are distinct with the AMP (free-ADP) site requiring an intact 6-amino group on the purine ring.

C. ASSAY The most generally reliable assay, applicable under the widest possible variation of conditions are methods dependent upon measuring changes in the amounts of the adenine nucleotides brought about in a measured interval of time by adenylate kinase as determined after separation of the nucleotides by chromatographic procedures. The method has been used with Dowex-1 resin (IS), with electrophoretic techniques 84. J. A. Secrist, 111, J.

11, 3499 (1972).

R. Barrio, N. J. Leonard, and G. Weber, Biochemistry

8.

ADENYLATE KINASE

301

(851, thin-layer chromatography (86), and by chromatography on paper (87). Less tedious spectrophotometric methods for measuring the reaction catalyzed in the direction of formation of ATP have involved use of the enzymes, hexokinase with excess glucose and glucose-6-phosphate dehydrogenase together with TPN (88), or by coupling with the creatine kinase reaction. In the latter case excess creatine is added together with creatine kinase to yield creatine phosphate from the ATP formed by the adenylate kinase reaction. The amount of creatine phosphate is measured as inorganic phosphate by acid molybdate decomposition of the labile creatine phosphate ( 4 ) . A pH-stat assay (75) couples the reaction of adenylate kinase with ADP as substrate with the hexokinase reaction carried out at pH 8 in which one mole of hydrogen ion is released for every mole of ATP formed by adenylate kinase. The rate at which standardized alkali is added to maintain the pH is a measure of adenylate kinase activity. I n another assay, AMP is utilized in contrast to the use of ATP in the above assays. Adenylic acid deaminase is used to specifically deaminate the AMP formed leading to a decrease in absorption a t 265 nm ( 4 , 89). The method suffers from a relatively small change in absorption during the deamination and from the differences in optimum pH being near 7-8 for the kinase and rather sharply a t pH 5.9 for the deaminase. A more widely used coupled enzyme assay for measuring the adenylate kinase reaction in the direction of utilization of ATP uses pyruvate kinase with excess phosphoenolpyruvate together with lactate dehydrogenase and excess DPNH. The decrease in DPNH is measured by the decrease in absorbance a t 340 nm with time (90,91). The various assays have all been used in more than one laboratory, and depending on the desired ends, the instrumentation available, and the preferences of the researcher certain methods seem to have their adherents. Isotopic methods can, of course, be adapted to those assays that depend upon separation of the nucleotides and have been used in studies of initial rates as well as of isotope exchange rates a t equilibrium (18, SO, 9 2 ) . 85. 86. 87. 88. 89. 1955.

T. R. Sato, J. F. Thomson, and W. F. Danforth, Anal. Bwchem. 5, 542 (1963). K. Randerath, Nature (London) 194, 768 (1962). H. A . Krebs and R. Hems, BBA 12, 172 (1953). T . Bucher and G. Pfleiderer, “Mcthods in Enzymology,” Vol. 1, p. 435, 1955. G. Nikiforuk and S. P. Colowick, “Methods in Enzymology,” Vol. 2, p. 469,

90. I. T. Oliver. BJ 61, 116 (1955). 91. H. Adams, Biochem. 2. 335, 25 (1961). 92. D. G . Rhoads and J. M. Lowenstein, JBC 243, 3963 (1968).

302

L. NODA

D. EQUILIBRIUM CONSTANTS Apparent equilibrium constants Knp,,= [AMP] [ATPI/ [ADP] " for the yeast enzyme were determined in which each term on the right in the equation represented the total of that nucleotide in its various forms. At 0.032 M MgCL, 30" and unspecified nucleotide concentrations, Su and Russell observed with yeast enzymes (30) that both pH and MgS+ concentration affected Kapp.By Arrhenius plots the heat of the forward reaction was found by them to be 917 cal/mole. For the liver enzyme a t 30" and initial [Mg2+]: [ATP] ratio of 1 : l or a [l\lg*+]: [ATP] ratio of 1:2, Markland and Wadkins (18) found an average value of KaPp= 0.82 for the equilibrium condition approached from either direction. Eggleston and Hems (83) and Callaghan and Weher ( 1 5 ) reported a value of 0.44.For bound adenylate kinase of r a t liver mitochondria, Siekevitz and Potter (93) estimated an equilibrium constant of about 1. As would be expected on the basis of effect of pH and of metal ion concentration on the ionic species of substrate present, the apparent equilibrium constant varies with change in these parameters.

E. MECHANISM Adenylate kinase is a phosphoryl transferring enzyme in which one substrate is bound to a divalent metal. There arc two independent sites for binding substrate-one for MgATP or MgADP and the other site to bind AMP or ADP. The mechanism is random Bi-Bi as shown by Fig. 1, DE

ME

FIG.1. Reaction mechanism of adenylate kinase [from Rhoads and Lowenstein (92) with permission], where E represents enzyme and symbols to right of E signify

binding of chelated nucleotide; M, D,, Dz. and T represent AMP, ADP, MgADP, and MgATP, respectively. 93. P. Sickcvitz and V. R. Potter, JBC 200, 187 (1953).

8.

ADENYLATE KINASE

303

according to the presentation of Rhoads and Lowenstein ( 9 2 ) . Using rabbit muscle adenylate kinase and by measuring isotope exchange rates a t equilibrium, Rhoads and Lowenstein (92)concluded that ordered Bi-Bi and ping-pong Bi-Bi mechanisms are excluded. They found that for rabbit muscle adenylate kinase the mechanism is random Bi-Bi as shown in Fig. 1. With liver adenylate kinase, Markland and Wadkins (18) tentatively interpreted their isotope exchange studies as consistent with an ordered Bi-Bi mechanism, but there has been no confirmation. The yeast adenylate kinase according to Su and Russell (SO) was found to catalyze in agreement with the concept of a random Bi-Bi mechanism. While Su and Russell concluded that the rate limiting step with yeast adenylate kinase was the interconversion of the ternary complex (i,e., phosphate transfer and not the binding of substrates or dissociation of products), Rhoads and Lowenstein, using rabbit muscle adenylate kinase, concluded that the binding and dissociation are a t least involved in the determination of the overall rate. These observed differences for the two enzymes may possibly result from slight differences in catalytic and/ or binding forces arising from differences in structure such as -SH substitutions for -%% groups, pointed out elsewhere in this review, or from factors related to the observation that excess Mgz+ inhibits the forward reaction of muscle adenylate kinase but not that of yeast adenylate kinase. The metal ion is essential for phosphoryl transfer. Mildvan (94) has reviewed the role of metals in enzyme catalysis and dealt specifically with adenylate kinase. This enzyme, like creatine kinase, belongs to the class of phosphoryl transferring enzymes in which the substrate lies between enzyme and metal in contrast to some other enzymes like pyruvate kinase in which the metal is bound directly to the enzyme. The NMR data for adenylate kinase clearly establish that the metal is very rigidly held when the metal nucleotide is bound to the enzyme. Cohn et al. (77u)by the use of a spin-labeled iodoacetarnide derivative (N-l-oxy1-2.2.5.5-tetramethyl-3-pyrrolidinyl) covalently bound to the sulfhydryl of creatine kinase, were able by NMR studies to determine distances among the atoms of Mn-ATP bound to creatine kinase. They concluded that the metal-bound nucleotide on the enzyme favors the syn configuration bringing the phosphate groups close to the purine ring and the manganese interacts with the a- and P-phosphate groups (not with the y-phosphate even in a quaternary complex), and thus the metal does not interact at all with the phosphoryl group transferred. Since creatine kinase and adenylate kinase are in the same class of 94. A . S. Mildvan. “The Enzymes,” 3rd ed., Vol. 2, p. 445, 1970.

304

L. NODA

phosphoryl transferring enzymes, the similarities of the two enzymes with regard to the role of metal appear to justify the supposition that the binding and phosphoryl transfer mechanism of MgATP by adenylate kinase must be essentially the same as for creatine kinase. Further, the kinetic data for the several adenylate kinases showing a sharp optimal ADP: metal ratio indicates that the ADP accepting the phosphoryl group should not be bound by divalent metal. The free ADP (AMP) binding site of adenylate kinase shows greater specificity than the MgADP (MgATP) site for the purine ring. Difference spectra in the region 250-300 nm provide evidence that the purine rings seem to move into a hydrophobic region when substrate binds to enzyme. It had been observed in our laboratory that the difference spectra for ATP in 90% dioxane compared to aqueous solution a t pH 8 showed the same peak at about 280 nm and a trough a t about 250 nm that is observed when adenine nucleotides bind to rabbit adenylate kinase. It seems probable by the above evidence as well as by analogy to other enzymes that there is a hydrophobic cleft into which the purine rings fit in adenylate kinase. It seems worthwhile to point out other observations with the creatine enzyme system that are probably analogous to the adenylate kinase system. Reed and Cohn (96), from their electron paramagnetic resonance studies of the binding of Mn-ATP to creatine enzyme, found evidence of substitution of ligand groups from the enzyme into the manganese coordination sphere only when creatine was added in addition to the metal nucleotide. With the abortive complex, enzyme-MnADPcreatine, addition of anions such as nitrate and chloride produce further changes in the electron paramagnetic resonance spectrum. The anions appear to bind at the vacant phosphoryl site leading to a complex resembling the transition state. If the creatine kinase sulfhydryl groups are blocked with iodoacetamide the spectral changes observed with binding of the second substrate are missing. It seems quite reasonable to speculate that with adenylate kinase the phosphoryl transfer takes place by a transition state resembling the nitrate form of the abortive quaternary complex of creatine kinase. With the data presently available, it seems adequate to suppose that phosphoryl transfer in adenylate kinase occurs by an SN2 type of displacement in which the phosphoryl group being transferred is attacked by the oxygen atom of the second nucleotide which is bound to the enzyme. Considering the findings of Leigh (96) for creative kinase and assuming an analogous 95. G. H. Reed and M. Cohn, JBC 247, 3073 (1972). 96. J. S. Leigh, Jr., Ph.D. Thesis, University of Pennsylvania, Philadelphia, Pennsylvania, 1971.

8.

305

ADENYLATE KINASE

structure for adenylate kinase, we may presume the metal is bound only to the a-,p-phosphorous atoms of ATP or ADP, and this serves to fix the configuration of the one nucleotide in the syn form to bind at the one particular site of the enzyme. In the transition state the metal might further interact with the enzyme and oxygen atoms. Recently, a report by Hampton et al. (97’)has appeared showing that 8,5’-cycloadenosine 5’-phosphate (an analog of AMP with ring closure between the (3-5’ of ribose and the C-8 atom of the imidazole ring of adenine) is a good substrate for porcine adenylate kinase and several other enzymes for which AMP is a substrate. The authors estimated for adenylate kinase that one of the C-5’ epimers can participate in the catalyzed phosphate transfer more rapidly than AMP. They concluded that, in view of the rigidity of the AMP analog with the fixed relative positions of the fused rings, i t appears that the catalytic process can proceed efficiently without rotation a t the 9,l’ nor at the 4’3’ bonds and without conformational changes of the ribose moiety. In harmony with proton magnetic resonance measurements for AMP in solution, these results of Hampton and co-workers showed that for enzyme-bound AMP the phosphate group is closer to H-8 than to H-2. ACKNOWLEDGMENT

The help of David Garver in the preparation of this manuscript is gratefully acknowledged.

97. A. Hampton, P. J. Harper, and T. Sasaki, Biochemistry, 11, 4965 (1972).

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Nucleoside Diphasflhokinases R . E . PARKS. J R . I . Introduction .

.

R . P. AGARWAL

.

.

.

.

.

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.

A . Historical Development . . . . B. Distribution . . . . . . . I1. Molecular Properties . . . . . . . A . Occurrence of NDP Kinase Isozymes B . Purification Procedures . . . . C. Physical Properties . . . . . D . Phosphorylated Enzyme . . . . I11. Catalytic Properties . . . . . . . A . Reaction Catalyzed . . . . . B. Specificity . . . . . . . C . Methods of Assay . . . . . D . Kinetics and Catalytic Mechanism . E . Metal Requirements . . . . . F. Sulfhydryl Groups . . . . . G . Conformational Changes . . . . IV . Functions in the Cell . . . . . .

. . . . .

. . . . . . . . . . . . . . . . . . . .

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. . . . . . . . .

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307 307 309 313 313 314 315 315 320 320 320 321 326 329 330 331 331

.

1 Introduction

A . HISTORICAL DEVELOPMENT The term “nucleoside diphosphokinase” (NDP kinase. ATP :nucleoside diphosphate phosphotransferase. EC 2.7:4.6) ( 1 ) is used to designate a family of relatively unspecific enzymes that catalyze the transfer of 1 . Since NDP kinases are not specific for ATP. 11. more correct terminology would be nucleoside triphosphate :nucleoside diphosphate phosphotransferase . 307

R. E. PARKS, J R . AND R. P. AGARWAL

308

the terminal phosphate group of 5’-triphosphate nucleotides to 5’-diphosphate nucleotides (2) by the following general mechanism: NlTP

+ NpDP 2 NlDP + NzTP

(1)

where N, and N, are purine or pyrimidine ribo- or deoxyribonucleosides. All N D P kinases that have been examined to date function through the formation of enzyme-bound high-energy phosphate intermediates. The first N D P kinases clearly identified were reported independently in 1953 by Berg and Joklik for yeast (3) and by Krebs and Hems for pigeon breast muscle (4). Over the next few years, various workers demonstrated that the enzyme is ubiquitously distributed in nature and is found in animals, plants, and microorganisms (see Section 1,B). A notable achievement was the crystallization of N D P kinase from brewer’s yeast by Ratliff et al. ( 5 ) .Today, the only crystalline preparations of N D P kinase available are those from yeast, although several laboratories (5, 6) have purified N D P kinases from various sources to apparent homogeneity (6-10). The only precise studies of the subunit structure, physicochemical properties, amino acid end groups, and composition of an N D P kinase have been performed with a crystalline yeast preparation which has been found to be a homologous hexamer with a monomer of about 17,000 daltons (11, 12). A significant development in the understanding of the biochemical behavior of N D P kinases came with the discovery that a purified human erythrocytic enzyme follows a ping-pong reaction sequence and functions through the formation of a stable phosphate to enzyme bond (7, 13, 14). All N D P kinases studied subsequently have been shown to function by 2. Throughout this article it is assumed that actual substrates arc Mg-nucleotide complexes, e.g., Mg-ATP. 3. P. Brrg and W. K. Joklik, Nature (London) 172, 1008 (1953). 4. H.A. Krebs and R . Hems, BBA 12, 172 (1953). 5. R. I,. Ratliff, R. H. Weaver, H. A. Lardy, and S. A. Kuby, JBC 239, 301 (1964). 6. B. Edlund, I,. Rask, P. Olsson, 0. WBlinder, 6 . Zetterqvist, and L. Engstrom, EUT.J. Biochem. 9, 451 (1969). 7. N. Mourad and R. E. Parks, Jr., JBC 241, 271 (1966). 8. R. P. Agarwal and R. E. Parks, Jr., JBC 246, 2258 (1971). 9. J. Sedmak and R. Ramaley, JBC 246, 5365 (1971). 10. B. Edlund, Actu Chem. Scund. 25, 1370 (1971). 11. R. H.Yue, R. L. Rntliff, and S. A. Kuby, Biochemistsy 6, 2923 (1967). 12. R. Palmieri, R. Yue, H. Jacobs, L. Maland, L. Wu, and S. A. Kuby, Fed. Proc., Fed. Ames. Sac. Exp. Biol. 29, A914 (1970). 13. N. Mourad and R. E. Parks, Jr., BBRC 19, 312 (1965). 14. N. Mourad and R. E. Parks, Jr., JBC 241, 3838 (1966).

9.

309

NUCLEOSIDE DIPHOSPHOKINASES

a similar reaction mechanism (8, 9, 15-18). The amino acid phosphorylated, in most instances, has proved to be histidine with the phosphate either in the 1 or the 3 position (6, 9, 10, 19-23). These observations permitted formulation of the following reaction sequence (13): NiTP+E=N,DP+ NiDP+E-PP=NzTP+

E-P E

(2)

(3) A recent development of considerable importance to future studies of N D P kinases is the discovery of marked electrophoretic heterogeneity of the enzyme in a number of tissues (22, 24, 25) ; for example, in human erythrocytes six peaks of N D P kinase activity were demonstrated with isoelectric points ranging from 5.4 to 8.3. The enzymic behavior and molecular properties of these presumed isozymes differ so greatly that questions have been raised concerning the true physiological role of this enzyme (22, 24). For an excellent and detailed review of studies with N D P kinases prior to 1960, the reader is referred to the chapter by Robert H. Weaver in the second edition of “The Enzymes” (26).

B. DISTRIBUTION The ubiquitous distribution of N D P kinases is shown in Table I. (27-56). Since the various laboratories have employed different assay 15. M. G . Colomb, A. ChBruy, and P. V. Vignais, Biochemistiy 8, 1926 (1969). 16. E. Garces and W. W. Cleland, Biochemistiy 8, 633 (1969). 17. A. Goffeau, P. L. Pedersen, and A. L. Lehninger, JBC 242, 1845 (1967). 18. 0. Wilinder. 8. Zetterqvist, and L. Engstrom, JBC 244, 1060 (1969). 19. A. W. Norman. R. T. Wedding, and M. K. Black, BBRC 20, 703 (1965). 20. 0. Whlinder, JBC 243, 3947 (1968). 21. 0. Wilinder, JBC 244, 1065 (1969). 22. Y. C. Cheng, B. Robison, and R. E. Parks, Jr., Biochemistry 12, 5 (1973). 23. 0. Whlinder, Actn Chem. Scand. 23, 339 (1969). 24. Y. C. Chrng, R . P. Agarwal, and R . E. Parks. Jr., Biochemklry 10, 2139 (1971). 25. R. E. Parks, Jr., P. R. Brown, Y. C. Cheng, K. C. Agarwal, C. M. Kong, R. P. Agarwnl, and C. C. Parks, Comp. Biochem. Phgysiol. (1973) (in press). 26. R. H. Weaver, “The Enzymes,” 2nd ed., Vol. 6, p. 151, 1962. 27. H . P. Agnrwal, E. M. Scholar. K. C. Agarwal, and R. E. Parks, Jr., Biochem. Phrrrmacol. 20, 1341 (1971). 28. R . P. G1:izr anti C. L. Wadkins, JBC 242, 2139 (1967). 29. D. R. Sansdi, D. M. Gibson, P. Ayengar, and M. Jacob, JBC 218, 505 (1956). 30. H. Nakamura and Y. Sugino, JBC 241, 4917 (1966). 31. P. Berg and W. K. Joklik, JBC 210, 657 (1954).

310

R. E. PARKS, JR. AND R . P . AGARWAL

TABLE I DISTRIBUTION OF NUCLEOSIDE DIPHOSPHOKINASES Activity. (units*/g tissue)

Source

Specific activity4 (unitsb/mg protein)

Mammals Human Eryt>hrocytesc Bovine Liver Liver mitochondria Heart mitochondria Thymus Brain (acetone powder) Rabbit Muscle Red bone marrow Erythrocytes" Pig Kidney Heart Rat Liver Liver mitochondria-whole Inner membrane [low-speed pellet (2,000-12,000 x g 10 min)] Outer membrane [high-speed pellet (144,000 X g 1 hr)l High-speed supernatant (144,000 X g 1 hr) Liver nuclei Liver cytosomes and microbodies Liver endoplasmic reticulum Liver supernatant Kidney Heart Spleen Brain Lungs Erythrocytesc Intestinal mucosa Baboon Erythrocytesc

33.0 (7); 75.0 (8); --d; (20,24)76.0 (27) -d

0.05 (7); 0.11 (8, 27)

(20)

5 . 4 (28) -d (16,6 9 ) 14.5 (SO) -d (31)

0 . 6 (28) 0.225 (16) 0.12 (SO)

(3,31) (32) 70.0 (33)

0.1-0.13 (31)

-d

-d -d

(34, 36) (34,

10.0 (7); 62.0 (22) -d

0.11 (33)

(36-42); 4 . 3 (28)

0 . 0 2 (7); 0.43 (22); 0.017 (43) 0.001 (43)

0.075 (43)

0.078 (43) 7.44 (22) 3 . 1 (22) 0.0 (22) 47.0 (22) 27.0 (7); 6 2 . 0 (22) 25.0 (7); 68.0 (22) 30.0 (7); 63.0 (22) 17.0 (7); 64.0 (22) 30.0 (22) 30.0 (7); 47.0 (22) -d

0.10 (7); 0.38 (22) 0.08 (7); 0.87 (22) 0.08 (7); 0.16 (22) 0.15 (7); 0.84 (22) 0.24 (22) 0.05 (7); 0.31 (22)

(4)

68.0 (33)

0.10 (33)

9.

31 1

NUCLEOSIDE DIPHOSPHOKINASES

TABLE I (Continued) Activity" (unitsb/g tissue)

Source Monkey Erythrocytesc Dog Erythrocytesc Cat Erythrocytesc Spleen of leukemic mice Novikoff hepatoma Seal, Phoca vitulina Erythrocytesc

Specific activitya (unitsb/mg protein)

5 1 . 0 (33)

0.09 (33)

110.0 (33)

0.15 (33)

90.0 (33) Highly active (44)

0.17 (33)

(46) 25-45 (26)

-d

Birds Chicken Liver mitochondria Pigeon Breast muscle Erythrocytes' Avian myeloblastosis virus Hagfish, Myxine glutinosa Eryt hrocytesc Dogfish, Squalus amnthias Erythrocytesc Eel, Anguilla rostrata Erythrocytesc

2.03 (46) -d

(4)

6 . 5 (33) -d

0.01 (33)

(47) Fishes

19.0 (26)

2 5 . 0 (26) 4 0 . 0 (26)

Plants Sugar beet leaf Silver beet leaf Pea, Pisum sativuni Seed (dry flour) Root Shoot Wheat Seed Shoot Barley Shoot Broad bean Root Shoot Sugarcane Shoot Root Potato tuber

-d

(48)

-d

(49)

100.0 (10);-d (49) -d (49)

(49)

0.8 (10)

-d

-d

(49) (49)

-d

(49)

-d

(49) (49)

-d

-d

-d -d -d

(49) (49) (49)

(Continued)

3 12

R. E. PARKS, J R . AND R. P. AGARWAL

TABLE I (Continued) Activitya (unitsb/g tissue)

Source Impatiens holstii leaves Jerusalem artichoke, Helianthus tuberosus Mitochondria Brewer’s yeast, Saccharmyces calbergenis Baker’s yeast Bacillus subtilis Escherichia coli Uninfected Infected Mierowccus luteus Streptowccus pneumoniae Schistosomu munsoni

-d

(60)

-d

(19)

Microorganisms -d (3,3 1 ) ; 115.0‘ ( 6 )

Specific activity(unitsb/mg protein)

0.88 (31);0.86, ( 6 )

4 . 4 (6) -d (9,61)

0 . 5 (6) 0.21 (9)

(47, 68, 63,63a) (47,61-64) -d (47) -d (66) 450.0 (66)

3.3 (63)

-d -d

The italic numbers in parentheses are reference numbers. activities have been converted to a common international unit unless otherwise mentioned. One unit of enzymic activity is that amount of enzyme which catalyzes the reacttion of 1 pmole of nucleoside triphosphate and of nucleoside diphosphate per minute. No attempt has been made to standardize the conditions such as temperature and pH. Hence, strict comparisons are not possible and activities will differ from one report to another. Units/ml packed cells. Activity present but values not reported. Units/g dry yeast. Autolysate.

* Enzymic

6

32. H. Klenow and E . Lichtler, B B A 23, 6 (1957). 33. P. R. Brown, R. P. Agarwal, J. Cell, and R. E. Parks, Jr., Comp. Biochem. Physiol. 43B, 891 (1972). 34. D. M. Gibson, P. Ayengar, and D. R. Sanadi, BBA 21, 86 (1956). 35. F. E. Hossler and R. Rendi, BBRC 43, 530 (1971). 36. T. L. Chan, J. W. Greenawalt, and P. L. Pedersen, J. Cell Biol. 45, 291 (1970). 37. R. P. Glaze, Fed. Proc., Fed. Amer. Poc. Exp. Biol. 26, 863 (1967). 38. E. Herbert, V. R. Potter, and Y . Takagi, JBC 213, 923 (1955). 38a. E. S. Canellakis and R. Mantsavinos, BBA 27, 643 (1958). 38b. L. I. Hecht, V. R. Potter, and E. Herbert, BBA 15, 134 (1954). 38c. R. Mantsavinos and E. S. Canellakis, JBC 234, 628 (1959). 39. P. L. Pedersen and C. A. Schnaitman, JBC 244, 5065 (1969). 40. P. L. Pedersen and C. A. Schnaitman, in “Energy Transduction in Respiration and Photosynthesis” (E. Quagliariello el al., eds.), p. 831. Adriatira Editrire, Bari. 1971. 41. C. A. Schnaitman and P. L. Pedersen, BBRC 30, 428 (1968).

9.

NUCLEOSIDE DIPHOSPHOKINASES

313

conditions, it is not possible to make strict comparisons of the reported values. However, it is apparent that the activity of this enzyme is unusually high in many tissues. For example, in a study of the distribution of N D P kinases in various tissues of the rat, activities ranging from 30 to 68 units/g of tissue were determined (7, 2 2 ) . Also, the NDP kinase activity in the human erythrocyte, approximately 76 units/ml of packed cells, is one of the highest enzymic activities detected in this cell (27).

II. Molecular Properties

A. OCCURRENCE OF N D P KINASEISOZYMES

A number of observations suggested that isozymes of N D P kinase occur in various tissues. For example, during attempts a t purification of N D P kinases occasional aberrant peaks of activity were observed (6, 7, 28, SO). I n a detailed study with the N D P kinase of human erythrocytes, the technique of isoelectric focusing revealed the occurrence of a t least six enzymic peaks with isoelectric points ranging from PI 5.4 to 8.3 (Fig. 1). Studies of the kinetic parameters and physicochemical properties of these activity peaks suggest that they do not represent, classic isozymes but rather a family of related enzymes that function through high-energy phosphate intermediates; for example, 42. C. I,. Wadkins and A. L. Lehninger, Proc. N a t . Acad. Sci.

U. S. 46, 1576

( 1960).

43. C. A. Schnaitman and J. W. Greenawalt. J . Cell Biol. 38, 158 (1968). 44. P. A. Bianchi, M. V. Farina, and E. Polli, BBA 91, 323 (1964). 45. D. H. Ives, JBC 240, 819 (1965). 46. K. Kurahashi, R. J. Pennington, and M. F. Utter, JBC 226, 1059 (1957). 47. L. K. Miller and R. D. Wells, Proc. N a t . Acad. Sci. U. S. 68, 2298 (1971). 48. D. P. Burma and D. C. Mortimcr, ABB 62, 16 (1956). 49. R. J. A. Kirkland and J. F. Turner, BJ 72, 716 (1959). 50. N. C. Ganguli, JBC 232, 337 (1958). 51. J. Sedmak, N. Fernnld, and R. Ramaley, ABB 130, 488 (1969). 52. M. P. Argyrakis, BBA 166, 593 (1968). 53. 1,. J. Bello and M. J. Bessman, BBA 72, 647 (1963). 53a. I. R. Lehman, M. J. Bessman, E. S. Simms, and A. Kornberg, JBC 233, 163 (1958). 54'. R. Somerville and G. R. Greenberg, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 18, 327 (1959). 55. F. S. Markland and C. L. Wadkins, JBC 241, 4124 (1966). 56. 4. W. Senft, G. W. Crabtree, K. C. Agarwal, E. M. Scholar, R. P. Aganval, and R. E. Parks, Jr.. Biochem. Pharmacol. 22, 449 (1973).

314

R. E. PARKS, JR. AND R. P. AGARWAL

3.0

1.0

Ip '

I

I I

5.0

3

23

43 Fraction number (1.3 ml)

63

FIG.1. Electrofocusing profile of human erythrocytic nucleoside diphosphokinase. From Cheng et al. ($4).

marked differences in the Michaelis constants with various nucleotide substrates were observed. With four of the isoaymes, linear Arrhenius plots were seen, whereas biphasic plots were seen with two of the isozymes. Molecular weight studies demonstrated a range of about 80,000100,000 daltons which indicates that the phenomenon could not have been caused by aggregation (24). More recent studies have demonstrated that the heterogeneity of N D P kinase is not confined to the human but occurs in erythrocytes throughout the animal kingdom (22, 25). Furthermore, a study of the N D P kinases from a variety of tissues of the rat and in the subcellular fractions of rat liver revealed striking heterogeneity of the N D P kinases. However, each organ and subcellular fraction has its own unique N D P kinase isoayme pattern (22).

B. PURIFICATION PROCEDURES Purification procedures have been developed for N D P kinases from a large number of sources. Detailed procedures have been described for the isolation of crystalline N D P kinases from brewer's and baker's yeast (6, 6). Apparently homogeneous preparations have been obtained from calf liver mitochondria (28),human erythrocytes (7, 8 ) , pea seed ( l o ) ,

9.

NUCLEOSIDE DIPHOSPHOKINASES

315

and Bacillus subtilis (9). Partial purification has been achieved with N D P kinases from a variety of other sources: beef heart mitochondria (I6),calf thymus (%I), rabbit muscle (.!?I), and several plant sources (49). The recent discovery of the marked heterogeneity of N D P kingse from several sources (22, 24, 26) suggests that caution should be taken in the selection or development of a purification scheme. The technique of isoelectric focusing has proved especially useful in monitoring for the presence and distribution of isozymes at various stages of purification, procedures. As occurred in early studies with erythrocytic N D P kinase (7), the inclusion of a cellulose ion exchange procedure in purification may result in the loss of several isozymes (7, 8, 24). Attention is directed to the potential usefulness of calcium phosphate gel adsorption procedures in the purification of N D P kinases. Where tested, these methods have given virtually quantitative recoveries of enzyme with excellent purification and retention of all the isozymes (8, 22, 24, 27).

C. PHYSICAL PROPERTIES Several physical properties have been reported for preparations of homogeneous or crystalline N D P kinases as shown in Table I1 (57, 68). Most N D P kinase preparations studied to date have molecular weights in the range of about 80,000-110,000 daltons. It is of interest that a number of phosphate-binding studies indicate that 3-4 moles of phosphate bind per mole of enzyme, suggesting a tetrameric structure (6, 10, 1 6 ) . However, the structure of the crystalline yeast enzyme is that of a homologous hexamer with subunits of about 17,000 daltons ( I d ) . Excellent agreement was obtained between the molecular weight calculated from the amino acid composition and by physical methods ( 1 2 ) . The terminal carboxyl group of the brewer's yeast N D P kinase monomer is a glycine residue, and it was shown that no disulfide bonds were present in the enzyme (12). D. PHOSPHORTLATED ENZYME

As a consequence of the discovery that the erythrocytic N D P kinase follows a ping-pong reaction sequence, it was predicted that a phosphorylated enzyme intermediate was involved (IS). This prediction was verified by incubating [ y - V ] A T P with purified N D P kinase from human erythrocytes and by separating the phosphorylated enzyme on a 57. P. L. Pcdrrsen, JBC 243, 4305 (1963). 58. P. Andrews, BJ 91, 222 (1964).

SOME PHYSICAL PAR.4METERS O F

Enzyme source Human erythrocytes

Beef heart mitochondria Bovine liver mitochondria

Ref. s,24

15 57

Pea seed Brewer's yeast Bacillus subtilis Hagfish erythrocytes

9 25

Eel erythrocytes

25

Rat erythrocytes

22

Rat kidney

22

10 11

TABLE I1 NUCLEOSIDE DIPHOSPHOHINASES FROM VARIOUS

Isoelectric point Molecular (PI) weight

5.4 5.8 6.3 6.8 7.3 8.3 9.5 -

-

8.0 -

5.2;6.0; 6.7;7.3 5.0;5.8; 6.4;8.2 5.3;5.5; 5.65;5.8; 6.05;6.2 5.3;5.8; 6.05;6.2;

80,ooOn 93,w 84,W 80,OOOO 84,000= 100,W 103,000 100,000~ 109,00w 1o4,oooc 70,000 102,000 100,000 -

-

SOURCES

Diffusion Sedimentation Apparent specific coefficient coefficient partia: volume Frictional (dS+ x lo7) (sp,., x 10") Va*, ratio (cm' sec-1) (set) (cma g-l) (flfo) -

5.9 5 . 6 4 d +_ 0.15

5.40e 0.28

9.

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

I

NUCLEOSIDE DIPHOSPHOKINASES

I

317

TABLE I1 (Cmtinued)

Enzyme source

Ref.

Supernatant

Pig kidney

Diffusion Isoelectric coefficient point Molecular (c&.~x 107) (an2see-1) (PI) weight

5.30;5.50; 5.80;5.90; 6.05;6.20; 6.50 56

21,Ooof 92, OOOo

138,W By molecular sieving (68).

* From Sm,w value of 5.64.

From Sm,sp value of 5.40. By analytical ultracentrifuge. * By sucrose gradient. f Monomer. Tetramer. * Hexamer.

Sedimentation Apparent specific coefficient partial volume Frictional ( ~ x 10l '8), ~ VSPP ratio (set) (em3 g-1) (flfo)

9.

NUCLEOSIDE DIPHOSPHOKINASES

319

Sephadex column ( 7 , 13, 1 4 ) . Shortly thereafter these observations were confirmed with an NDP kinase isolated from artichokes (19). I n addition, it was demonstrated with this preparation that the high-energy phosphate residue is a phosphorylated histidine ( 1 9 ) .Since these original observations, every NDP kinase preparation that has been examined has been shown to function through a high-energy phosphate-enzyme intermediate (5, 9, 10, 16, 18, 2U, 21, 23, 36, 51, 57, 59). I n a series of investigations with various NDP kinase preparations, Wetlinder and his colleagues have identified in alkaline digests of phosphorylated NDP kinases l-phosphohistidine, 3-phosphohistidine7 and N-c-phospholysine (6, 10, 2U). With a NDP kinase purified from bovine liver, l-phosphohistidine appeared to be the major phosphorylated residue ( 2 3 ) . Similarly, with a homogeneous preparation isolated from B. subtilis, the principal phosphorylated residue was l-phosphohistidine (9, 5 1 ) . In preliminary studies with NDP kinase isozymes from human erythrocytes, both l-phosphohistidine and 3-phosphohistidine residues were identified (60). A possibility that must be considered in these studies is the migration of the phosphate residue from the l to the 3 position of histidine, which is the more stable form, a migration that might occur during the isolation procedure. With most NDP kinases that have been examined, the phosphorylated enzyme intermediates have proved to be alkali stable and acid labile, which is consistent with the formation of phosphate to nitrogen bonds as in phosphorylated histidines (16, 23). However, there have been reports of NDP kinases and of ATP-ADP exchange enzymes that function through alkali-labile and acid-stable phosphorylated enzyme intermediates (35,59). The acid stability of these phosphorylated enzymes suggests that they may function through carboxyl phosphate residues. The occurrence of apparent NDP kinases with drastically different stability characteristics and phosphorylated amino acid residues suggests that these NDP kinases may serve a function in the cell other than the transfer of phosphate groups among nucleotides. It has been shown with a number of NDP kinase preparations that more than one mole of phosphate binds per mole of enzyme; for example, with a pea seed NDP kinase, 3.1 moles of phosphate bound per mole of enzyme ( l o ) , and with baker’s yeast 3.5-3.6 moles of phosphate per mole of enzyme were determined ( 6 ) . With the human erythrocytic enzyme, approximately 2 moles of phosphate were bound per mole of enzyme ( 1 4 ) . Since physical studies have shown that the 59. M. G. Colomb, J. G . Laturaze, and P. V. Vignais, BBRC 24, 909 (1986). 60. Y. C. Cheng, B. Robison, and R . E. Parks, Jr., unpublished data.

320

R. E.

PARKS,

J R . AND

R.

P. AGARWAL

brewer's yeast enzyme is a homologous hexamer (Id),one should expect that 6 moles of phosphate should bind per mole of enzyme. However, Garces and Cleland have reported four phosphate-binding sites per mole of enzyme (16). In a series of studies with a rapid mixing technique, the rate of phosphorylation and dephosphorylation was determined with an N D P kinase isolated from bovine liver (18).Here it was shown that the turnover number for the phosphorylation reaction is 2,700 min-' and for dephosphorylation 24,000 min-l. The turnover number of the overall reaction is about 1,300 min-', which is strong evidence that the phosphoenzyme is the true intermediate for the N D P kinase reaction (18).It should be noted that the turnover number of crystalline brewer's yeast N D P kinase is very much greater, i.e., 600,000 min-' or about 150,000 min-' per active site (16).

111. Catalytic Properties

A. REACTION CATALYZED The reaction catalyzed by N D P kinases is the readily reversible transfer of the terminal phosphate of a 5'-triphosphate nucleotide to a 5'-diphosphate nucleotide as shown in Eq. ( 1 ) . A number of studies have reported that the equilibrium constant ( K e g )of this reaction is approximately 1.0 (9, 31, 4 9 ) . In a detailed study with the crystalline yeast enzyme, Garces and Cleland (16) found that the overall equilibrium constant a t pH 8.0 is 1.28. However, the partial equilibrium constant for the phosphorylation of free enzyme by ATP is 0.188, and for dephosphorylation of the phosphorylated enzyme by UDP it is 6.76 as shown in the following equations: E

-

E P

-

+ A T P e E P + ADP + U D P e E + UTP

(Keq= 0.188) (Keq= 6.76)

(4)

(5) These equilibria favor the free enzyme and suggest that the phosphorylated enzyme has a free energy of hydrolysis about 1.0 kcal more negative than the terminal phosphate bond of ATP (16).

B. SPECIFICITY All the N D P kinases that have been studied to date are relatively nonspecific and can react with di- and triphosphate ribonucleotides or

9.

NUCLEOSIDE DIPHOSPHOKINASES

321

deoxyribonucleotides that contain either purine or pyrimidine bases (3, 6, 7, 16, 24, 68, 50, 31, 49). In addition, a number of nucleotides that contain purine or pyrimidine analogs have substrate activity ( 7 ) . However, despite this relative lack of substrate specificity, one may not conclude that N D P kinases do not discriminate between the various nucleotides since widely different kinetic parameters have been’ determined (Table 111). Furthermore, the N D P kinase isozymes of human erythrocytes vary greatly in their Michaelis constants (Table 111) and V,,,,, values with different nucleotides. Although substrate specificities have been described for a number of enzyme preparations from a variety of tissues, little consideration has been given to the isozymic nature of N D P kinases, and it is possible that the substrate specificities described for a specific N D P kinase may, in fact, reflect the presence of mixtures of isozymes that have different reactivities with nucleotide substrates. I n view of the marked electrophoretic heterogeneity of N D P kinases in many tissues (22, 2 4 ) , the whole question of the nucleotide specificity of the various reported preparations should be reopened. It should be noted that monophosphate nucleotides are not capable of serving as substrates for N D P kinases although they may act as weak competitive inhibitors (7).

C. METHODS OF ASSAY Various procedures have been developed for the measurement of N D P kinase activity. They fall into three major groups: (1) coupled enzymic assays, (2) isotopic assays, and (3) staining procedures for localizing N D P kinase activity after chromatography or electrophoresis. 1. Coupled Enzymic Assays

Several coupled enzymic procedures have been developed in which the reaction rate is followed spectrophotometrically by measuring the appearance or disappearance of NADH or NADPH. These procedures have the advantage of being dynamic assays that enable continuous measurements, considerable sensitivity, rapidity, and ease of operation. In addition, they avoid the use of radioisotopes. Of the several methods available, the pyruvate kinase-lactate dehydrogenase and the hexokinase-glucose-6-phosphate dehydrogenase systems have been most popular. a. Pyrwate Kinase-Lactate Dehydrogenase Method. The pyruvate kinase-lactate dehydrogenase procedure (7) follows the formation of ADP from ATP or G D P from G T P in a coupled reaction system con-

w

E3 E3

TABLE I11 MICHAELIS CONSTANTS OF NUCLEOSIDE DIPHOSPEOKINASES FROM VARIOUS SOURCES~ Source of enzyme Human erythrocytes (Refs. 7, 8, 2.4)pH Nucleotides

5.4

ATP

0.b -

CTP

-

GTP

0.2'

ITP dTTP UTP

-

5.8*

6.3

6.8

7.3

8.3

0.12 3 . 0 ~ 0.25c 0.08c 0.1P 0.4- 0.lc 1.0c -

_

_

-

_

0.074; 0.10" 0.05c 0.16" 0.08' 0.lP _ _ _ _ _ _ _ 1.4 _ - _ -

Beef Calf Brewer's liver Cdf heart yeast B. subtilis mitochondria thymus mitochondria (Ref.16) (Ref.9) (Ref. 17) (Ref.30) (Ref.16) 0.31 -

0.15 0.22c 0.28; 0 .42c 0.059;

0.1W 0.25

0.30 0.11 0.13

Rat liver (Ref. 22) PH

m

6.05 6.50

1 .8-3. 5d; 2.0-8. OC

-

-

-

-

-

-

-

-

0.11 0.10

-

-

-

-

-

-

-

-

1.4~

*s

0.40" 0.12c

4

sd

-

_

-

-

-

0.5W 0.13"

sd cd

0.05-1.5d 0.01-0.17"

dCTP dGTP 5F-UTP ADP CDP GDP IJDP IDP dCDP dGDP dTDP 8-axaGDP 6-azaUDP

-

-

-

-

5.0 0.13 O . l l c 0.26; 0 .55c _ 0.1 3.3 -

All K , values are expressed in mM and are apparent values unless otherwise mentioned. The enzyme was mentioned as homogeneous NDP kinase (7) but was found later to be a mixture of PI 5.8 and 6.3 with PI 5.8 as a main component (8). c Corrected K , (apparent K,,, value increases as the second substrate approaches saturation concentration). In presence of a fixed amount of Mge+. e In presence of an excess of M@+. a

b

324

R. E. PARKS, J R . AND R. P. AGARWAL

taining phosphoenolypyruvate (PEP), NADH, pyruvate kinase, lactate dehydrogenase, and the nucleotide substrates plus the appropriate salts and buffers according to the following reactions:

+ +

+ + +

dGDP ATP c-) dGTP ADP ADP PEP ts ATP pyruvate Pyruvate NADH lactate NAD

(6) (7) + (8) The rate of decrease in absorbance a t 340 nm is followed spectrophotometrically. Although this procedure is very convenient, it has certain limitations. The diphosphate nucleotide substrate must be one that is a very poor substrate for pyruvate kinase, and the triphosphate nucleotide substrate must liberate a disphosphate nucleotide that is a good substrate for pyruvate kinase. This restricts somewhat the nucleotide combinations that may be studied. Diphosphate nucleotides such as dGDP, 8-azaGDP, dTDP, dCDP, and dADP fulfill these conditions (7, 61).Of the diphosphate nucleotides readily available, dTDP in concentrations as great as 1 mM gives negligible background velocities with the pyruvate kinase indicator system (8).

-

b. Hexokinase-Glucose-6-Phosphate Dehydrogenase Assay. The hexokinase-glucose-6-phosphate dehydrogenase method was developed originally by Berg and Joklik (S) who measured the formation of ATP from ADP and ITP or UTP in the presence of hexokinase-glucose-6-phosphate dehydrogenase indicator system according to the following equations:

+ + +

+

ITP ADP IDP ATP ATP glucose --t glucose 6phosphate Glucose &phosphate NADP -+ 6-phosphogluconate

+ ADP + NADPH + H+

(9) (10)

(11) Although this method is convenient, the only diphosphate nucleotide that can be studied is ADP and triphosphatc nucleotide must be a poor substrate for hexokinase. Various descriptions of this method have been given (7, 61). c. Other Coupled Enzymic Assays. Other coupled enzymic assays for N D P kinase have been developed, but their use has been limited to a few publications; for example, Sanadi et al. (29) have coupled the N D P kinase reaction with an a-ketoglutarate-succinate thiokinase system. Ratliff et al. ( 5 ) coupled the N D P kinase reaction to the phosphoglycerate kinase and triosephosphate dehydrogenase reactions, whereas Berg and Joklik coupled the N D P kinase reaction to the adenylate kinase 61. M. Chiga, A. Oda, and R . L. Holtaer, ABB 103, 366 (1963).

9.

NUCLEOSIDE DIPHOSPHOKINASES

325

and adenylate deaminase reactions (31). Nakamura and Sugino (SO) have developed a procedure that employs P3'P-labeled dTDP. Upon completion of the N D P kinase reaction, the products are digested with a snake venom preparation that liberates both pyrophosphate and phosphate which are separated by a specific method. The reaction is followed by measuring the incorporation of 32Pinto the pyrophosphate fraction (62). 2. Isotopic Procedures

One of the most commonly employed procedures for N D P kinase assays is the ATP-ADP exchange or a related isotopic method (1.4, 28, 63). I n a typical procedure, a l"C-labeled disphosphate nucleotide is incubated with a nonlabeled triphosphate nucleotide. Upon completion of the reaction, the di- and triphosphate nucleotides are separated, usually by chromatography, and the extent of incorporation of radioactivity into t.he triphosphate nucleotide is determined (8, 44, 5 2 ) . Some laboratories have employed 32P-labeled nucleotides for following the reaction ( 9 , 16, 59, 63). The isotopic method has the advantage that it does not impose the restrictions of a coupled enzymic assay procedure and makes possible measurement of the reactions of any nucleotide pair depending on the availability of labeled substrates. This procedure has the disadvantage of being somewhat cumbersome and time-consuming and of employing radioisotopes. In comparison with the coupled-enzymic methods, isotopic methods are inconvenient for some kinetic analyses such as initial velocity studies. A number of isotopic methods have been described (3, 8, 9, 16, 16, 19, 28, 44, 52, 53, 63).

3. Staining Procedure for N D P Kinase A convenient staining procedure has been developed for the identification of N D P kinase activity bands following gel electrophoresis or chromatography. This method is similar to the hexokinase-glucose-6phosphate dehydrogenase procedure described above with the exception t.hat the NADPH formed is detected by reaction with tetrazolium dyes (24). A useful method has been described where the complete indicator system is incorporated in an agarose overlay which becomes stained during the N D P kinase reaction (24). Since this staining method usually employs UTP and ADP as substrates, it is necessary to run control reactions for contaminating adenylate kinase (by omitting UTP) . 62. Y. Sugino and Y. Miyoshi, JBC 239, 2360 (1964). 63. M. Chiga and G. W. E. Plaut, JBC 234, 3059 (1959).

326

R. E. PARKS, JR. AND R. P. AGARWAL

D. KINETICSAND CATALYTIC MECHANISM 1 . Effect of Substrate Concentration There have been a number of reports describing the kinetic behavior of N D P kinase isolated from erythrocytes (7, 8, 24), yeast (16), calf liver mitochondria (1'7, 28, 64, 66),calf thymus (SO),beef heart mitochondria (15),and Bacillus subtilis (9). All enzymes that have been described to date display the classic kinetic behavior of a ping-pong reaction sequence (66) according to Scheme 1. With several N D P kinases,

A

A If

Y

NTP P

E-

SCHEME 1

including six isozymes from human erythrocytes, marked differences were observed in the kinetic parameters with a variety of nucleotide substrates as seen in Table 111. I n each case, the reactions were demonstrated to function via phosphorylated enzyme intermediates. Figure 2 shows the pattern of parallel lines observed with a human erythrocytic N D P kinase when the dGDP concentration was varied a t several concentrations of G T P ( 7 ) . In comparable experiments with other N D P kinase preparations, similar patterns of parallel lines have been described ( 9 , 1 5 4 8 ) .Figure 3 demonstrates the occurrence of the abortive complex, E-NDP, a t high concentrations of a diphosphate nucleotide such as ADP. Here, the inhibition is partially overcome by increasing the concentration of the triphosphate nucleotide, ATP. Similar results have been obtained with other N D P kinase preparations (9, 16-17, 64). I n studies of competing substrates, both di- and triphosphate nucleotides 64. A. Goffeau, J. Brachet, P. L. Pedersen, and A. L. Lehninger, Arch. I n t . Physiol. Biochim. 76, 179 (1968). 65. P. L. Pederscn, A. Goffeau, and A. L. Lehninger, Fed. Proc., Fed. Amer. SOC.E x p . B i d . 26, 609 (1967). 66. W. W. Cleland, BBA 67, 104 (1963).

9.

327

NUCLEOSIDES DIPHOSPHOKINASES

I60

o GTP concn. 0 GTP concn. e GTP c o x n 0 GTP concn.

= 1.0 x I O - ~ M

= 5.0 x = 1.0 x

M M

= 2.5 x

M

Ib

2b

0

I20

-VI 80

40

;

Ib

2;

3b

315

I mM dGDP

FIQ.2. Plots of reciprocal of the initial velocity (v = -AA340 min-') versus reciprocal of concentrations of dGDP (millimolar). Illustrates family of parallel line characteristics of the ping-pong reaction sequence. From Mourad and Parks (7).

have given the predicted classic competitive inhibition patterns (7, 9, 16).Studies of the inhibition of erythrocytic N D P kinase by the monophosphate nucleotide, GMP, gave K i values of 4 X M when d T D P or dGDP was the variable substrate and 6.5 X lO-'M when ATP was the variable substrate. I n both cases, noncompetitive patterns were ob240 200 160 -

-I

"

a

o GTP concn = I o x 0 GTP concn = 25 x 0 GTP concn = 5.0 X

M M M

120-

80 40

-

01 0

I

5

I

10

I

15

I

20

I ADP m M

FIG.3. Demonstration of abortive complex found at high concentration of ADP. See text for discussion. From Mourad and Parks (7).

328

R. E. PARKS, JR. AND R. P. AGARWAL

200

I60

-VI

I20

80

40

o0r

'

I

5

I

I

I

10

15

20

L r n M ATP

(ATP TDP = 1 2rl)

FIQ.4. Inhibition of erythrocytic NDP kinase by GMP with di- and triphosphate nucleotide substrates varied in a constant molar ratio. ATP and dTDP were varied in a ratio of 1:24. ATP and GMP were added in the concentrations indicated. Lineweaver-Burk plot is typical for competitive inhibition. From Mourad and Parks (7).

tained. However, when di- and triphosphate nucleotide substrates ATP and d T D P were varied in a constant ratio a t different concentrations of GMP, the reciprocal plot yielded a family of straight lines that met a t the l/v intercept, a pattern of competitive inhibition (Fig. 4) ( 7 ) . These results indicate that the inhibitor, GMP, competes for the nucleotide substrate-binding sites of both the free and the phosphorylated enzyme. A rate equation offered by Garces and Cleland (16)for the mechanism of Scheme 1 which predicts initial velocities in the absence of products but which allows for the formation of abortive complexes by both substrates is v =

VAB

KoBU

+ B/Kib) -I- KbAbA(1 + A/Ki,)

-I- AB

(12)

where K , and Ka are Alichaelis constants for N,TP (A) and N,DP (B), and Ki, and Kib are dissociation constants for N,TP and N,DP from abortive complexes with phosphorylated enzymes and free enzyme, and V is the maximal velocity. 2. pH Optima and E f f e c t of Temperature

Several laboratories have described the p H optima for the reactions of various preparations of N D P kinase. In general, the opt,ima range from

9.

NUCLEOSIDE DIPHOSPHOKINASES

329

pH 6-9, and in most cases between p H 6.5 and 8.0 (8, 16, 88, 30,31, 49). A Dixon analysis (67) of the effect of pH on the kinetic parameters has been performed with the PI 7.3 isozyme from human erythrocytes (8). In experiments with dTDP as the varying substrate, little change was observed in the Michaelis constant between pH 5.5 and 9.0. The plot of log V,,, versus pH revealed a relatively broad optimum with downward breaks at pH values of about 6.3 and 9.2. The downward break a t pH 6.3 is consistent with the ionization of a histidine residue a t the active site. It is intriguing that WBlinder (80) has identified phosphorylated residue of histidine and lysine in alkaline hydrolysates of human erythrocytic N D P kinase phosphorylated by reaction with [ y S 2 PATP. ] Since several studies have indicated that sulfhydryl groups play an important role in the activity of this enzyme, i t is of interest that with the PI 7.3 isozyme a clearcut break is not seen in the plots of pK, or log V,,, versus pH, from pH 8 to 9, which would be consistent with the occurrence of a sulfhydryl group (8). Detailed studies of the temperature dependence of N D P kinase have been performed with the six isozymes from human erythrocytes and with a homogeneous preparation from B. subtilis (8, 9, 2.4). With the B. subtilis enzyme and the PI 7.3 and 8.3 isozymes from human erythrocytes, biphasic Arrhenius plots were observed. In each case, lower activation energies were observed at the higher temperatures. With the human erythrocytic isozymes, the break points occurred a t about 31", with the B. subtilis enzyme a t about 25". It is of interest that the other four human erythrocytic N D P kinase isozymes yielded linear Arrhenius plots. The fact that Arrhenius plots of two of the human erythrocytic isozymes are biphasic and the remaining four are linear has not yet been explained but suggests the existence of a fundamental difference in the physicochemical behavior of these isozymes.

E. METALREQUIREMENTS Many reports have appeared describing the metal dependency of N D P kinases (8, 9, 16, 17, 28, SO, S1, 49, 6 2 ) . I n no case has a requirement for a monovalent cation been described. However, all N D P kinases have an absolute requirement for divalent cations. Where examined, Mg2+ and Mn2+ have similar activity (8, 28, SO, 49, 62).Some differences have been observed from one N D P kinase preparation to the next in the activity of other divalent cations such as Ca'+, Co2+, and Zn2+ (8, 9, $8,SO, 31, 49, 5 2 ) . In studies with NDP kinase from beef heart mito67. M. Dixon, BJ 55, 161 (1953).

330

R. E. PARKS, J R . AND R. P. AGARWAL

chondria, Colomb et al. (16) uncovered evidence indicating that the true substrates for the enzyme are Mg”-nucleotide complexes, such as Mg-ATP. In studies with a purified N D P kinase from liver mitochondria, it was found that Mg-ADP decreased the sensitivity of the enzyme to sulfhydryl reagents, whereas in the presence of Mg2+ alone the inactivation of the enzyme was enhanced. It was suggested that the Mg2+ stabilizes one of the equilibrium forms of the enzyme that possesses an essential sulfhydryl group that is more sensitive to reaction with mercurial agents (68).

F. SULFHYDRYL GROUPS All N D P kinase preparations that have been examined give evidence of essential sulfhydryl groups (8, 14, 16, 68, 49, 57, 64, 68). In most cases, the enzymes are inhibited by mercurial reagents such as p-chloromercuribenzoate but they may be reactivated by the addition of a thiol such as cysteine, dithiothreitol, or mercaptoethanol (8, 14, 16, 68). With a homogeneous preparation of an N D P kinase from bovine liver mitochondria, titration studies revealed two classes of sulfhydryl groups. Four sulfhydryls per mole of enzyme are apparently free and react readily with p-chloromercuribenzoate, whereas a second group of four “hidden” sulfhydryl groups react with p-chloromercuribenzoate only when the enzyme is denatured with sodium dodecyl sulfate. Silver nitrate titrates both classes of sulfhydryl groups readily. Presumably the smaller size of the silver ion (Ag+) allows it to reach the “hidden” sulfhydryls that are inaccessible to the much larger p-chloromercuribenzoate (67). It is also of interest that a t low concentrations of a thiol reagent (below 10-‘M) a slight stimulation of enzymic activity was observed (68). Studies of the protective action of nucleotide substrates against the effects of thiol reagents have been performed with the human erythrocytic enzyme and bovine liver mitochondria1 enzyme (8, 1.6, 68). I n all cases, the presence of di- or triphosphate nucleotide substrates gave significant protection against inactivation by mercurials. Of particular interest is the observation that somewhat greater protection of the PI 7.3 isozyme from inactivation by p-chloromercuribenzoate was afforded by ATP or dTDP in the absence of a divalent cation ( 8 ) . These observations suggest the occurrence of a nucleotide binding site on the enzyme that is not identical with the catalytic site but which influences the conformation of the enzyme. In other studies with the PI 7.3 human erythrocytic isozyme, the sensitivity to thiol reagents were increased by the addition 68. A. Goffeau, P. L. Pedersen, and A. L. Lehninger, JBC 243, 1685 (1968).

9.

NL'CLEOSIDE

DIPHOSPHOKINASES

331

of 2 M urea. Moreover, in the presence of urea the enzyme could not be reactivated by addition of dithiothreitol which readily occurs in the absence of urea (8). This suggests the occurrence in the erythrocytic enzyme of a t least two types of sulfhydryls as was observed with the bovine liver mitochondria1 enzyme (67).

G. CONFORMATIONAL CHANGW Several pieces of evidence indicate that NDP kinases from a variety of sources are susceptible to conformational effects. Where studied, the enzyme has proved to be polymeric and in the case of yeast NDP kinase the enzyme is a homologous hexamer. This suggests the possibility of interactions between subunits. With a homogeneous NDP kinase isolated from calf liver, a sigmoid-shaped rather than hyperbolic substrate saturation curve was obtained with ATP. This yielded a Hill coefficient of 1.8 ( 1 7 ) . Conformational changes have also been observed with the PI 7.3 isozyme from human erythrocytes. As noted above, marked protection against the inactivation by mercurial reagents is brought about by the addition of nucleotide substrates. Furthermore, the greatly enhanced susceptibility to mercurial reagents in the presence of 2 M urea is significantly diminished in the presence of a nucleotide substrate such as ATP (see Section 111,F). Also, as noted above, with several N D P kinases diphasic downward Arrhenius plots are obtained which might indicate the existence of different conformational states as the temperature is varied (8, 9, 24).

IV. Functions in the Cell

It is generally assumed that N D P kinase is a major component of the enzymic pathway for the synthesis of triphosphate nucleotides, many coenzymes, and RNA and DNA. The activity of N D P kinases in many tissues is relatively high, usually 10- to 100-fold greater than the activity of the monophosphate nucleotide kinases. Furthermore, the N D P kinases studied to date do not display absolute specificity for nucleotide substrates, which is in sharp, contrast to the monophosphate nucleotide kinases which are highly substrate-specific. Therefore, it seems unlikely that major control points in the pathways of triphosphate nucleotide synthesis occur a t the level of N D P kinases and perhaps would be found a t the level of the monophosphate nucleotide

332

R. E. PARKS, JR. AND R. P. AGARWAL

kinases or a t an earlier step. However, there are several indications that N D P kinases are responsive to the energy charge of the cell (69) and show conformational changes (8, 17) that are often seen in regulatory enzymes. In view of the high activity of N D P kinases in many cells, one would expect to observe a very rapid, indeed, almost instantaneous, equilibration of the high-energy phosphate bonds among the various nucleotide species. This should result in the intracellular ATP :ADP ratio being reflected in the triphosphate: diphosphate nucleotide ratios in the guanine, uracil, etc., polyphosphate nucleotide series. However, where nucleotide pools have been examined in cells that contain high levels of N D P kinase, such corresponding polyphosphate nucleotide ratios have not always been seen (33). This may, in some instances, result from compartmentation in the cell. In the consideration of presumed N D P kinases, one must continually bear in mind the possibility that an enzyme that has low specificity for di- and triphosphate nucleotide substrates and that functions through the reversible formation of a high-energy phosphate-to-enzyme bond can behave as an N D P kinase. For example, an enzyme that functions via the following reaction sequence might be considered an N D P kinase if it has low specificity for nucleotide substrates and if one examines only reaction (13) of the sequence:

-

+

+

E N T P S E P NDP E-P+X-+E-X+ P, E X Y + E XY

-

+

+

(13) (14)

(15) I n this case, X and Y could be any non-nucleotide metabolites. Although to date there have been no definitive reports of this type of correlation between a presumed N D P kinase and other enzymic activities, the possible occurrence of such reactions must he borne in mind. For example, a succinic thiokinase that can react with adenine or guanine nucleotides (70)would behave as an N D P kinase in the presence of ATP and GDP. Mention should be made of the recent report of N D P kinase activity in highly purified preparations of DNA polymerase from bacterial sources ( 4 7 ) .It is conceivable that the N D P kinase activity with this enzyme represents only a segment of a total reaction sequence. An observation that suggests the occurrence of specialized functions for N D P kinases is the wide variation in the kinetic parameters of the human erythrocytic isozymes with various nucleotide substrates ($4). A finding that points to another function for an N D P kinase is the demonstration of an acid-stable, alkali-labile high-energy phos69. F. M.Thompson and D. E. Atkinson, BBRC 45, 1581 (1971). 70. B. F. Rurnham, Acta Chem. Scnnd. 17, 5123 (1963).

9.

NUCLEOSIDE DIPHOSPHOKINASES

333

phate-enzyme bond in an N D P kinase from pig kidney (35), whereas most other N D P kinases form acid-labile, alkali-stable enayme-phospha te bonds. There is still much uncertainty about the role played by N D P kinases in such functions as mitochondrial oxidative phosphorylation, membrane transport involving ATPases, the substrate-level phosphorylation of G D P by the succinic thiokinase reaction, and the transport of high-energy phosphate from mitochondria t o cytoplasm. Since N D P kinases can catalyze ATP-ADP exchanges, there is still some confusion involving certain unique ATP-ADP exchange reactions found in mitochondria and N D P kinase reactions. These ATP-ADP exchange reactions are highly specific for adenine nucleotides (71). They are inhibited by both oligomycin and 2,4-dinitrophenol and are membrane-associated (39, &Iwhereas ), N D P kinases are insensitive to these reagents, are nonspecific, and are not membrane-bound. An Mg*+-dependent mitochondria1 N D P kinase was separated from an oligomycin, 2,4-dinitrophenol-sensitivephosphorylating membrane fraction (41, 43). I n digitonin-treated mitochondria, N D P kinase is associated with the outer membrane and the soluble fraction, whereas adenylate kinase occurs entirely in the soluble fraction (41, 43). In view of the recent finding of marked heterogeneity of N D P kinases in many tissues and subcellular fractions, including mitochondria ( 2 2 ) , it is obvious that considerably more investigation will be required before definitive functions can be assigned to N D P kinases with confidence. ACKNOWLEDQMENTS This chapter was prepared with the assistance of funds from Grant CA 07340 from the National Cancer Institute of the U. S. Public Health Service.

71. G . 8. P. Groot. nnd S. G . VandenBergh, BBA 153, 2!2 (1968).

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3-Phosphoglycerate Kinase R. K. SCOPES

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I. Introduction . . . . . . , . 11. Biological Behavior of Phosphoglycerate Kinase . A. Reaction Catalyzed . . . . . B. Biological Occurrence . . . . . C. Species Variation and Genetics . . . 111. Isolation and Molecular Properties . . . A. Purification Procedures . . . . . B. Molecular Properties . . . . . . IV. Reaction Kinetics . . . . . . . . A. Kinetics of the Back and Forward Reactions B. Nucleotide Specificity . . . . . C. Metal Ion Specificity . . . D. Postulated Mechanisms of Reaction . . V. Conclusion . . . . . . . . .

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.

.

. . . .

. .

.

. . . .

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335 336 336 337 338 340 340 342 346 346 348 349 349 351

I. Introduction

The isolation of 1,3-diphosphoglycerate as an intermediate in the oxidation of glyceraldehyde 3-phosphate to 3-phosphoglycerate was the first indication that two separate enzymes were involved at this point in the glycolytic pathway ( 1 ) . Not long afterward, the enzyme S-phosphoglycerate kinase was isolated from yeast ( 2 ) . The crystalline yeast preparation (3) has been the principal supply of the enzyme and available for over 20 years; nevertheless, compared with other enzymes of the 1. E. Negelein and H. Bromel, Biochem. 2. 301, 135 (1939); 303, 132 (1939). 2. T. Bucher, Natzcrwksenachuften 30, 756 (1942). 3. T. Bucher, BBA 1, 292 (1947).

335

336

R. K. SCOPES

glycolytic pathway, PGK (Su) has been somewhat neglected until recently. This neglect is rapidly being put right, and the next few years should see the elucidation of its reaction mechanism a t the atomic level.

II. Biological Behavior of Phosphoglycerate Kinase

A. REACTION CATALYZED 3-Phosphoglycerate kinase catalyzes the transfer of "energy-rich" phosphate from the acid anhydride bond of 1,3-DPGA (Su) to the terminal phosphate of ADP. Divalent cations are essential to the reaction since the magnesium complexes of the nucleotides are most likely the true substrates (4-6'), see reaction (1).

iII

-0- 40

I

-0-P-O-

d

The forination of ATP, as in the direction of glycolysis, is generally termed the forward direction of reaction. The reaction equilibrium, essentially independent of pH, favors ATP production :

K,

3.1-3.4 X 103 a t 25" (3, 7) AGO = -4.80 f 0.05 kcal/mole = -17.5 f 0.2 kJ/mole However, in physiological conditions and in the usual assay procedures, the reaction is tightly coupled to the preceding glycolytic enzyme, GAPDH (Su),and the overall reaction is more nearly in balance: GAP-

=

+ NAD+ + P:- + MgADP-

3-PGA3-

+ NADH + MgATP*- + H+

(2)

3a. Abbreviations: PGK, 3-phosphoglycerate kinase, ATP-3-phospho-D-glycerate 1-phosphotransferase (EC 2.7.2.3.) ; 3-PGA, 3-phospho-D-glycerate; 1,3-DPGA, 1,3diphospho-D-glycerak; GAPDH, glyceraldehydephospliate dehydrogcnase (EC 1.2.1.12); and GAP, n-glyceraldehyde 3-phosphate. 4. M. Larsmn-Rainikiewicz, BBA 132, 33 (1967). 5. M. Larsson-Rahikicwicz and B. G. Malmstrom, ABB 92, 94 (1961).

10.

337

3-PHOSPHOGLYCERATE KINASE

[Although the equilibrium constant for GAPDH a t p H 7.0 is reported to be 0.5 ( 8 ) , with only two products from three substrates the equilibrium is dilution-dependent, being shifted toward GAP (3a) production at physiological substrate concentrations.] Thus the two enzymes can act concertedly in either direction, depending on such factors as ATPADP and NADH-NAD’ ratios, Pi level, and pH. 3-Phosphoglycerate kinase is involved in carbon fixation in many plant tissues (9); it is the enzyme reacting immediately after one of the primary carbon fixing enzymes, ribulosediphosphate carboxylase, EC 4.1.1.39, reaction (3).

+ HCOa- + 2 3-PGA3- + H+ + MgATP2- S 1,3-DPGA+ + MgADPH+ + 1,3-DPGA4- + NADPH eGAP2- + NADP+ + P,2-

Ribulose 1,5diphosphate43-PGAB-

(3)

(4)

(5) (6) This reverse reaction of PGK is favored during photosynthesis by the high ATP-ADP and NADPH-NADP’ ratios. GAP*- -+carbohydrate

+ Pi2-

B. BIOLOGICAL OCCURRENCE 3-Phosphoglycerate kinase has been shown to occur in a variety of higher plants (10)and in blood (3), but it exists in highest concentrations in yeast, about 2 mg/g wet weight (If) and in muscle, 1-1.5 mg/g (1.2, 13). Because of compartmentalization within the cells, it probably exists a t concentrations up to four times greater than these (l4), which is up to 0.1 mM. This is of the same order as the normal concentration of one of its substrates in the forward direction, MgADP, and far greater than the concentration of 1,3-DPGA ever reaches (16). As a consequence, the forward reaction rate is first order with respect to 1,3-DPGA concen6. M. Larsson-Rainikiewicz, BBA 85, 60 (1964). 7. W. K. G. Krietsch and T. Bucher, Eur. J. Biochem. 17, 568 (1970). 8. K. Burton and H. A. Krebs, BJ 54, 94 (1953). 9. M. Calvin and J. A. Bassham, “Photosynthesis of Carbon Compounds.” Benjamin, New York, 1962. 10. B. Axelrod and R. S. Banduski, JBC 204, 939 (1953). 11. R. K. Scopes, BJ 122, 89 (19’71). 12. R. Czok and T. Bucher, Advan. Protein Chem. 15, 315 (1960). 13. R. K. Scopes, BJ 113, 551 (1969). 14. R. K. Scopes, in “The Physiology and Biochemistry of Muscle as a Food” (E. 6.Briskey, R. G. Cassens, and B. B. Marsh, eds.), Vol. 2, p. 471. Univ. of Wisconsin Press, Madison, 1970. 15, H. J. Hohorst, M. Reim, and H. Bartels, BBRC 7, 142 (1962).

338

R. K. SCOPES

tration in these tissues, despite a very low K, value for this substrate (see below). The specific activity of the enzyme is high, particularly in view of the fact that there is only one active center on the molecule of 47,000 daltons (see below). The maximum theoretical forward direction throughput in muscle is about 5000 pmoles/g/min at 37". However, the maximum possible rate of glycolysis in muscle, calculated from phosphorylase content and activity (16, l 7 ) , would produce only about 200 pmoles 1,3-DPGA/ g/min, and the maximum rate of production of MgADP calculatcd from myofibrillar ATPase is about 250 pmoles/g/min (18) (part of which is rephosphorylated by pyruvate kinase). Thus, there is a t least 25 times more PGK than necessary to cope with even the fastest glycolytic rate in muscle, and this "overkill" must be even greater in yeast. The same applies to GAPDH which also has a theoretical maximum throughput of about 5000 pmoles/g/min. This enzyme has a specific activity of only about 0.1 times that of PGK, but its concentration in muscle is some 10 times greater (12).The relative proportions of these two enzymes and of the subsequent enzymes of glycolysis have been demonstrated to be nearly constant in a wide variety of tissues (12,19).It has been suggested that there may be specific interactions between the enzymes of this group, particularly between PGK and GAPDH, to aid transfer of common substrates. However, the large excess of enzymes would seem to make such an association unnecessary, a t least in muscle and yeast. C. SPECIESVARIATIONAND GENETICS Krietsch and Bucher (7) have isolated PGK from yeast and rabbit muscle and compared many of their kinetic and molecular properties; differences found were mostly slight. Amino acid analysis shows that yeast PGK contains far fewer sulfur atoms than muscle enzyme (a common situation vk-&-vis yeast-muscle enzyme comparisons) and is insensitive to thiol reagents. The muscle enzyme, on the other hand, is rapidly inactivated by p-chloroniercuribenzoate, 5,5'-dithiobis(Z-nitrobenzoic acid), or traces of heavy metals during purification. Comparison of electrophoretic mobilities on starch gel has indicated only slight variations in mammalian muscle PGK's with a somewhat more acidic nature of the enzyme in many fish species (20, 2 1 ) . A selec16. E. H. Fischer and E. G. Krebs, JBC 231, 65 (1958). 17. H. D. Engers and N. B. Madsen, BBRC 33, 49 (1968). 18. J. R. Bendall, BJ 81, 520 (1961). 19. D. Pette, W. Luh, and T. Bucher, BBRC 7, 419 (1962). 20. R. K. Scopcs, BJ 107, 139 (1988). 21. R. K. Scopes, unpublished data (1972).

10.

339

3-PHOSPHOGLYCERATE KINASE

tion of PGK mobilities is shown in Fig. 1. I n all cases the band on starch gel has a characteristic shape with a sharp curved cathodic edge, and an indistinct anodic edge, indicating interaction with both buffer ions and the starch itself. Crude extracts of whole muscle normally give only a single band staining for PGK activity, but purified preparations run in the absence of thiols often give two bands or a less distinct result (17, 13). Multiple bands of erythrocyte PGK appear, which resolve to a single band when ATP is present during electrophoresis ( 2 2 ) . The yeast enzyme, prepared from an ammoniacal cytolysate, gives only a single band with about the same mobility as the nonmammalian muscle enzymes, however, separate components have been isolated from the commercial products by column electrophoresis (23). It seems likely that these minor components are degraded forms of the original enzyme, produced during autolysis of the yeast a t neutral pH. Genetic investigations using gel electrophoresis to detect mobility variations have recently provided evidence that the PGK gene is located on the X chromosome. A study on human erythrocytes failed to find any polymorphism (22), but similar investigations with kangaroos (24) demonstrated mobility variation both between and within species. Family studies of thc polymorphisms showed clearly that PGK is X chromosome-linked; in females the paternal gene is not expressed; thus, females do not have a double dose of the enzyme. More precise location of the PGK genc, on thc long arm of thc X chromosome, is indicated from a cytological study of human-mouse and human-hamster hybrid Rabbit

Horse pig

Carp

Tortoise

Yeast

Pike

.

.

Frog

tm

+ FIG.1. Mobilities of PGK on starch gel clectrophoresis, pH 8.3 ( 1 8 ) . In 4 hr at about 2 V/mm, the pike enzyme moved 25 mm. 22. E . Beutler, Bwchem. Genet. 3, 189 (1969). 23. M. Larsson-Rainikiewicz, Eur. J . Biochem. 15, 574 (1970). 24. D. V. Cooper, J. L. Van de Berg, G . B. Shnrman, and W. E. Poole, Nature (London) 230, 155 (1971).

340

R. K. SCOPES

cells (25). Earlier evidence for X chromosome linkage in humans came from studies of a hereditory hemolytic anemia associated with PGK deficiency ( 2 6 ) . Another variant, discovered in a New Guinea population, has been found to have a considerably faster mobility in starch gel electrophoresis (27). This variant has been studied in detail, and exhibits some unusual features (28, 29). The isoelectric point appears to be substantially less than that of the normal human erythrocyte enzyme (4.7 compared with 7.9),yet amino acid analysis and tryptic peptide mapping indicates that only one residue is involved, and that the substitution is neutral, being Thr- Asn. In the absence of citrate, the mobility difference in gel electrophoresis is abolished, which suggests that the variant has a strong affinity for citrate ions and perhaps also for the ampholytes used in isoelectric focusing.

111. Isolation and Molecular Properties

A. PURIFICATION PROCEDURES 3-Phosphoglycerate kinase has been isolated in crystalline form from yeast (3, 7, 11), rabbit (7, IS),pig ( I S ) , horse (So),and ox (21) muscles, and from human erythrocytes (98, 31). Purified preparations have also been made from chicken muscle (32), pigeon breast muscle ($I), pea seeds ( l o ) , and sheep brain (33). The pure enzymes from yeast and muscle catalyze the phosphorylation of 3-PGA by ATP with theoretical maximum velocity of 1200-1300 pmoles/mg/min a t 30" (7, 11, 13, 23) [calculated using the expression 25. K. H. Grzesrhik, P. W. Allderdice, A. Grzesrhik, J. M. Opitz, and 0. J. Miller, €'roc. Nat. Acad. Sci. U.S. 69, 69 (1972). 26 H.4. Hsieh, D. E. Paglia, H. M. Anderson, M. A. Banghan, E . R. Jaffe, and 0. M. Garson, N . Engl. J . M e d . 280, 528 (1969). 27. S.-H. Chen, L. A. Malcolm, A. Yoshida, and E. R. Giblett, Amer. J . Hum. Genet. 23, 87 (1971). 28. A. Yoshida and S. Watanabe, JBC 247, 440 (1972). 29. A. Yoshida, S. Watanabe, S.-H. Chen, E. R. Giblett, and L. A. Malcolm, JBC 247, 446 (1972). 30. C. C. F. Blake, P. Evans, and R. K. Scopes, Nature (London) 235, 195 (1972) 31. T. Hashimoto and H. Yoshikawa, BBA 65, 355 (1962). 32. C. Gosselin-Rey, BBA 87, 140 (1963). 33. Y. Leli&vre, BBA 206, 187 (1970).

10. 3-PHOSPHOGLYCERATE KINASE

341

(see reference 4 ) ] .This corresponds to a turnover number of 1000 sec-l. Crystalline preparations occasionally contain some inactive PGK protein which lowers the specific activity. The enzyme is most conveniently assayed by the reverse reaction, coupling to GAPDH and measuring the oxidation of NADH spectrophotometrically (3).In the forward direction the maximum reaction velocity is between 21/2 and 3 times greater (7) ; it also can be measured by coupling with GAPDH in conditions favoring the forward direction. Alternatively, freshly prepared 1,bDPGA can be used, detecting the ATP produced either by a suitable radioisotope method (34) or by using the glucose-hexokinase-glucose-6-phosphate dehydrogenase system, spectrophotometrically or fluorimetrically. To purify the enzyme from muscle, low ionic strength extracts are fractionated with ammonium sulfate, heat treated, and subjected to gel filtration and ion exchange chromatography to give a crystallizable preparation (7,13). Yeast can be autolyzed after drying, with toluene, or simply with ammonia to yield a solution which also is fractionated by successive ammonium sulfate, ion exchange, and gel filtration steps (11, 36). Overall recovery of crystalline PGK from muscle is up to 400 mg/kg and from yeast 800 mg/kg wet weight. Triosephosphate isomerase, EC 5.3.1.1, has similar physical properties to PGK and accompanies the enzyme through to the ion exchange chromatography steps. I n the correct conditions, PGK is adsorbed on CM-cellulose whereas triosephosphate isomerase is not, and conversely for DEAE-cellulose. The muscle enzymes crystallize readily from ammonium sulfate solution into needles which may grow to several millimeters in length. Of those investigated, only the horse muscle enzyme has given crystals of sufficient cross section to be useful for X-ray crystallography (30).The yeast enzyme also gives needles in many conditions but can be persuaded to grow into more chunky forms ( 1 1 ) ; these have also proved satisfactory for X-ray crystallography (36). In both cases, the larger crystals grow more readily in the presence of 0.5-30/, v/v 1,4-dioxan (in addition to the ammonium sulfate). The human erythrocyte PGK has also crystallized as needles (28, 31). 34. D. R. Rao and P. Oesper, BJ 81, 405 (1961). 35. W. K. G . Krietsch, P. G. Pentchev, H. Klingenburg, T. Hofstiitter, and T. Biicher, Eur. J . Biochem. 14, 289 (1970). 38. H. C. Watson, P. L. Wendell, and R. K. Scopes, JMB S7, 623 (1971).

342

R. K. SCOPES

B. MOLECULAR PROPERTIES 1. Molecular Weight 3-Phosphoglycerate kinase has a molecular weight in the range of 45,000-50,000 daltons. Earlier sedimentation measurements underestimated the value (6) ; gel filtration also suggests values lower than this

(13,37). The various values for the molecular weight of PGK that have been reported are listed in Table I (6, 7, 11, 13,23, 28, 37-40). The “subunit” size of PGK’s, as determined by SDS-gel electrophoresis is 48,00050,000 daltons, indicating that the enzyme is monomeric. Amino acid analysis, in papicular the single cysteine of the yeast enzyme, and X-ray crystallographic studies all strongly support the lack of subunit structure. 2. Primary Structure

The amino acid analyses of rabbit and yeast enzymes have been compared and found to be very similar (7). An independent analysis of the yeast enzyme gave almost identical results (23). TABLE I REPORTED MOLECULAR WEIGHTSOF PHOSPHOQLYCERATE KINASIC MW x lo-’ DALTONS Method Gel filtration Ultracentrifuge

Muscle Ref. 38 47 41 47

1s 7

Ref.

Erythrocytes

Ref.

37 47

37 7

50

38

34 45 47 46 45 47 50

6

50

28

23 7 7 2s 11 11

49 50

28 28

59

7

Amino acid analysis

45

7

Tryptophan contenta SDS*-gel electrophoresis

48 48

13 40

a

Yeast

Assumes 4 and 2 Trp residues in mammalian and yeast enzymes, respectively.

* SDS stands for sodium dodecyl sulfate.

37. J. B. Alpers, JBC 243, 1698 (1968). 38. S. Watanabe and A. Yoshida, Fed. Proc., Fed. Amei. SOC.Exp. Biol. 30, Abstr. 903 (1971). 39. C. T. Walsh and L. B. Spector, JBC 246, 1255 (1971). 40. R. K. Scopes and I. F. Penny, BBA 236, 409 (1971).

10.

343

3-PHOSPHOGLYCERATE KINASE

Recently, the human erythrocyte enzyme has been purified, and its amino acid composition is seen to be similar to that of the rabbit muscle (98).The amino acid compositions of the three types of PGK are listed in Table 11. The interesting feature of the yeast enzyme is that it possesses only one cysteine residue per molecule, and that is not a t the active center (see below). Methionine is also poorly represented in the yeast enzyme with the result that mammalian PGK's contain a t least five times as much sulfur as the yeast enzyme. No sequence of peptides from PGK has yet been reported. The only published information is that the C-terminus on the muscle enzyme is valine, and no N-terminal residues could be detected as dansyl derivatives (7),whereas on the erythrocyte enzyme, the C-terminus is isoleucine, and the N-terminus, N-acetylserine (98).

TABLE I1 AMINOACID COMPOSITION OF PHOSPHOQLYCERATE KINASP Rabbit muscle

Human erythrocytesd

Amino acid

Yeastb

Yeastc

Lysine Histidine Arginine Aspartic acid Threoiiine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Cysteine Tryptophan

44 8 13 42 19 25 39 18 39 45 38 3 21 41 7 18 1 2

46 9 14 41 18 25 38 18 37 43 38 3 23 41 8 19 1 2

40 39 38 13 18 38 4 16 8 4

40 5 11 45 17 24 34 19 41 41 41 13 18 40 4 15 10 4

423

424

420

422

Total

46 8 12 44 17 24 33

18

Nearest integer values normalized to residue numbers of 420-424 (7). From Larsson-Rainikiewicz (83). c From Krietsch and Biicher (7). d From Yoshida and Watanabe (88).

1'

b

344

R. K. SCOPES

3. Secondary Structure The magnitude of the Cotton effect in yeast PGK using optical rotatory dispersion has been reported; it was interpreted as indicating a moderate amount of a-helix in the protein (41). However, since the commercial preparation available a t that time was considerably contaminated with triosephosphate isomerase and other proteins (23,35) the results are of marginal value. Some further knowledge of secondary structure can be expected from X-ray crystallographic studies; the 6-A resolution map of the horse enzyme (SO, see below) indicates several possible helical sections in the polypeptide configuration. In particular, the connecting bridge between the two dissimilar globular portions may well be a rigid helix. 4. Tertiary Structure

The investigation of PGK by X-ray diffraction is at the moment of writing a t a stage where the first indications of the enzyme’s shape are beginning to emerge. The yeast enzyme is being studied a t Bristol, and the horse muscle enzyme a t Oxford, both as part of a concerted effort of X-ray crystallography of the glycolytic enzymes as a whole. At present the information from the two groups is complementary, and preliminary reports indicate they are also mutually confirmatory. The first published report of the X-ray work came from the yeast enzyme studies ( 3 6 ) ,in which it was demonstrated that crystals treated with Hg2+ ions gave a difference Patterson function showing a single binding site for mercury; clearly the single cysteine residue was reacting. Later, it was shown that adenosine-soaked crystals gave slight differences which, when the two-dimensional electron density distribution was calculated, showed slight conformational shifts a t a site remote from the mercury binding site (42). Aurocyanide ion also forms heavy metal derivatives with the yeast enzyme, two gold atoms being bound, one a t the same position as the adenosine site, the other close to but not a t the mercury site ( 4 3 ) . Since aurocyanide also inactivates the enzyme, it would be a reasonable assumption that the adenosine site corresponded with the adenine nucleotide binding site a t the active center. However, this position appears to be a secondary weak binding area for both Au (CN)?- and adenosine. Treatment with Au (CN),-ions gave only 41. B. Jirgenssons, JBC 240, 1064 (1965). 42. J. W. Campbell, E. Duke, G. Hodgson. W. D. Mercer, D. K. Stammers, P. L. Wendell, H. Muirhead, and H. C. Watson. Cold Spring Hnrbor S y m p . Q u m t . Biol. 36, 165 (1971). 43. H. C. Watson and P. L. Wendell, personal roniniunirntion (1972).

10.

3-PHOSPHOGLYCERATE KINASE

345

one heavy metal binding site at the position close to the mercury site, and corresponding with the active center. A 5 A resolution electron density map for the yeast enzyme has now been computed (4S), and shows satisfactory agreement with that from the horse muscle enzyme. The horse muscle enzyme grows in rhombus cross-sectional prisms, and these crystals have a different space group to the yeast PGK crystals. One of the dimensions of the unit cell is only 36.3 k (SO), much smaller than the expected diameter of a hydrated spherical protein (about 50A) of this size. This was the first indication that PGK was far from being spherical. Because this muscle enzymc has seven sulfhydryl residues, heavy metal derivatives were more complex than with the yeast enzyme. The Au(CN),- ion binds a t five places, and ethyl mercuric phosphate at two. Nevertheless, it was possible to calculate, using these derivatives, a three-dimensional electron density map a t 6 A resolution (SO). A photograph of a model built according to the electron density distribution is shown in Fig. 2.

FIG.2. A 6-A resolution inodrl of the electron density map of horse muscle phosphoglycerntc kinnse. The arrow points to the MgADP binding site located in a shallow cleft on one of the globular portions of the molecule. (From Blake, Evans, and Scopes (1972) Nature 235, 195.)

346

R. K. SCOPES

The molecule appears asymmetric, composed of two globular units joined by a section which is difficult to interpret but may be a helix portion-some rigid structure would bc expcctcd to join the two parts. Further investigations are undcr way to obtain highcr resolution models. It is clear that the yeast PGK cannot contain any cystine bridges, and the fact that 7 of thc 8 rabbit muscle cnzymc’s cystcinc residues can bc reacted with p-mercuribcnxoatc (7’) makes i t improbable that any such bridgcs cxist. Thc wholc structurc of PGK must bc maintained without covalent bonds bctwecn separate parts of thc polypeptide chain.

IV. Reaction Kinetics

A. KINETICSOF

THE

BACKAND FORWARD REACTIONS

The kinetics of the enzyme have mostly been studied on the back reaction, coupling to GAPDH with oxidation of NADH. Nearly all of this work has been carried out by Dr. Martha Larsson-Rainikiewicz in Professor Bo Malmstrom’s laboratory, and the author is grateful to her for corrections and suggestions in this section. Detailed accurate studies are confounded by the problems involved in estimating the concentrations of the ionic species MgATPz- and MgADP-, particularly when in the presence of high concentrations of K+ or Na+ ions [e.g., Larsson-Rainikicwicz (44)1. The possible formation of sodium and potassium complexes with ATP and A D P has been partly corrected for by using values for the dissociation constant of MgATP*- and MgADP- which are too high per sc. Nevertheless, thc basic kinetic conclusions would be unaltered if lower values for these constants were used (45). Michaelis constants for all the reactants have been determined, and the valucs reported are listed in Table 111. I n not all of these investigations was the magnesium concentration sufficient to ensure complete complexing of the nucleotides, which may explain some of the variation. The valucs are further complicated by the fact that in the presence of a moderate amount of free Mgz+ ions, the Michaelis constants for both MgATPz- and for 3-PGA are increased about twofold, but only in the higher substratc conccntration range (4, 5 ) . With 3-PGA, this is an indirect rcsult of nonconipetitivc inhibition by Mgz+ ions. For MgATP2however, the inhibitions arc inorc complcx. With a small cxccss of Mg 44. M. Lal.sson-Rninikieu,icz, Eur. J. Biochem. 22, 506 (1971). 45. W. J. O’Sullivan and D. D. Pewin, Biochemistry 3, 18 (1964).

10.

347

3-PHOSPHOGLYCERATE KINASE

TABLE I11 REPORTED

K , VALUES FOR

PHOSPHOGLYCERATE KIN.wI.:

Forward Reaction

Yeast,

6.9

Muscle

7.0 7.5 7.0

0.20 0.40

0 . 0018

0

0.0020

$4

0.20 0.35

0,0022

7 34

Back Reaction

h',,,for MgATP2- (ATP) (mW Yeast

Miisrle

Erythrocytes Pea seeds

6.0 6.!) 7.0 x.0 7.8 7.5 6.0 7.0 8.0 7.5 7.5 7.3

0.11 0.32 0.33 0.40 (i.nnq

0.4s 0.48 0.48 0.42 0.37 (1 .w) 4.1

K,, for 3-PGA (mM) 0.85 0.20 1.300

1.07" 0.63 (1.23O) 0.69 (1.2W) 1.36" 1.226 1 .07a 0 . 4 2 (1.37") 1.1" 7.6

a Represerits the higher vahies found for ATP and 3-PGA at higher substrate conreutxations and in the presence of excess Mgz+ ions.

(Mg2+ concentrations maintained a t 1 miM) , a weak uncompetitive inhibition was observed (46). Uncompetitive inhibition was also obtained when magnesium was replaced by zinc, making ZnATP2- the nucleotide substrate. In this case the inhibition by uncomplexed Zn2+ions was very strong, with a K i of approxiinatcly 0.02 mM, compared with a value of 3 mM for hlg". At very low levcls of ATP and magnesium competitive inhibition by Rig" may bc involved (6),whereas a t high levels of each, noncompctitive inhibition is indicated (4). The Michaelis constants of each substrate are nevertheless independent of the concentration of the other substrate, which suggests that the reaction mechanism is of thc random ordcr typc a t least a t low levels of Mg2*. Somc anoinalics in thc behavior (in particular thc changes in Km in thc prcscncc of C X C ~ S Smagnesium) coulcl bc intcrprcted in terms of two binding sitcs for each substratc ( 4 ) , although there is no other evidence in favor of this theory. As would be anticipated, the back reaction is inhibited by ADP, but

348

R. Ii. SCOPES

not in the normal competitive way with ATP. Instead, ADP acts competitively with 3-PGA and noncompetitively with ATP (44). Thc I
B. NUCLEOTIDE SPECIFICITY Several other purinc nuclcotidc triphosphatcs can takc thc placc of ATP in the back reaction, but none as efficiently. The order of reactivity was found to be (at 2.5 mM nucleotide) ATP > ITP > GTP > dGTP > dATP for both ycast and niusclc cnzyincs (7) ; ITP had a slightly higher K , than ATP, but not sufficient to account for its lowcr activity in the system. Guanosine triphosphate had a significantly lower I<,, but V,,,,, was only about 50% that obtainccl with ATP. Only a tracc of activity (ca. 1%) was obtained with UTP, and none a t all with either C T P or TTP. The nuclcotidc diphosphates inhibited the back reaction in much the same way, except that thc cxtcnt of inhibition (using 10 m M N D P ) was in the order GDP > A D P > IDP (7). Monophosphates also inhibited (see also reference 44).

10.

3-PHOSPHOGLTCERATE KINASE

349

C. METALIONSPECIFICITY 3-Phosplioglyccratc kinasc has an absolute rcquircnicnt for mctal ions, and it has been demonstrated that the true substrates are most likely the magnesium complcxes of ADP and ATP (4-6). Kinctic studies show that, as with many 0 t h kiiiascs, Mn2+ will rcplacc Mg2+with alniost idciitical kinetics, tlic apparcnt I<, of &In2+ bcing slightly lower than that of Mg'+, reflcctiiig tlic smaller dissociation constant for MnATP'- compared with MgATP2- ( 5 , 4 6 ). CaATPZ-, ZnATPz-, CoATP", CdATP", and NiATP2- have all been shown to be alternative substrates, although NiATP'- was poor (46). No measurable activity was given by Be2+ and Fez+; Zn2+ ions strongly inhibited tlic enzymc uncompetitively, whereas other free divalent cations showed various inhibitory effects less strongly.

D. POSTULATED MECHANISMS OF REACTION As indicated in Section IV,A, PGK probably has a rapid random reaction mechanism with no mutual interference in binding of either ATP or 3-PGA. However, the forward reaction has not been studied in such detail, and from the kinetics of the back reaction there seem to be separate binding sites for ATP and ADP. These results are compatible with the scheme originally suggested by Bucher ( S ) , in which there are two binding sites for adenine nucleotides and one for 3-PGA. This is illustrated in Fig. 3. In the forward reaction it is postulated that 1,3DPGA binds across sites 1 and 2, with the acid-anhydride phosphate a t site 2, Fig. 3 (I). (In Bucher's original paper sitc 1 bound ADP and site 3, 3-PGA. Since the recent paper from his laboratory numbers the sites in the opposite way, the later formulation has been used here.) Then MgADP- binds to sitc 3 and the transfer of phosphate takes place as the adenine nucleotide leaves site 3 and links to 2 through the terminal phosphate. In the back reaction, MgATP and 3-PGA bind to sites 2 and 1, rcspcctively (1111, leaving opcn sitc 3. From the kinetics wc must conclude that site 3 now has a highcr affinity for MgADP than when ATP is not present a t site 2 (Ki for MgADP being 0.02 mM K , for MgADP 0.2 mM). Thus the back reaction, in the presence of ADP, leads to the formation of an abortive enzymcMgADP-MgATP-3-PGA complex (IV). Further support for this theory of two binding sites for the nucleotides comes from thc observation (7) that MgADP, but not 46. M. Larsson-Rainikiewicz, Eur. J . Biochem. 17, 183 (1970).

350

R. K . SCOPES 1,3-DPGA

pG A

+

WADP

1

pGA

1'

1

P

2

MgADP

~

3-PGA

+ MgATP

3

(1)

pGA

%ADP

i

1:

(Ira)

G '

1 1 A l

MgATP J 2 MgADP

I

3

(N)

FIG.3. Postulated reaction scheme for PGK. Formation of (11) from (I) could occur either by direct transfer of the acid-anhydride phosphate of 1.3-DPGA to Mg.4DP or through the phosphorylated intermediate (Ha) . MgATP, protects the muscle enzyme from inactivation by 5,5'-dithiobis (2-nitrobenzoic acid). This suggests that the reactive -SH groups are shielded by MgADP occupying site 3, but not by MgATP occupying site 2 (presumably NIgATP will not bind to site 3 a t all). Essential involvement of these -SH groups either in binding or in enzymic catalysis is unlikely. Krietsch and Bucher assumed from their results that the -SH group in the yeast enzyme was not involved in catalysis since the p-chloromercuribenzoate-treated enzyme was fully active. Nevertheless, the recent X-ray crystallographic analysis shows that this single cysteine, binding mercury, is quite close to the active site ( 4 3 ) . Evidence for a phosphoenzyme intermediate has been presented (39). The evidence is not, conclusive, especially since the preparation on which the experiments were carried out must have been far from pure. Such an intermediate would not be obligatory in the reaction sequence since there is no kinetic evidence for ping-pong reaction mechanisms. Nevertheless, it seems reasonable that the reaction could proceed via the transitory intermediate (IIa) in Fig. 3; the phosphate could be attached to a carboxyl of glutamic or aspartic acid by an acid-anhydride bond as present in the substrate 1,3-DPGA. Dissociation of 3-PGA and MgADP from the intermediate would then leave the phosphoenzyme.

10.

3-PHOSPHOGLYCERATE KINASE

35 1

The various kinetic and other observations could also be interpreted in terms of only two binding sites, one for the phosphoglycerate, the other for adenine nucleotide, together with a conformational change associated with substrate binding. In such an interpretation, the binding of, say, MgATP would cause a conformational change, whereas MgADP would not [cf. cracking of PGK crystals in the presence of ATP salts but not with ADP (SO)]. The conformational change would be involved in the reaction as an integral step in the catalysis. Such a system must a t present be purely speculative in the absence of further data.

V. Conclusion

Phosphoglycerate kinase is an enzyme of almost universal occurrence, being involved in carbon fixation in plants, for ATP generation in anaerobes utilizing all or most of the Embden-Meyerhof glycolytic pathway, and similarly as a vital link in the aerobic energy-producing chain in animals. From present X-ray crystallographic results it appears that the molecule has an asymmetric shape with about half the protein far removed from the active site. Since the protein does not normally exist as polymers, this extra part is not required for self-association but may have a role in combining with enzymes sharing a common substrate, notably glyceraldehyde-3-phosphate dehydrogenase. However, in muscle and yeast a t least, both of these enzymes are present in vastly greater quantities than appear to be necessary from the potential maximum throughput of these tissues, thus, a special association between them would appear to be unnecessary. It is possible that earlier in evolution the two enzymes were not present in such an excess, and a n association between them may have been not only desirable but even necessary for activity of either part. That is to say that PGK-GAPDH may have originally been a single enzyme complex which later separated into the two parts. As a phosphotransferase, the enzyme has fairly simple kinetic properties which can mostly be interpreted in terms of three binding sites for substrates. It is possible that the full details of the reaction mechanism a t the atomic level will be available before long from the X-ray diffraction investigations, which should provide a basis for the description of phosphotransferase reactions in general.

This Page Intentionally Left Blank

11 Pyruvate Kinase F. J. KAYNE I. Introduction . . . . . . . 11. Molecular Properties . , . . A. Enzyme Purification . . B. Composition . . . . . C. Structure . . . . . D. Cheinical Modificat.ion . . E. Conformational Change . . 111. catalytic Properties . . . . . A. Stoichiometry and Specificit,y B. Assay . . . . C. Thermodynamics . . . . D. Kinetics . . . . . . E. Control . . . . . . F. Catalytic Mechanism . .

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. .

.

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.

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. . . . . . .

353 355 355 358 358 360 361 364 364 371 371 372 378 379

1. Introduction

As with most other enzymes described in the preceding edition of this series, a large number of studies have been undertaken on the enzyme pyruvate kinase (ATP:pyruvate phosphotransferase, EC 2.7.1.40) since the review by Boycr ( 1 ) . However, the greatest amount of interest followed the realization of the importance of the reaction step in the glycolytic pathway catalyzed by this enzyme in the control of glycolysis and gluconcogenesis (2). This slightly reversible stcp is in fact bypassed in gluconcogcnic tissues and organisms by reactions of pyruvate 1. P. D. Boycr, “Tlic Enzyines,” 2nd cd., Vol. 6, p. 95, 1962. 2. M. C. Scrutton and M. F. Utter, AW~ZL. Rev. Biocheni. 37, 249 (1968). 353

354

F. J. KAYNE

C02-

coz-

I

I

I1

I

H++ ADP + C-OP0:- 2 ATP + C=O CHI carboxylase and P-enolpyruvate carboxykinase. The product pyruvate feeds into a number of metabolic pathways thus placing this enzyme a t a primary metabolic intersection. The previous review on pyruvate kinase outlined well the historical developments leading to the characterization of this phosphorylation step in glycolysis as well as many of the kinetic studies on the most readily obtainable enzyme, that from rabbit muscle. Although the enzyme’s presence had been demonstrated in many cells ( 1 ) it was some time before considerable purifications were obtained from mammalian tissues other than muscle or from other eukaryotic and prokaryotic cells. These preparations were mainly derived from experience with the purification of other unstable proteins as well as development of various column chromatographic methods. One additional source of interest in the study of pyruvate kinase arises from its variability in the human erythrocyte. Some recent publications have described the presence of genetic variants of this enzyme associated with inherited anemias of somewhat different pathologies (3,4 ) . As one might expect from the enzyme’s key role in glycolysis, changes in its catalytic ability lead to severe metabolic imbalance. Its function also suggests it to be ubiquitous, and, indeed, in organisms where it has been shown not to be present (6, 6) its role can be fulfilled by pyruvate phosphate dikinase. The enzyme has a number of attributes which made it of particular interest to various investigators. It was the first and perhaps the most clearly apparent example of an enzyme with an absolute requirement for a monovalent cation for its catalytic activity. The skeletal muscle enzyme was quite easily obtained in good yields (about 0.5 g/kg) of high purity and was stable over a period of a year. It was also recognized quite early to be a multisite subunit containing protein. In contrast, the enzyme from other sources has been somewhat difficult to stabilize both during and after purification. Because of this, relatively few structural and physical details have been studied for these enzymes in spite of CHs

3. P.Cartier,

A. Najman, J. P.Leroux, and H.Temkine, Clin. Chim. A C ~ 22, Q

(1968). 4. G. E. J. Staal, J. F. Koster, and J. G. Nijessen, BBA 258, 685 (1972). 5. R. E. Reeves, JBC 243, 3202 (1968). 6. R. E. Reeves, R. Menzies, and D. S. Hsu, JBC 243, 5486 (1968).

165

11.

PYRUVATE KINASE

355

the obvious interest in the allosteric kinetics usually exhibited by the enzymes from these sources. The following discussion will tend to emphasize the enzyme derived from skeletal muscle chiefly because it is the one for which most molecular data have been obtained.

II. Molecular Properties

A. ENZYME PURIFICATION Skeletal muscle pyruvate kinase is most commonly prepared from rabbit muscle by the method of Tietz and Ochoa (7) [see also Boyer (I)3 . The basic procedure utilized no chromatographic steps and can be conveniently scaled up to produce gram quantities of enzyme in a few days starting with commercially available frozen muscle. The product can be crystallized by slow addition of (NH,),SO, (7) or by allowing the enzyme to stand in the cold at high concentrations (>120 mg/ml) in dilute imidazole buffer, or, alternatively, fractionated once again in (NH,),SO, (8) and used directly. The product is considered quite pure by the usual criteria i.e., yielding a single sedimentation boundary and the appearance of only very faint additional bands upon polyacrylamide gel electrophoresis. However, Bondar and Pon (9) showed some evidence for the presence of an endogenous inhibitor which could be removed by chromatography on a CM-Sephadex column. It has also been suggested that the yellow color of concentrated solutions of purified protein resulted from a tight binding of some chromophore with an absorption spectrum similar to that (A,,, 412 nm) of pyridoxal 5'-phosphate (0.25 equivalent/mole enzyme) (8). A more recently reported preparation (10) is claimed to be substantially free of myokinase although it is not clear that this contamination really represents a problem with the Tietz and Ochoa preparation. The specific activity of the rabbit muscle preparations has been reported to be between 150 and 300 pmoles pyruvate/min/mg protein under optimal concentrations of substrates and activators (K++ Mg2+) a t 25" and p H around 7.5. Such enzyme is available from commercial sources. Recently, the yeast enzyme has been crystallized for the first time 7. A . Tietz and S. Ochoa, ABB 78, 477 (1958). 8. F. J. Kayne, ABB 143, 232 (1971). 9. R. J. L. Bondar and N. Pon, BBA 191, 743 (1969). 10. F. Noll, W. Heumann, and C . Littmann, Actn Biol. M e d . Ger. 16, 115 (1965).

356

F. J. KAYNE

by Roschlau and Hess ( 1 1 ) with a resulting activity of 250 pmoles/ min/mg protein. Other procedures for obtaining highly purified preparations from this source have been given by Hess and co-workers (12, I S ) and Hunsley and Suelter ( 1 4 ) . The former group has utilized the fortunate property of this enzyme’s ability to bind tightly to Blue Dextran (Pharmacia), a characteristic apparently shared by the erythrocyte enzyme (16). Affinity chromatography (12) or a column separation of the complex, followed by its dissociation, results in a very specific purification step. Hunsley and Suelter (14) have relied on the classic gradient elution methods with stabilization of the protein in an aqueous-glycerol medium during the preparation. One point worth noting here is the use of Saccharomyces carlsbergensis by the first group and S. cerevisiae by the second. This may be the source of some differences between the preparations as well as the properties of the enzyme. Tanaka et al. (16) have crystallized pyruvate kinase from both liver and muscle tissues of the rat. Purification of this liver enzyme can also be carried out by elution from a CM-cellulose column using the allosteric effector fructose 1’6diphosphate ( 1 7 ) . Other highly purified preparations have been reported for the enzyme from rat liver (18, 1 9 ) , pig liver (20), frog muscle (211, erythrocytes (22-27), hepatoma cells ( S r a ) , and bacteria (28, 29). Par11. P. Rijschlau and B. Hess, Hoppe-Seyler’s 2. Physiol. Chem. 353, 435 (1972). 12. P. Roschlau and B. Hess, Hoppe-Seyler’s 2. Physiol. Chem. 353, 441 (1972). 13. H. Bischofberger, B. Hess, P. Roschlau, H-J. Wieker, and 13. ZimmermannTelschow, Hoppe-Seylei’s 2. Physiol. Chem. 351, 401 (1970). 14. J. R. Hunsley and C. H. Suelter, JBC 244, 4815 (1969). 15. K. G. Blume, R . W. Hofhauer, D. Busch, H. Arnold, and G. W. Lohr, BBA 227, 364 (1971). 16. T. Tannka, Y. Harano, F. Sue, and H. Morimura, J. Biochem. (Tokyo) 62, 71 (1967). 17. H. Carminatti, E. Rozengurt, and L. Jimenea d r Asua, FEBS Left.4, 307 (1969). 18. L. Jimenez de Asua, E. Rozcngurt, and H. Carrninatti, JBC 245, 3901 (1970). 19. W. A. Susor and W. J. Rutter, BBRC 30, 14 (1968). 20. C. Kutzbach and B. Hem, Hoppe-Seyler’s 2. Physiol. Chem. 351, 272 (1970). 21. L. E. Flanders, J. R. Bamburg, and H. J. Sallach, BBA 242, 566 (1971). 22. M. D. Prager and W. R. Whigham, BBA 132, 181 (1967). 23. K. Ibsen, K. Schiller, and E. A. Venn-Watson, A B B 128, 583 (1968). 24. R. D. Koler and P. Vanbellinghen, Advan. Enzyme Regul. 6, 127 (1968). 25. J. C. Dubin and S. Bernard, Bull. SOC.Chim. Biol. 52, 659 (1970). 26. G. E. J. Staal, J. F. Koster, H. Kamp, L. Van Milligen-Bocrsmn. and C. Veeger, BBA 227, 86 (1971). 27. K. W. Jacobson and J. A. Black, JBC 246, 5504 (1971). 27a. K. Imamura, K. Taniuchi, and T. Tanaka, J. Biochem. ( T o k y o ) 72, 1001 ( 1972). 28. M. Ide, A B B 140, 408 (1970). 29. F. W. Tuominen and R. W. Bernlohr, JBC 246, 1733 (1971).

11. PYRUVATE

357

KINASE

tially purified preparations from various sources, which have been reported since the last review, are given in Table I (61,SO-SO). This does not include those sources from which crude extracts have been prepared. It also may be worth noting that a procedure has been given for the covalent attachment of the rabbit muscle enzyme to Whatman No. 1

TABLE I PARTIALLY PURIFIED PREPARATIONS OF PYRUVATE KINASE Source (tissue)

Ref.

Escherichia wli B E . wli K12 Acetobacter xylinum Azotobacter vinelandii Bacillus subtilis Thiobacillus neapolitanus Brevibacterium favunz Euglena gracilis Mucor rouxii Coprinus lagopus Trypanosoma brucei Loach (embryo)

30 31 3% 33 34 36

36 37 38 %I

40

Source (tissue) Ascites tumor, Yoshida Ascites tumor, Ehrlich Desert locust (fat body, flight muscle) Oyster (mantle, adductor) Alaskan king crab Rainbow trout, antarctic fish ( T . bernacchii)(muscle) Rana pipiens (heart) Rat (adipose tissue) Rat (kidney)

Ref. 42 43

44 46 46 47

21 48, 49 60

41

30. P. Maeba and B. D. Sanwal, JBC 243, 448 (1968). 31. M. Malcovati and H. L. Elomberg, BBA 178, 420 (1969). 32. M. Benzimnn. BJ 112, 631 (1969). 33. C-L. Liao and D. E. Atkinson, J. Bacteriol. 106, 37 (1971). 34. M. Diesterhaft and E. Freese, BBA 268, 373 (1972). 35. A. Cornish and E. Johnson, ABB 142, 584 (1971). 36. H. Ozaki and I. Shiio, J. Biochem. ( T o k y o ) 66, 297 (1969). 37. E. Ohrmann. Arch. Mikrobiol. 67, 273 (1969). 38. S. Passeron and E. Roselino, FEBS L e t t . 18, 9 (1971). 39. G. R . Stewart and D. Moore, J. Gen. Microbial. 66, 361 (1971). 40. I. W. Flynn and I. B. R. Bowman, Trana. Roy. SOC. Trop. M e d . H y g . 65, 255 (1971). 41. L. S. Milman and Yu. G. Yurowitzki, BBA 146, 301 (1967). 42. G. Xeri, A. Coputo, and T. Terranova, Life Sci. 9 (pt 2),507 (1970). 43. F. L. Bygrave, BJ 101, 488 (1966). 44. E. Bailey and P. Walker, BJ 111, 359 (1969). 45. T. Mustafa and P. Hochachka, JBC 246, 3196 (1971). 46. G. Somero, BJ 114, 237 (1969). 47. G. Somero and P. Horhachka, BJ 110, 395 (1968). 48. C. I. Pogson, BJ 110, 67 (1968). 49. R. Marco, J. Carbonell, and P. Llorente, BBRC 43, 126 (1971). 50. 1,. Jimenez de Asua, E. Rozengurt, and H. Carminatti, FEBS L e t t . 14, 22 (1971).

358

F. J. KAYNE

filter paper disks ( 5 1 ) .The resulting immobilized enzyme exhibited some loss of long-term stability although the apparent Kmfor ADP remained unchanged. B. COMPOSITION Amino acid analyses have only been reported for the enzyme from rabbit muscle (52) and yeast (13, 1 4 ) . Substantial differences are only seen in the lower relative number of methionine and histidine and increased percentage of tyrosine residues in the yeast enzyme compared with the muscle enzyme. Cottam et ul. (52) have reported the N-terminal amino acid of the rabbit muscle enzyme to be acetylated, as have Bornmann et al. (5%) for the yeast enzyme. These workers have identified valine as the C-terminal amino acid. No covalently bound prosthetic groups or nonamino acid components have been demonstrated in any of the enzyme preparations. The extinction coefficient given for the muscle enzyme as A,,, = 0.54 for 1 mg/ml protein solution, 1 cm path ( I ) , has been generally accepted and is confirmed by the relatively low aromatic amino acid content.

C. STRUCTURE At the time of the original review, the enzyme from rabbit muscle was reported to have a molecular weight of 237,000 daltons and consist of a t least two subunits. This value for the molecular weight of the native protein has been confirmed by equilibrium as well as velocity sedimentation measurements (51) and the native enzyme is clearly dissociated in 6 M guanidine HCl to monomers of 57,000 daltons (52,53). These studies have given evidence for a preferential dissociation into dimers a t intermediate concentrations of urea and that these dimers are active. However, there is the possibility that reassociation occurs in the presence of both substrates used in the assay. Reassociation of the monomers to the active tetramer has been accomplished by dialysis, gel filtration, or dilution of the dissociating agent (52, 54). Although the idea of active dimers 51. R. J. H. Wilson, G. Kay, and M. D. Lilly, BJ 109, 137 (1968). 52. G. L. Cottam, P. Hollenberg, and M. J. Coon, JBC 244, 1481 (1969). 52s. L. Bornmann, P. Roschlau, and B. Hess, Hoppe-Seyler’s 2. Physiol. Chem. 353, 696 (1972). 53. M. A. Steinmete and W. C. Deal, Jr., Biochemiatry 5, 1399 (1966). 54. G. S. Johnson, M. S. Kayne, and W. C. Deal, Jr., Biochemistry 8, 2455 (1969).

11.

PYRUVATE KINASE

359

could suggest somc! subunit asymmetry, the one report of peptide niapping experiments (52) has shown only that iiumbcr of peptides expected for identical monomers by tryptic digestion of the protein. Certainly, there is no evidence for differing subunit molecular weights. Although such a large protein represents a formidable task a t tlie present time for the X-ray analysis of its structure, a few laboratories have made preliminary studies of the rabbit (55, 56) and human (57) muscle enzymes. There is, however, disagrecnicnt as to the size of tlic unit cell although if that reported by RlcPlierson and Rich (56) is correct, a complete structural determination may bc feasible in tlie not-toodistant future. The preliminary data on tlic human musclc enzyme arc indicative of identical subunits. Structural studies on the enzyme from othrr sources have just begun to appear in the literature. A nuinher of laboratorics haw rcportcd the molecular weights for the yeast cnzymc to bc around 160,000-190,OOO daltons (58, 591. Current opinion considers the cnzymc as being coniposed of four identical (5%) mononicrs with a niolccular wciglit of 40,000-50,OOO daltons. These values are supported by the recent calculations made from the low-anglc X-ray diffraction pattern (600).The tetramer is rcportccl to bc soinewliat asymmctric with a radius of gyration of 43.5 A. Sucrose density gradient centrifugation of thc Escherichia coli enzyme (50) indicates a molecular weight of 100,OOO daltons for the active species with dissociation into units of approxiinatcly half that size in the absence of reducing agent. Thc crythrocyte enzyme has recently been reported to have a molccular weight of 195,000 daltons (15) while that from rat liver and musclc was rcportcd as 208,000 and 250,000 daltons, respectively, by Tanaka et al. (16)on the basis of equilibrium sedimentation mcasurements. A recent preparation from pig liver was characterized as having a molecular weight of 265,000 tlaltons by gel filtration with subunits of 62,000 as determined by electrophoresis with sodium dodecyl sulfate (200).No pcptide mapping or other structural studies have hcen reported on thc enzymes from other sources. 55. P. Hollenberg, M. Flnshner, nnd M. J. Coon, JBC 246, 946 (1971). 56. A. McPherson, Jr. and A. Rich. JBC 247, 1334 (1972).

57. J. W. Campbcll, E. Due&, G . Hodgaon, W. D. Mercer, D. K. Stnnimers, P. L. Wendell, H. Muirhead, and H. C. Wntson, Cold Spring Harbor Symp. @ant. BWZ. 36, 165 (1971). 58. H. Bischofbergcr, I3. Hess, and P. Rosclllnu, Iloppe-Seyler’s 2. Physiol. Chem. 352, 1139 (1971). 59. R. Kuczenski and C. H. Snclter, Biochemistry 9, 2043 (1970). 60. K. Miiller, 0. Kmtky, P. RBschlau, nnd B. Hess, Hoppe-Seyler’s Z . Physiol. Chem. 353, 803 (1972).

360

F. J . KAYNE

D. CHEMICAL MODIFICATION Muscle pyruvate kinase has long been known to react with the usual sulfhydryl reagents ( I ) , but the -SH groups are not unusually reactive and there is no direct 1:1 stoichiometry of -SH groups reacted with inhibition of catalysis. More recent reports of -SH reactivity under various conditions have been given by Mildvan, Leigh, and Cohn who used the kinetic protection method to determine activator (61) and substrate (62) dissociation constants with the muscle enzyme. Tanaka et al. (16) found that the rat liver enzyme was 30 times more sensitive to p-hydroxylmercuribenzoate inhibition than the rat muscle enzyme. Johnson and Deal (63) have reported that pyridoxal 5’-phosphate forms a Schiff base with the c-NH, group of a lysine residue of the muscle enzyme which results in an apparent inhibition (reversible) of the activity. The rate of this interaction is decreased in the presence of ADP, ATP, or P-enolpyruvate, but to a relatively small extent. This inhibition by pyridoxal 5’-phosphate was observed earlier in a survey of glycolytic enzymes (64). Similarly, Hollenberg e t al. (55) have shown that treatment with trinitrobenzene sulfonate causes trinitrophenylation of the r-NH, group of a lysine which apparently blocks a portion of the nucleotide binding site. This inactivation is prevented by the presence of ADP, MgZ+,or K+. Reaction of the same reagent with the yeast enzyme (64a) shows a similar effect, with four lysine residues reacting with biphasic kinetics. In this case, inactivation is only around 60% and is likewise prevented by ADP or Mg”. Flashner and Coon (65) have made a preliminary study of the reaction of dithiobisnitrobenzoate with the rabbit muscle enzyme and observed four reactive sulfhydryl groups in the presence of P-enolpyruvate, K+ and Mg2+. The enzyme thus reacted is active unless the substrate and metals are removed a t which time inactivation apparently occurs with an interchange of disulfide groups. A similar observation was also made earlier by Bondar and Pon who showed 4 of the approximately 34 cysteine residues reacted with this reagent in the presence of 61. A. S. Mildvan and R. A. Leigh, BBA 89, 393 (1964). 62. A. S. Mildvan and M. Cohn, JBC 241, 1178 (1966). 63. G. Johnson and W. C. Deal, Jr., JBC 245, 238 (1970). 64. W. Domschke and G. Domagk, Hoppe-Seyler’s Z . Physiol. Chem. 350, 1111 (1969). 64a. P. Roschlau and B. Hew. Hoppe-Seyler’s Z . Physiol. Chem. 353, 914 (1972). 65. M. Flashner and M. J. Coon, Asbtr. 162nd ACS Meet., Washington Biol. No. 53 (1971).

11.

PYRUVATE KINASE

36 1

Mg2+ without causing inhibition (66). Bondar and Suelter (67) have made a preliminary study of this reaction with the yeast enzyme and found a biphasic behavior of the reaction kinetics. The rates of activity loss were influenced by the presence of metal ion cofactors, substrates, and fructose 1,6-diphosphate. The dithionitrobenzoate reactions with the yeast enzyme have also been described by Wieker and Hess (68) who have distinguished three cysteine residues on the basis of their different reactivities and protection from reaction by substrates and divalent and allosteric activators. They also conclude, as with the rabbit muscle enzyme, that substitution of one of the groups occurs via an intermediate enzyme-dithionitrobenzoate complex. Jacobson and Black (27) found no significant difference in the rate of iodoacetamide reaction between the rabbit muscle and erythrocyte enzymes. Significant protection was observed in the former enzyme with K+ or Mg2+-ADPand in the latter, with P-enolpyruvate-K+ or P-enolpyruvate-Mg*+-ADP. In somewhat related work, Flynn and Bowman (40) have shown that inhibition of a trypanosome pyruvate kinase by arsenicals is competitive with respect to P-enolpyruvate and noncompetitive with respect to ADP. Actually, relatively little has been learned from chemical modification experiments on this enzyme up to the present time. This suggests the absence of particularly reactive groups in the vicinity of the active site as well as limited exposure of groups critical to the maintenance of the active conformation of the enzyme.

E. CONFORMATIONAL CHANGE Pyruvat,e kinase was one of the first enzymes for which a conformational change was detected by ultraviolet difference spectroscopy upon binding specific ligands (69). I n fact, similar tryptophan difference spectra were observed upon binding monovalent or divalent cations or changing the temperature of a solution of the enzyme (70-7s). I n gen66. R. J. L. Bondar and N. Pon, Abstr., 156th A C S Meet., Atlantic City, Biol. No. 172 (1968). 67. R. J. L. Bondar and C. H. Suelter, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 30, 1104 (1971). 68. H.J. Wieker and B. Hess, Hoppe-Seyler's Z . Physiol. Chem. 353, 769 (1972). 69. C. H. Suelter and W. Melander, JBC 238, PC4108 (1963). 70. F. J. Kayne and C. H. Suelter, JACS 87, 897 (1965). 71. F. J. Kayne and C. H. Suelter, Biochemistry 7 , 1678 (1968). 72. R. H. Wilson, H. J. Evans, and R. R. Becker, JBC 242, 3825 (1967). 73. J. Melchoir and C. Davis, Fed. Proc., Fed. Amer. Soc. Ezp. Biol. 26, 832 (1967).

362

F. J. KAYNE

eral, these difference spectra were typical of the solvent-perturbed spectra of the aromatic amino acid residues; hence, a change in their solvent environment was suggested. Similar conclusions were also reached in studies of tryptophan fluorescence (74) and the ultraviolet circular dichroism spectrum (75). Evidence from sedimentation velocity and optical rotatory dispersion studies (71) suggested, however, that any protein conformational changes were relatively small and did not markedly alter these structural properties. The thermodynamic data obtained from equilibrium studies of the conformational changes could apparently be described adequately by a model in which the enzyme existed in two conformations, the equilibrium between which were strongly temperature-dependent ( 7 1 ) . A subsequent kinetic study by the temperature-jump technique indicated more than three distinct relaxation times occurred during the transition ; hence, intermediate conformational states were populated to a significant degree (76). The number of conformationally related relaxation times is somewhat surprising but might be related to the conformational linkage between subunits in this tetrameric enzyme. The relaxation times for these conformational changes were in the range of 10 msec to l min. Other workers (77) have suggested that a conformationally related relaxation time around 0.1 msec exists, although this was not seen in the former study. Other direct evidence has been found suggesting that the muscle enzyme exists in more than one conformational state. Reed and Cohn (78) have seen such indications in the presence of both isotropic and anisotropic Mn2+ EPR signals for the Mnz+-pyruvate kinase complex in the presence of various substrates and inhibitors with the weakly activating monovalent cations. In this case, signals from enzyme-bound Mn2+ are seen which can only arise from the simultaneous presence of differently liganded Mn2+species. Kuczenski and Suelter have extended similar conformational studies to the pyruvate kinase from yeast (79).Small changes (ca. 2 3 % ) in tryptophan fluorescence are seen upon titrating the enzyme with monovalent or divalent cations. However, a much larger (about 127%) quenching is seen upon binding the allosteric activator fructose 1,6-diphosphate 74. C. H. Suelter, Biochemistry 6, 418 (1967). 75. R. A. Wildes, H. J. Evans, and R. R. Becker, BBA 220, 850 (1971). 76. F. J. Kayne, in ‘‘Probes of Structure and Function of Mncromolccnlcs and Membranes” (B. Chancc, T. Yonctnni, and A. S. Mildvnn. rds.), Vol. 2, p. 147. Academic Press, New York, 1971. 77. G. G. Hammes and J. Simplico, BBA 212, 428 (1970). 78. G. H. Reed and M. Cohn, Fed. Proc., Fed. Amer. SOC.B x p B i d . 31, 451 (1972). 79. R. Kuczenski and C. H. Suelter, Biochemistry 10, 2862 (1971)

11.

PYRUVATE KINASE

363

(FDP). Binding constants have been obtained for this ligand under a number of conditions of substrate and activator concentrations. This enzyme was recently recognized as being cold sensitive (80); upon incubation a t 0”, activity is lost over a period of tens of hours in a biphasic manner. This loss is accelerated as much as 1000-fold by the allosteric effector FDP and inhibited by addition of divalent activators or cryoprotectants such as glycerol and dimethylsulfoxide (81).This is apparently the basis for the stabilization by glycerol during the preparation. A somewhat similar case may arise with enzyme as isolated from rat adipose tissue by Pogson (48, 82). Separation and isolation of two kinetically different forms of the enzyme can be achieved by extraction in the presence and absence of EDTA. The interconversion from the form exhibiting cooperative interactions with P-enolpyruvate into that showing Michaelis-Menten kinetics with P-enolpyruvate is mediated by FDP, and the reverse by EDTA, ATP, or citrate. The effccts of varying temperature on these experiments are not known. Somero and Hochachka (&’, 47) have studied the enzyme from the Alaskan king crab and fishes of different thermal environmental require-. ments. Their results indicate a similar, but temperature-dependent, interconversion between two forms of the king crab enzyme. I n this case, the forms arc diffcrentiatcd by their kinetic characteristics. The “cold” variant exhibits hyperbolic kinetics with P-enolpyruvate and a minimum & a t 5 ” , while the “warm” variant has sigmoid kinetics and a minimum K , at 12”. Actually, few other studies have been made of the temperature dependence of the kinetic parameters for the enzymes from various sources. While apparent Michaelis constants for the substrates of the muscle enzyme may not be very temperature dependent, an Arrhenius plot of the catalytic reaction velocity does show a curvature and this has been interpreted to have some causal relationship to the equilibrium between the forms of the enzyme observed by ultraviolet diffcrence spectroscopy (71). The significance of these various thermal effects is not simply understood. Based on our present knowledge the only statement which can be unequivocally made is simply that these effects are seen in the enzymes isolated from different sources, and that these effects can usually be related to an equilibrium involving some ligand binding which in turn may havc some physiological or control significance. The similarities may be quite fortuitous and their presence reflects only the fact that 80. R. Kuczenski and C.H. Suelter, Biochemistry 9, 939 (1970). 81. M. J. Ruwnrt and C . H. Sueltcr, JBG 246, 5990 (1971). 82. C. I. Pogson, BBRC 30, 297 (1968).

364

F. J. KAYNE

multiple conformations do exist and are probably involved in the regulatory functions of the protein.

111. Catalytic Properties

A. STOICHIOMETRY AND SPECIFICITY 1 . Reactions Catalyzed

In addition to the normal reaction, e.g., in the glycolytic pathway ( I ) , pyruvate kinase from muscle also catalyzes the phosphorylation of fluoride or hydroxylamine from ATP [see Boyer (1)1. Both of these reactions require bicarbonate as a cofactor, are not apparently reversible, and exhibit somewhat different divalent metal requirements from the normal reaction. The only subsequent studies published, those by Cottam et al. (85), demonstrated the product of the hydroxylamine reaction to be O-phosphorylhydroxylamine.The product of the other reaction, fluorophosphate, a competitive inhibitor of the normal reaction with respect to P-enolpyruvate, has been used in the interpretation of the structure of the components at the active site of the enzyme (84, 86). Proton exchange at the methyl group of pyruvate is also catalyzed by the enzyme (86),and this activity has been used by Robinson and Rose (87) in isotope exchange studies of the catalytic reaction. 2. Substrate Specificity

Early work on the enzyme, reviewed by Boyer ( I ) , demonstrated a somewhat broad specificity for the nucleotide substrate of the reaction. This was further extended by the studies of Plowman and Krall (88) who determined specific activities and apparent Michaelis constants for several diphosphates. The maximum velocities at pH 7.5 were in the order: ADP N GDP > IDP > d ADP > U D P > C D P > dCDP; with CDP ca. 30% and dCDP 0.3% that of ADP. However, the K , for ADP was a factor of four smaller than for GDP or any of the others. In addition, the same order of relative rates was also found in studies using 83. 84. 85. 86. 87. 88.

G. L. Cottarn, F. Kupiecki, and M. J. Coon, JBC 243, 1630 (1968). A. S. Mildvan, J. S. Leigh, and M. Cohn, Biochemistry 6, 1805 (1967). T. Nowak and A. S. Mildvan, Biochemistry 11, 2819 (1972). I. A. Rose, JBC 235, 1170 (1960). J. L. Robinson and I. A. Rose, JBC 247, 1096 (1972). K. M. Plowman and A. R. Krall, Biochemistry 4, 2809 (1965).

11. PYRUVATE

KINASE

365

crude extracts of the brain enzyme (89). This suggests that the nucleotide binding portion of the active site does not have very stringent steric (or catalytic) requirements. The phosphate ester portion, as might be expected, does show some structural restrictions. The only work reported on such modifications concerns studies of the rabbit muscle enzyme with which Setondji et al. (90)demonstrated that adenosine 5'-hypophosphate could act as phosphoryl acceptor from PEP, resulting in the formation of adenosine 5'-hypophosphate phosphate (AOPPOP). The Ki reported for this substrate, in the presence of ADP, was 1.8 X 10-2M while the apparent K,. was 1.7 x M. Until recently few studies were done with P-enolpyruvate analogs, apparently because none was readily available. Four groups (91-95) have synthesized a number of analogs and found that P-enol-a-ketobutyrate (91-93, 95), P-enol-a-ketovalerate (93, 95), P-enol-a-ketocaproate (96), P-enol-3-Br-pyruvate (96), and P-enol-3-F-pyruvate (94) could act as poor substrates with maximal velocities as phosphoryl donors less than 1s/. that of P-enolpyruvate. Apparent Michaelis constants were not, however, very much greater than that for the normal substrate. A number of other analogs were found to serve as weak inhibitors or were completely inactive. I n the studies of hydroxylamine phosphorylation, Cottam et al. (83) found that both N-methylhydroxylamine and, to a much lesser extent, N,N-dimethylhydroxylamine could substitute as the acceptor for the ATP phosphate. Actually, these results indicated quite a high degree of specificity toward the phosphate donor component of the reaction. There is no carbon asymmetry in the enolpyruvate or pyruvate, but the introduction of deuterium and tritium into the molecule allows a determination of any stereospecificity in the reaction, Indeed, Rose (96) has done just this and the results have been confirmed by Bondinell and Sprinson (91) and Stubbe and Kenyon (9,9), both of whom studied the stereochemistry of the product of the reaction using P-enol-a-ketobutyrate as the substrate. The proton addition to P-enolpyruvate or z-P-enol-a-ketobutyrate occurs from a specific direction, i.e., the 2-si face (3-re of the P-enol-a-ketobutyrate) of the molecule : 89. G. Weber, Proc. N a t . Acad. Sci. U . 8. 63, 1365 (1969).

90.J. Setondji, P. Remy, J-P. Ebel, and G. Dirheimer. BBA 232, 585 (1971). W. Bondinell and D. Sprinson, BBRC 40, 1464 (1970). J. Stubbe and G. Kenyon, Biochemistry 10, 2669 (1971). H. D. Soling, U. Walter, H. Sauer, and J. Kleineke, FEBS L e t t . 19, 139 (1971). J. Stubbe and G. Kenyon, Biochemistry 11, 338 (1972). A . E. Woods, V. B. Chatman, and R. A. Clark, BBRC 48, 1 (1972). I. A. Rose, JBC 245, 6052 (1970).

91. 92. 93. 94. 95. 96.

366

F. J. KAYNE

H+

One preliminary report has been made which suggests that this face may be directly accessible to the solvent (97).In this case, the enzyme apparently catalyzed a stercospecific reduction of pyruvate by NaBH,, forming 73% of the D-isomer of lactate. 3. Cofactors

Pyruvate kinase was the first enzyme for which an absolute requirement for a monovalent cation was established [see reviews by Boyer ( I ) , Suelter (98),and Evans and Sorger ( 9 9 ) ] .The one well-documented exception to this requirement is the enzyme from Acetobacter xylinum (32) where no such K' or NH,' requirement was found. Other reports have been given of the inability of the monovalent ion to stimulate the reaction catalyzed by the enzymes from E . coli (SO), A . vinelandii ( S S ) , B. flavum ($69,and a plant latex (100). Various cations can serve as the required activator, and the enzyme usually shows an apparent preference (in terms of V,,,,,) for those of a particular ionic size. Table I1 illustrates this in the case of the rabbit muscle enzyme, where in the presence of tris+ or (CH,),N+ the activity is less than 0.02% that in the presence of K+. Concentrations under 0.1 M of these ions do not significantly inhibit the activated velocities, but the indicated V,,, should be considered only an apparent value. The actual part played by the monovalent ion in enzyme catalysis has been a puzzle since it was first recognized. One of the earliest suggestions, by Kachmar and Boyer (101), that perhaps the cation serves to regulate the critical displacement of an enzyme group involved in catalysis is still quite probable. Suggestions of a unique role in a larger conformational aspect of the protein are made somewhat unlikely by the observations that those conformational changes seen can be caused by other changes in solution conditions. Thallium (I) is a good substitute for K+ in the catalytic reaction (a), and a study of the nuclear mag97. T. M. Phillips and G . W. Kosicki, Fed. Proc., Fed. Amer. Soc. Em. Biol. 29, 462 (1970). 98. C . H. Suelter, Science 168, 789 (1970). 99. H.J. Evans and G.J. Sorger, Annu. Rev. Plant Physiol. 17,47 (1966). 100. J-L.Jacob and J. d'Auaac, Bull. SOC.Chim. Biol. 51, 511 (1969). 101. J. F. Knchmar and P. D. Boyer, JBC 200, 069 (1953).

11.

367

PYRUVATE KINASE

TABLE I1 OF PYItUV.lTIs; KIN.W I k i C T l O N 1IICLITIVE ACTIVATION VARIOUS MONOVALENT CATIONS.

Cation

Li+ Na+ K+ NHI+ ltb+

T1+

cs+

Crystal radius

V,,, (cation)c/

(A)b

Vmax(K+)

0.68

0.97 1.33 1.43 1.47 1.47 1.67

BY

Optimum cation conc. (mW

0.02 0.08 1.00 0.81 0.65 0.61 0.09

100 100

100 50 100 3 100

From Kayne (8),measured with C1- as anion; M$+, 8 mdl. Value from “Handbook of Chemistry and Physics,” 48th ed. (1967). Maximal specific activity with constant ionic strength, about 0.15 M.

netic resonance of the 205Tl+bound to the enzyme has been made (109, 10s). Paramagnetic broadening of the resonancc signal by the Mn2+spin was used to estimate the monovalent-divalent cation separation distance and the values obtained were 8.2 and 4.7A in the absence and presence of P-enolpyruvate respectively. Although the absolute values are subject to some uncertainty, the rclative differcnce is quite significant, and a substantial conformational change is therefore indicated upon the addition of substrate. In any case, the monovalent cation is clearly located very near the active site, and this lends support to the postulation by Suelter (98) of a catalytic role (more than simple charge neutralization) for the metal. Of course, if such were the case, one would have to postulate that those enzyme variants not requiring a monovalent cation would operate by supplying some needed catalytic function to the transition state through an enzyme functional group. Nowak and Mildvan (85) have made thc suggestion that the monovalent cation acts by coordination of the substrate (P-enolpyruvate or analog such as P-lactate or P-glycolate) a t the carboxyl group. This conclusion was reached through a study of the proton relaxation rates of H 2 0 in the Mn2+ coordination sphere and paramagnetic broadening of the slP and ‘H nuclei of such substrates and analogs. While the experimental evidencc is consistent with such a coordination, there is, unfortunately, no dircct proof bccausc of an ambiguity which arises here 102. F. J. Knyne nnd J. Reuben, JACS 92, 220 (1970). 103. J. Reuben and F. J. KaJ-nc, JBC 246, 6?27 (1971).

368

F. J . KATNE

a5 in many such systems. In this case, K+ is shown to raise the affinity of the enzyme-Mn2+ complex for the substrate or analog (and vice versa), except in cases where the carboxyl group is blocked. Unfortunately, we do not know if the K+ only allows an enzyme liganding group to interact with the carboxyl group. Another study of this cofactor binding site was made by Bryant (104) who looked a t the aoK+NMR. Howcvcr, in this case, quite high K+ concentrations were required for thc mcasurcincnt (2 M ) and the results are therefore somewhat ambiguous because of the uncertain effects of such high ionic strengths. The author did report that the enzyme broadened the 3DK+signal under those conditions, and this effect was not substantially changed in the presence of Mg2+-ATP but reduced in the presence of Mn2+plus Mg?+-ATP. The latter observation would be consistent with what is apparently a morc effective competition of the Mn2+ for the monovalent site (8). The enzyme from all sources also requires a divalent cation for activity. The early kinetic work on this requiremcnt indicated that the nucleotide substrates could interact with the enzymc as the divalent metal salts (1, 105) and also suggested a random order of metal-nucleotide addition was most fcasiblc. Nuclear magnetic resonancc studies by M. Cohn's group (62, 84, 106) demonstrated thc likelihood of the involvement of an enzyme-mctal-substrate bridgc complex a t the active site with a random order of metal and substrate addition. On kinetic grounds, Clcland (107)has questioned this suggcstion, and has proposed that the metal acts only to complex the nucleotide. A strong argument against this, howcver, is tlic fact that both ADP and ATP bind to the muscle enzyme in the absence of any divalent cation (108) (even with only (CH,),N+ present) with affinities not much less (ca. 1 mM) than the reported substrate I(, or product K i valucs. As far as the environment of the enzyme-bound divalent cation is concerned, Reuben and Cohn (109) studied this in detail by observing the temperature and frequency dependence of the H,O proton relaxation rates in the presence of Mn2+. Analysis of these factors suggested the bound Mn2+has three water molecules coordinated, and the conclusion was that the enzyme prohably provides thrce ligands in thc binary complex. A determination of Mil2+ binding by E P R showed three to four

104. R. G. Bryant, BBRC 40, 1162 (1970). 105. J. B. Melclioir, Biochemistry 4, 1518 (1965). 106. A. S. Mildvan and M. Colin JBC 240, 238 (1965). 107. W. W. Cleland. Annu. Rev. Biochem. 36, 77 (1967). 108. N. C. Price, FBBS Lett. 24, 21 (1972). 109. J. Reubcn and M. Colin, JBC 245, 6539 (1970).

11. PYRUVATE

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sites per protein tetramer. Cottam and Mildvan (110) studied the binding of the Mn2+to pyruvate kinase by proton relaxation rate and electron paramagnetic resonance measurements. They found that addition of urea to the enzyme, sufficient to give rise of indications of unfolding or dissociation into dimers, resulted in an increase in the apparent Mn2+ dissociation constant but no change in the number of cations bound. However, with an older preparation of low specific activity, a decrease in the number of sites with no apparent change in K D for Mn2+was seen. Similar studies of the yeast enzyme have also been carried out in Mildvan’s laboratory (111).Basically, these have indicated a similar coordination of the Mn2+in the ternary complexes as compared with the muscle enzyme. Fructose 1,6-diphosphate has little effect on this, only decreasing the apparent substrate dissociation constant about 3-fold. A greater difference in behavior was seen, however, with ATP and ADP in the ternary complexes where their dissociation constants were about an order of magnitude smaller than their respective inhibitor or Michaelis constants suggesting multiple modes of binding ATP or the occurrence of a steady state mechanism, possibly not random. The recent work by Nowak and Mildvan, previously mentioned (85), has also led to the suggestion that the divaleiit cation, Mn2+in this case, binds directly to the phosphoryl groups undergoing transfer. However, there are still some questions as to the exact values of these internuclear distances, i.e., Mn2+-31P,Mn2+-lH,and Mn2+-l3C,especially in light of the observation of more than one species of bound Mn2+being present with these analogs. One can say with certainty, though, that the enzyme donates some ligands (69) and this cation is also near the active site itself. The failure to find a typical tetrahedral-type absorption spectrum for the CO‘+complex of the enzyme further suggests the existence of an octahedral environment for the divalent cation (106). The identity of these ligaiiding groups is unknown, just as in the case of the monovalent cation. A few divalent metal ions can substitute for the apparent physiological activator, Mgz+.Boyer (1) mentioned some of these along with the differences in metal requirement for the “hydroxylamine kinase” and “fluorokinase” reactions. More recently, Bygrave (@) has studied the metal ion requirements of the tumor and muscle enzyme and shown that Ni2+will activate to a small degree along with the known substitutions of Mil2+or Co’+. Apparently, Zn2+does not activate the pyruvate kinase 110. G. L. Cottam and A. S. Mildvan, JBC 246, 4363 (1971). 111. G. I,. Cottam, A. S. Miidvan, J. R. Hunsley, and C. H. Suelter, JBC 3802 (1972).

247,

370

F. J. KAYNE

reaction but Cd2+will to some 13% of the Mg2+velocity, although it apparently will slowly inactivate the enzyme. In some cases, Mn2+has been reported to be the preferential activator. This was seen in a study of the enzyme from ascites tumor cells (@), in the rabbit muscle enzyme with the substitution of T1+for K+, and with the enzyme from Mucor rowhi in thc absence of FDP (112). The latter report showed considerable differences in the kinetic behavior of the enzyme in the presence of either of these divalent activators, a situation which was also seen in a highly purified preparation of the B. licheniforinis enzyme (113) where AMP reversal of the ATP inhibition was not seen with Mn2+.Such an observation strongly suggests a multiple role for the divalent activator, both in thc catalytic function as well as in the protein conformational equilibria related to the regulatory function.

4. Number of Active Sites During the early part of the time covered by this review, most workers used the value of two as the number of active sites for the muscle enzyme. For the most part, this was based on the work of Reynard et al. (114) in a measurement of the number of P-enolpyruvate binding sites by ultracentrifugation, which was reported to be 2.8 while a value of 2-4 was given for the number of pyruvate binding sites. With the evidence for the number of subunits being four as discussed earlier, additional studies have been made by workers who looked into this a little more closely, and thc findings are summnrizcd in Table 111. It TABLE I11 NUMDER

OF

BINDING SITES

IN l
KIN.WIC

Moles per 237,000 g

Ligand

Method

Ref.

4.2 f 0.2 3 to 4 4 . 3 f 0.7 2 to 4 4.0 3 .8 f 0 . 3 4.5 f 0 . 8

Zn2f Mn2+ Mn2+ Pyridoxal5’-POd (NOz)s-benzenesulfonate P-enolpyruvate Tl+

~~C~NMR EPR EPR Absorption spect,riim Absorption spectrum Equilibrium dialysis Equilibrium dialysis

116 109 110 6.3

66 8 8

112. S. Passeron and H. Tcrenzi, FEBS Lett. 6, 213 (1970). 113. F. W. Tuominen niid R. W. Bernlohr, JBC 246, 1746 (1971). 114. A. M. Reynard, L. F. Hass, D. D. Jacobson, nnd P. D. Boycr. JBC 236, 2277 (1961). 115. G. L. Cottain and R. Ward, ABB 132, 308 (1969).

11. PYRUVATE

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should be pointed out that these results are quite dependent on the particular conditions, and, indeed, Melchoir and Kamper (116) have claimed to observe up to 12 moles of Mgz+bound a t low K+ (0.025M) concentrations by the use of gel filtration techniques.

B. ASSAY Enzymic activity is usually measured continuously by coupling the pyruvate formation to the lactic dehydrogenase reaction (NADH utilization) or stopping the reaction and estimating pyruvate formation with 2,4-dinitrophenylhydrazine [see Boyer (1)] . More recently, workers have used pH-stat (70, 105) or pH change (11‘7) recording for following the H+ uptake, the disappearance of the 240-nm absorption of the en01 phosphate (118), and the use of 32P-labeled nucleotide substrate with subsequent thin-layer chromatographic separation of the products and determination of the incorporated label (119). Although the coupled lactic dehydrogenase assay is the quickest and most convenient, each of these other methods has obvious uses, for example, where it is desirable to eliminate any problems resulting from the presence of a coupling enzyme as with the first two methods and in the assay of crude preparations with the latter. Obviously, suitable precautions have to be taken in the application of any assay for it to be valid for a particular purpose. One cautionary note concerns the substrate levels utilired in the popular Bucher and Pfleiderer (120) assay, particularly the 0.2 mM ADP concentration, which is just about its I(, value. At least ten times this level would normally be used if some other variable is being studied. Of course, it is always a good procedure to check the optimal activator concentration levels when modifying the assay for any purpose or studying the enzyme from a source for which these characteristics are not well documented.

C. THERMODYNAMICS The equilibrium characteristics of the pyruvate kinase reaction were discussed by Boyer ( I ) in the previous review. No further studies have 116. J. B. Mrlchoir and D. Knmpcr, Ferl. Proc., Ferl. Anzer. Soc. E x p . B i d . 31, 886 (1972). 117. C. Carmcli and Y. Lifshitz, Annl. Biochem. 38, 309 (1970). 118. N. G. Pon and R. J. L. Bondar, Annl. Biochem. 19, 272 (1967). 119. J. Morningsinr, Jr. and D. Therriault, Annl. Biochem. 39, 135 (1971). 120. T.Biichcr and G. Pfleiderer, “Methods in Enzymology,” Vol. 1, p. 435, 1955.

372

F. J . KAYNE

apparently been done on the enzyme reaction or various binding equilibria. Since most binding constants for the various substrates products and effectors have been obtained by kinetic measurements, the observed temperature dependence may not necessarily be directly related to the changes in the thermodynamic parameters.

D. KINETICS 1. Substrates and Activators

Most relevant kinetic studies on the muscle enzyme have been discussed under previous headings of this chapter and the earlier review ( 1 ) . Obviously, some interactions exist which are somewhat more complicated than those described by the usual Michaelis-Menten formulation, especially with regard to the monovalent-divalent cation antagonisms. Slightly cooperative behavior can be observed in the binding of the monovalent (8, 121) or the divalent (121) activator under certain conditions. The slight inhibition shown by Kachmar and Boyer (101) a t monovalent concentrations above ten times their activator (Michaelis) constants which are not simply nonspecific ionic strength effects, as well as the sharp optima in divalent activator concentrations, suggest the ability of each cation to bind with a lowered affinity at the other site (8). Probably, much of the uncertainty regarding the various kinetic aspects of cation binding is related to this behavior. Very recently, it was found that L-phenylalanine could noncompetitively inhibit the muscle pyruvate kinase (122,123). The concentration dependence curve for this effect follows a classic allosteric pattern with the Hill coefficient, n, increasing from 0.85 to 2.0 between pH 7 and 8, respectively. This cooperativity was significantly reduced when enzyme was used which had been stored in dilute concentrations or reassociated from the unfolded subunits (desensitization). Apparently, the phenylalanine is not binding a t the active site but at some other specific location. Alanine, serine, and cysteine were found to reverse the effect of phenylalanine. These workers (123) mcntionecl that relatively high (millimolar) concentrations of phenylalanine were required for half-inhibition but did show that this value was dependent on the P-enolpyruvate concentration. This 121. C. H. Suelter, R. Singleton, Jr., F. J. Kayne, S. Arrington, J. Glass, and A. S. Mildvan, Biochemistry 5, 131 (1M6). 122. E. Rozengurt, 1,. Jimenea de Asua, and H. Carminatti, FEBS Lett. 11, 284 ( 1970).

123. H. Carminatti, L. Jimenez de Asua, B. Leiderman, and E. Rozengurt, JBC 246, 7284 (1971).

11. PYRUVATE

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373

is thc only prcscntly known cxamplc of allostcric interactions in the muscle enzyme although it can be argued that the monovalent-divalent interactions are allosteric in naturc since protein-mediated effects are suggested (124.6). Interest in the kinetic properties of pyruvate kinase from other sources followed from the suggestion that this cnzymc is an important point in the control of glycolysis as was discussed by Hess and Brand (125). This laboratory was the first to report a direct effect on the yeast enzyme by a feed-forward activator, fructose 1,6-diphosphate (126). Subsequent work with the purified enzynic in their laboratory (12’7) and by Hunsley and Suelter (128, 129) found it to exhibit classic allosteric kinetics of the “K” type (130). Homotropic activation with P-enolpyruvate was observed with heterotropic activation by FDP. Around the same time, similar effects were seen with the rat liver enzyme (131, I%?). Although it is difficult to generalize, most of the pyruvate kinase ensymes from nonmusclc sources show a significant degree of cooperativity in binding P-enolpyruvate (but not the nucleotide), and FDP serves as an activator for most of these. This activation occurs as an apparent shift of the P-enolpyruvate saturation curve to a hyperbolic form with a lowering of the concentration required for half-maximal velocity. It is easy to find exceptions to this generalization. In some cases, one can imagine that insufficient purification has failed to remove endogenous 124. Recent experiments in this laboratory IF. J. Kayne and N. C. Price, Biochemistry 11, 4415 (1972)l have shown that these interactions can be followed by changes in tryptophan fluorescence emission and tryptophan ultraviolet absorption spectra. Fluorescence enhancement and absorption differences opposite to those produced by adding thc cation cofactors (70) nre seen upon binding of phenylalanine. These changes arc reversed in the presence of the divalent cation (Mg’’). The.= results extend thr observations of Carminatti et d. (123) by showing a much tighter binding of phenylalanine in the absence of P-enolpyruvate. Electron paramagnetic resonance studies show a concomitant reduction of the enzyme’s affinity for Mn2+ while preliminary NMR experiments suggest the phenylalanine site to be a relatively large distance from the MnZ+sitc (>12A) and thus, most probably, allosteric in nature. 125. B. Hess and K. Brand, in “Control of Energy Metabolism” (B. Chance, R. W. Estabrook, and J. R. Williamson, cds.), p. 111. Academic Press, New York, 1965. 126. B. Hess, R . Haeckel, and K. Brand, BBRC 24, 824 (1966). 127. R. Haeckel, B. Hess, W. Lauterborn, and K-H. Wiister, Hoppe-Seyler’s Z. Physiol. Chem. 349, 699 (1968). 128. J. R. Hiinsley and C. H. Sueltcr, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 26, 559 (1967). 129. J. R. Hunsley and C. H. Suelter, JBC 244, 4819 (1969). 130. J. Monod, J. Wyman, and J-P. Chnngeux, JMB 12, 88 (1965). 131. T.Tanaka, F. Sue, and H. Morimura, BBRC 29, 444 (1967). 132. C.B. Taylor and E. Bailey, BJ 102, 32 C (1967).

374

F. J. KAYNE

FDP from the system and the resulting kinetic picture is similar to that of the muscle enzyme. In mammalian tissues, there is some evidence for isoenzymes (133), and these may have kinetic properties of the muscletype enzyme (16), and, finally, one must consider the presence of interconvertible forms with such kinetic characteristics. Since the initial reports of FDP activation of the yeast and rat liver enzymes, many enzymes of various degrees of purification from different sources have been found to exhibit such effects. These include: E . coli K12 (31), various fermentive yeasts (134), E . gracilis (37), M . rouxii (I%), C . Zagopw (39), T . brucei (do), loach embryo (41), desert locust (flight muscle, fat body) (&), oyster (45),trout, and T . bernacchii ( 4 7 ) , R. pipiens (eggs and liver) ( I % ) , rat (adipose tissue) (49, 82), rat (kidney) (60), erythrocytes (4, 26, 137, 138), and lung cells (tissue culture) (139). In general, where a metabolic regulation of the type required in the change from glycolysis to gluconeogenesis is needed, positive cooperativity in P-enolpyruvate binding is observed. The ability of FDP to activate is quite general, but one major exception is the substitution of 5’-AMP activation seen with the bacterial enzymes from B . licheniformis (113), A . vinelandii (33),B . liquefaciens (28), B . flavurn ( 3 6 ) ,E . coli B (30), and, to a slight extent, with T. neapolitanus ( 3 5 ) .3’,5’-cyclic AMP was observed to activate the enzyme from loach embryos where the FDP effect is also observed (41). Monophosphorylated hexoses have also reported to weakly activate the enzymes from erythrocytes (26), rat liver (140), and T. neapolitanus ( 3 5 ) . A recent publication has suggested that glucose l16-diphosphate can activate the erythrocyte enzyme (141) and that intracellular levels of this metabolite make control by F D P unlikely. At least two organisms exhibiting very low levels of phosphofructokinase do not show FDP activation of their pyruvate kinase: T. neapolitanw (33) and Acetobacter zylinum ( 3 2 ) . N o particular metabolic activators have been identified for the enzyme from the latter 133. J. Osterman and P. J. Fritz, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 31, 893 (1972). 134. D. S. Bnrbalace, G. H. Chambliss, and R. J. Brady, BBRC 42, 287 (1971). 135. H. Terenzi, E. Roselino, and S. Passeron, Eur. J. Biochem. 18, 342 (1971). 136. L. Schloen, J. Bamburg, and H. Sallach, BBRC 36, 823 (1969). 137. G. F. Munro, and D. R. Miller, BBA 206, 87 (1970). 138. D. E. Paglia and W. N. Valentine, Blood 37, 311 (1971). 139. G. A. Dunaway, Jr. and E. C. Smith, Life Sci. 10 (pt 2), 1163 (1971). 140. L. Eggleston and H. Woods, FEBS Lett. 6, 43 (1970). 141. J. F. Koster, R. G. Slee, G. E. J. Staal, and T. J. C. VnnBerkel, BBA 258, 763 (1972).

11. PYRUVATE

KINASE

375

organism. A point of interest here is the apparent “linkage” between the absence of an FDP effect and the absence of a monovalent cation requirement. Whether or not this has any mechanistic implications regarding the activation is not known. Unfortunately, not enough studies have been carried out to do inore than suggest the possibility of such a linkage. In all of the cases discussed thus far, the allosteric kinetics have been expressed chiefly as a positive cooperativity of the P-enolpyruvate binding. Although cooperative binding has also been reported with ADP for a few enzymes from bacteria and yeast (28, 113, 127) these allosteric interactions arc apparently weaker than those observed for Y-enolpyruvate binding. I n general most other enzymes have been reported not to show cooperative binding with the nucleotide. It is apparent from many of the studies just cited that a number of factors including endogenous substrates, cofactors, effectors, pH, ionic strength, temperature, preparation methods, and pretreatments of various conditions can strongly influence the allosteric behavior exhibited by the enzyme. When this is not recognized, especially in the cases of the partially purified enzymes, the interpretation as well as the observations regarding the kinetic behavior can bc open to question. Some of the more recent studies with the purified enzymes have involved experiments conducted using a wide range of conditions and show the complexity of these interactions. pH effects on the allosteric interactions of the yeast (12.9,142) and liver (143) enzyme, temperature effects on those of the yeast enzyme (80, 81, 144), as well as various factors influencing the enzymes from adipose tissue (48), erythrocytes ( 2 6 ) , oysters (45), and bacteria (32, 36, 113) have recently been reported. 2. Inhibitors

Inhibition of the muscle enzyme by ATP has been known for some time (I) and its competition with both P-enolpyruvate and ADP was the basis for postulating a common binding position for the phosphoryl group undergoing transfer. This observation has been challenged by the claim that the competition is the result of reduction of free Mg2+ by chelation with the nucleotide (145). Boyer hits attempted to answer these criticisms by showing ATP competitively inhibits the enzyme when the 142. 143. 144. 145.

H J . Wieker and B. Hrss, Bioclwmktry 10, 1243 (1971). E. Rozengurt, L. Jiinenea de Asua, and H. Carminatti. JBC 244, 3142 (1969). R. Kuczcnski and C . H. Suelter, Biochemktry 10, 2867 (1971). T. Wood, BBRC 31, 779 (1968).

376

F. J. KATNE

hIg2+concentration is buffered by the novel use of glycerol l-phosphateMg2+ (146). A small noncompetitive component in the inhibition with respect to P-enolpyruvate was observed a t 25" and pH 7.8 in this case, and also in the work by Holmsen and Storm (147) a t p H 8.5 and high Mg2+ concentrations. These authors apparently see ATP competition with both substrates but claim the data do not fit the equation of Reynard et al. (114). It is not clear, though, that their data do differ substantially from that reported by Boyer (146) with the exception of the significantly different K,,, values reported for P-enolpyruvate which may be a result of the different pH values used. One other piece of evidence implying that ATP is isosteric wit11 the ADP site and, as expected, not so with the P-enolpyruvate site is the studies of substrate protection against. trinitrophenylation inhibition mentioned earlier (55). These show almost identical protection by ATP and ADP, both values differing greatly from those with P-enolpyruvate. Inhibition by ATP also occurs with the pyruvate kinases exhibiting allosteric substrate kinetics. As one might expect, the kinetic picture becomes more complicated since in a number of cases it has been shown that the allosteric activator can overcome the ATP inhibition (see 26, 33, 39, 44, 113, 148). I n most cascs, ATP inhibition occurs in the millimolar range. The "liver" type of enzyme was reported to show a greater affinity for the inhibitor than the muscle type (16), but there is apparently no evidence yet that any additional binding site is involved. The only point that is somewhat surprising here is that such interaction is observed between the effector and nucleotide inhibitor while little or no interaction between effector and nucleotide substrate site is seen. As with the homotropic and heterotropic activation of P-enolpyruvate binding, these effects can be altered by changes in solution conditions. Some divalent cations can be potent inhibitors of the enzyme (1) and this is true for the allosteric rat liver enzyme where quite low concentrations of Cu2+were found to inhibit (149). This effect could partially be reversed by F D P although some irreversibly inactivated component was present (143). Of the other common metabolites tested and reported, the L-amino acids alanine and phenylalanine have been observed to also inhibit the enzyme from sources other than muscle (21, 45, 49, 90, 123, 139, 148, 146. P. D. Boyer, BBRC 34, 702 (1969). 147. H. Holmsen and E. Storm, BJ 112, 303 (1969). 148. P. Llorente, R. Marco, and A. Sols, Eur. J . Biochern. 13, 45 (1970). 149. S. Passeron, L. Jimenea de Asun, and H. Cnrminatti, BBRC 27, 33 (1967).

11.

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150-153). I n the case of rat brain (152), prostate and seminal vesicles (151),and rabbit muscle (123), the inhibition caused by phenylalanine can be reversed by alanine. With the rat adipose tissue, this can be reversed by low concentrations of F D P (49). p-Chlorophenylalanine was found to be as effective as phenylalanine with crude preparations of the rat brain enzyme (152). A number of other metabolites might serve as inhibitors of the pyruvate kinase reaction but few surveys have been conducted with purified preparations of the enzyme such as Pogson (48) has done for the rat adipose tissue preparation. Weber’s group (150) has reported that the free fatty acids myristate, laurate, and octanoate, as well as acetyl-CoA, can inhibit t.he rat liver enzyme. The latter metabolite was also shown to inhibit a crude preparation of the muscle enzyme. Oleate and palmitate were reported to inhibit the rat heart enzyme (154) but only after a moderately long preincubation period. In their review on the regulation of pyruvate kinase, Seubert and Schoner (155) discounted these first results due to the apparent irreversibility of the inhibition. Waygood and Sanwal (156) have now just reported that succinyl-CoA can reversibly inhibit the E . coli and rat liver enzymes. This observation suggests the necessary negative control function for the enzyme of this bacterium and may be related to the earlier reports of such inhibition with acetylCoA. One early report suggested that the muscle enzyme could be noncompetitively inhibited by diethylstilbesterol (157), but further studies have apparently not been made. A more recent report (158) shows that phenylethylbiguanide and related compounds inhibit pyruvate kinase like Ca*+; i.e., they are competitive with respcct to the divalent metal (106).The data also show that 4 moles of Ca2+or Mn*+ are bound per mole of tetramer (12 moles of biguanide are bound) in the presence or absence of K+. The latter observation does not correlate with Melchoir’s report (116) of up to 12 MnZ+bound in the absence of K+. 150. G. Weber, M. A. Lea, and N. B. Stamm, Advan. Enzym. Regul. 6, 101 (1968). 151. R. Vijayvargiya, W. S. Schwark, and R. L. Singhal, Can. J . Biochern. 48, 1268 (1970). 152. W. S. Schwark, R. 1,. Singhal, and G. M. Ling, Life Sci. 9 (pt l), 939 (1970). 153. W. Schoner, U. Haag, and W. Seubert, Hoppe-Seyler’s Z . Physiol. Ckem. 351, 1071 (1970). 154. E. Tsutsumi and F. Takenaka, BBA 171, 355 (1969). 155. W. Seubert and W. Schoner, Curr. Top. Cell. Regul. 3, 237 (1971). 156. E. B. Waygood and B. D. Sanwal, BBRC 48, 402 (1972). 157. D. V. Kimberg and K. L. Yielding, JBC 237, 3233 (1962). 158. F. Davidoff and S. Carr, Proc. Nut. Acad. Sci. U.S. 69, 1957 (1972).

378

F. J. KAYNE

E. CONTROL Clearly, much of the interest in the enzymology of pyruvate kinase relates to its functioning in the control of glycolysis and gluconeogenesis. Most of these aspects have been reviewed (2, 166, 169) but should be mentioned briefly here. Some time ago, Kerson e t al. (160) made a computer simulation study of the mammalian enzyme which demonstrated its kinetic functioning in glycolysis. This may have been a bit premature because of the subsequent discovery of the allosteric inhibition of the muscle enzyme (lZZ-l24). In muscle, an active pyruvate kinase is likely to be desired under most conditions and any control functions other than inhibition when ATP levels (and available energy supply) are high are not understood. This is especially true regarding any control functions for the monovalent or divalent cations since the authors (160) had pointed to the latter as being able to exert strong control on the reaction velocity. Many workers have thought that the monovalent cations serve some control function, changing either the level of activation or inhibition. This has been quite difficult to show since few types of cells exhibit substantial changes in the cation levels which are easily implicated in control functions. For the most part, it is thought that the levels (especially those of the monovalent cation) are sufficiently high such that small changes in these concentrations will not result in changes in the level of activation. However, this question really depends on the free concentrations of the cations and the measurement of their “pool” size. Refinements in these measurements should allow a determination of the plausibility of such a control mechanism. I n the case of other pyruvate kinases, high ATP levels inhibit the enzyme. However, it is not clear that this negative control operates a t all in gluconeogenesis since ATP levels can be high or low. Changing FDP levels will, of course, vary the activity of the FDP-activated enzyme. Intracellular pH may also exert a considerable control over the activity of the enzyme in light of its pronounced effect on the activation by FDP (129, 14.2, 14.3) and inhibition by ATP or alanine (155).Interaction of other metabolic cycles in the control function is evident in the alanine and phenylalanine inhibition and release, but the physiological basis for this is not known. Finally, a few metabolic effectors are known to stimulfttc the synthesis of the enzyme itself. High carbohydrate diets (16, 159. C. Villar-Palnsi and J. Lamer, Annu. Rev. Biochem. 39, 639 (1970). 160. L. A. Kcrson, D. Gnrfinkel, and A. S. Mildvnn, JBC 242, 2124 (1967).

11.

PYRUVATE KINASE

379

161-163) are seen to increase the activity of the liver enzyme, insulin can stimulate its synthesis in rat liver (164) and rat liver cell culture (165), and estradiol, in the r a t uterus (166). The presence of multiple forms of the enzyme might be appropriately discussed a t this point. Conformationally different forms of the enzyme which are apparently interconvertible were discussed in Section I1,E. The mammalian enzyme probably exists as distinct isoenzymes (133, 167) with different electrophoretic patterns. What is not clear, however, is how many of the other reports of kinetically differing forms of the enzyme from a single tissue involve proteins which differ in their primary structure or simply in conformation or bound ligands. Reports of the observation of such multiple forms include B. Zicheniformis (29), M . rowii (38, 155), E . coli K12 (N), rat liver (19, 162, 168), erythrocytes ( 4 , 15, 169, 170),and rat kidney (50). Interconversion between forms have been noted in some of these cases (4, 16, 29, 135, 170). In the case of the liver enzyme, Hess and Kutzbach (168) have shown that of the two forms isolated by isoelectric focusing, one has 2 moles of bound FDP while the other is FDP free. Fresh tissue homogenates, however, mainly contain the FDP loaded species. Such observations suggest that caution should be used in the interpretation of the electrophoretic data which show the appearance of proteins of different charge characteristics.

F. CATALYTIC MECHANISM Since the last review, there have been basically few major advances in understanding the catalytic mechanism of this enzyme. Additional factors affecting the reaction are now known, but the overlapping ATP-P-enolpyruvate site model (I) has not been significantly revised. Basically, this is a mechanism where both substrates are present simultaneously a t the active site in order for phosphoryl transfer to occur. 161. H. A. Krrbs and L. V. Eggleston, BJ 94, 3C (1965). 162. E. Bailey, F. Stripe, nnd C. B. Taylor, BJ 108, 427 (1968). 163. B. Szepesi nnd R. Frcedland, Proc. Soc. Ezp. Biol. Med. 132, 489 (1969). 164. G. Weber, N. B. Stamm, and E. A. Fisher, Science 149, 65 (1965). 165. 1,. Grrschenson and M. Anderson, BBRC 43, 1211 (1971). 166. I;.Jimenez de Aaua, E. Rozengurt. and H. Cnrniinntti, BBA 170, 254 (1968). 167. 1,. Jimencz dc .&in, E. Roarngnrt, J. Dcvallr, and H. Carminatti, BBA 235, 326 (1971). 168. B. Hess and C. Kutzbach, Hoppe-Ssyler's Z. Physiol. Chem. 352, 453 (1971). 169. J. F. Kostcr, G. E. J. Stnnl, and 1,. VanMilligen-Boersmn. BBA 236, 362 (1971). 170. K . Ibsen, K. Schillcr, and T. Haas, JBC 246, 1233 (1971).

380

F. J. KAYNE

The phosphoryl group of P-enolpyruvate occupies the same site as the terminal phosphate of ATP would. This picture closely resembles that obtained for thc catalytic sitc of creatine phosphokinase (I7Oa) wherc such a common phosphoryl group subsite is suggested. Previous experiments have indicated that there is no stable phosphoryl-enzyme intermediate ( I ) , but these do not rule out the existence of a transient phosphoryl cnzymc prcscnt only when both substratcs are bound. Mac farlane and Ainsworth (I7Ob) have now reported a complete kinetic analysis of thc (FDP K+)-activated enzyme from baker’s yeast. They have concluded that the rcaction mechanism is an ordered typc with P-enolpyruvatc binding first followed by ADP and Mg2+. Pyruvate release then takes place before MgATP and a dead-end enzyme-pyruvate complex is also suggested. They also express their idea that Mg‘+, by binding separately, bridges the two substrates and makes a later shift of the cation from the e/3to the /3-y positions of the ATP phosphates unnecessary. However, this might not be a true requirement for the catalytic step, and such a shift could occur after product release. The most recent studies by Robinson and Rose (87) with the rabbit muscle enzyme are of considerable interest here. They have shown that tritium can be lost from the C-3 of P-enolpyruvate during the forward reaction prior to the release of pyruvate. Tritium could also be incorporated from tritiated water into P-enolpyruvate when a trapping system for pyruvate was used. Thus, enByme-bound ATP-pyruvate could revert to substrate and exchange protons with the medium. The exchange characteristics suggested that the product release was rate limiting in regard to both the forward reaction as well as the enolization reaction (86) (when activated by ATP) ; P-enolpyruvate or ADP release was rate limiting for the back labeling. Although a random order of substrate interaction may occur, the authors suggested that “equilibrium kinetics” may not formally be obeyed in the case of the reaction catalyzed by this enzyme. They also observed the effect of various divalent activators on the partitioning of P-enolpyruvate tritium into H,O or pyruvate and showed that this was directly related to the electronrgativity of thc metal ion. They suggestcd that this effect was an indirect one, i.e., that hydroxide in the metal ion coordination sphere or an amino acid residue coordinated to the metal ion could be acting as the base. The study of a model system for phosphoryl transfer, that consisting of the well-known Hg2+-catalyzed hydrolysis of P-enolpyruvate, made

+

170a. J. S. Leigh, Jr., P1r.D. Thesis, University of Pennsylvania, Philadelphia, Pennsylvttnin, 1971. 170b. N. Macfarlane and S. Ainsworth, BJ 129, 1035 (1972).

11.

PYRUVATE KINASE

381

by Benkovic and Schray ( I W ) , showed that Hg2+ addition to the enol double bond resulted in a large catalytic enhancement (los) of the hydrolytic rate. These results led to the suggestion by Robinson and Rose (86) that protonation of the double bond might precede phosphoryl transfer. They were able to show however, that loss of tritium from P-enolpyruvate only occurs during the course of the net reaction; hence, phosphoryl transfer precedes protonation. These authors have also suggested as a result of preliminary experiments that there are no immediately apparent differences between the catalytic mechanisms of the yeast and muscle enzyme. The studies from Rose’s laboratory have allowed a good deal of insight into the mechanism from the standpoint of the enolization (pyruvate) reaction. This reaction’s activator requirements, which are similar to those of the normal reaction, suggest a similar mechanism is involved. But this factor must also be carefully considered since the isotope effect studies indicate an apparent change in the rate-determining step when ATP is compared with any of the other anions (phosphate) used by the enzyme to catalyze the enolization of pyruvate. One may get an impression of the characteristics of the enzyme and its active site from the various physical and chemical studies which have just been discussed. The enzyme appears to be in general, a spherical, quite symmetrical globular protein of four identical subunits. Each of these contains a catalytic site and one or more regulatory (allosteric) sites. The observation of cooperative binding in almost every one of these enzymes indicates subunit interactions which are ligand sensitive. As far as the active site is concerned, one infers a relatively rigid configurational arrangement of the two substrates and two activating cations. Accessibility to the solvent must be relatively good because of the somewhat high turnover number (ca. 250 equivalent/site/sec) , the report of the stereospecific reduction of ppruvate (W), and the tritium exchange results just discussed (87). I n contrast, however, an exclusion of solvent (H,O) is implied by the limited amount of HzO in the Mn2+ coordination sphere measured by proton relaxation rate enhancements of the binary complex ( l o g ) , and, of course, by the fact that H,O would make an excellent phosphoryl acceptor if available to the group undergoing transfer. Indeed, no phosphatase activity is observed for either P-enolpyruvate or nucleotides. Nuclear magnetic resonance studies of some P-enolpyruvate analogs, which are competitive inhibitors with respect to this substrate ( l 7 2 ) , suggest hindered methyl rotation with 171. S. Benkovic and K. Schray, Biochemistry 7, 4097 (1968). 172. T. Nowak and A. S. Mildvan, JBC 245, 8057 (1970).

382

F. J. KAYNE

L-phospholactate. This is presumably the result of steric interaction with an enzyme group involved in the protonation or polarization of the enol double bond (173). The strong specificity and affinity for the P-enolpyruvate substrate in the forward reaction suggests a relatively complete, sterically restricted liganding. This is probably not true of the nucleotide substrate where the purine or pyrimidine moiety is most likely bound away from the phosphoryl transfer site itself. The trinitrophenylation studies (55) suggest a reactive lysine residue is possibly in the vicinity of the nucleotide base binding subsite. I n considering the reverse reaction, however, one notes that F- or hydroxylamine in the presence of HC0,- can apparently bind a t the pyruvate site (83).Even though one can easily imagine these combinations of substrates could occupy a similar-sized site as pyruvate, the liganding groups must exhibit a certain amount of flexibility to permit such coordination. The primary question concerning the mechanism thus still remains to be elucidated: This is whether the enzyme facilitates the phosphoryl transfer from P-enolpyruvate to ADP by forming intermediate covalent complexes with the phosphoryl group or supplies the proper electronic effects to facilitate the direct transfer. And, although it is very tempting to diagram the most probable configuration a t the active site, the lack of information regarding the liganding groups, the question still remaining as to the nature of the divalent cation interaction with the substrates, as well as the uncertainty as to the position of the monovalent cation, lead to the conclusion that the best interests might be served by leaving this picture blank, and with it the possible tendency to misleading prejudices for future studies.

173. T. Nowak and A. S. Mildvan. Biochemistry 11, 2813 (1972).

Creatine Kinase (Adenosine 5'-Triphosphate- Creatine Phospho transferase) D . C. WATTS I . Introduction . . . . . . . . . . . I1. Structure . . . . . . . . . . . . A . Isoenzymes. Interspecific Hybrids. Conformers. and Genetic Variants . . . . . . . . B . Amino Acid Composition . . . . . . . C . Primary Structure . . . . . . . . D . Secondary and Tertiary Structure . . . . E . Subunit Shape and Organization . . . . . F. Molecular Weight . . . . . . . . I11. Purification. Assay. and Enzyme Stability . . . . A . Creatine Kinase from Rabbit Muscle . . . . B . Other Muscle-Type Creatine Kinases . . . . C . Creatine Kinase from Ox Brain . . . . . D . Other Brain-Type Creatine Kinases . . . . E . Hybrid Creatine Kinases . . . . . . . IV . Substrate Specificity . . . . . . . . . A . Guanidine Substrates and the Organization of the Creatine Binding Site . . . . . . . B . Nucleotide Substrates and Related Inhibitors . . V. The Activating Metal Ion . . . . . . . . VI . Enzyme Kinetics . . . . . . . . . . A . Active Forms of the Substrates . . . . . B . Substrate Binding . . . . . . . . C . Temperature Effects . . . . . . . . D . ATP and ADP as Inhibitors . . . . . . 383

. . .

.

384 386

. . . . . . . . . . . .

. . . . . . . . . . . .

386 390 392 393 394 395 395 395 400 401 402 403 403

. . . . . .

403 407 409 412 412 414 420 422

.

. . . . .

.

. . . . .

384

D. C. WATTS

E. Effects of Anions . . . . . . . . . F. Equilibrium of the Reaction . . . . . . . VII. Chemical Investigations of the Enzyme Mechanisin . . . A. Reactive Groups in the Native Enzyme Essential for Catalytic Activity . . . . . . . . . B. Substrate-Induced Conformationnl Changes . . . C. Formation and Topography of the Catalytic Site . . D. The “Essential” Thiol Group . . . . . . . E. Mechanism of Transphosphorylation . . . . .

.

423 428 431

. .

431 436 439 442 451

. .

. .

.

1. Introduction

Creatine kinase (adenosine 5’-triphosphate-~reatinephosphotransferase, EC 2.7.3.2) catalyzes the reaction MgATP2-

+ creatine*

MgADP-

+ phosphocreating- + H+

(1) with the reaction proceeding from left to right being arbitrarily designated the forward reaction. The enzyme has a wide tissue distribution (1, 2 ) and can generally be associated with the physiological role of ATP regeneration in conjunction with contractile or transport systems. It may represent as much as 10-2076 (w/v) of the soluble sarcoplasmic proteins (3-6) of muscle. The enzyme was first crystallized from rabbit skeletal muscle (S), and most investigations have been on the enzyme from this source However, it has also been isolated in an essentially pure state from the skeletal muscle of ox ( 7 ) , human and monkey ( 8 ) , mouse (9),frog and turtle ( l o ) , carp (6, 11), dogfish (6),and chicken ( l a ) ,the smooth muscle of ox (13),the heart muscle of pig (14) and chicken (15),and the brain of cliickcn (If?), ox (17,18), rat (19),and rabbit (16).All ~

1. R. Richterich, U. Wiesmann, and B. Cants, in “Homologous Enzymes and Biochemical Evolution” (N. Van Thoai and J. Roche, eds.), p. 243. Gordon %I Breach, New York, 1968. 2. D. M. Dawson and I. H. Fine, Arch. Neurol. 16, 175 (1967). 3. S. A. Kuby, L. Noda, and H. A. Lardy, JBC 209, 191 (1954). 4. R. Czok and T. Bucher, Advnn. Protein Chem. 15, 315 (1960). 5. C. Gosselin-Rey and C. Gerday, BBA 221, 241 (1970). 6. B. Simonarson, Ph.D. Thesis, University of London, 1971. 7. A. R. Tliomson, J. W. Eveleigh, and B. J. Miles, Nature (London) 203, 267 (1964). 8. I. Kumudavalli, B. H. Moreland, and D. C. Watts, BJ 117, 513 (1970). 9. B. T. Hooton and D. C. Watts, BJ 100, 637 (1966).

12.

CREATINE KINASE

385

these enzymes arc considered to occur in the cell cytoplasm and are readily extracted a t low ionic strength. Creatine kinase with properties distinct from the soluble enzyme has also been found associated with the mitochondria of various tissues (19-24) and where the tissue is rich in mitochondria, as in rat heart or pigeon breast muscle, it may represent 25-50% of the total enzyme (20). The sarcoplasmic reticulum of rabbit skeletal muscle is also reported to contain creatine kinase (25) representing about 1% of the total tissue enzyme, but no properties have been found to distinguish it from the soluble enzyme. Because so little is known about the properties of the particulate enzyme, this review will primarily be concerned with the cytoplasmic forms. These may generally be easily isolated in good yield and have increasingly attracted attention, with the enzyme from rabbit muscle proving an ideal system for investigations using traditional kinetic methods, a variety of inhibitors but thiol reagents in particular, and physical methods measuring proton relaxation rate and electron paramagnetic resonance among others. However, the secrets of creatine kinase are not easily revealed and the progressive awareness of conformational changes, subunit interactions, and unsuspected effects such as those of anions have demanded a continuous reappraisal of past achievements. This review attempts to paint the picture as it is currently seen but makes no promises as to how long the colors will glow true in the future.

10. H . Kulhcrtus and A. DistBche, Arch. Int. Physwl. Biochim. 70, 246 (1962). 11. C. Gosselin-Rey, G . Hamoire, and R . K. Scopes, J . Fish. Res. Bd. Can. 25, 2711 (1968). 12. 13. P. Roy, J. F. Laws, and A. R. Thomson, BJ 120, 177 (1970). 13. B. Focant, FEBS Lett. 10, 57 (1970). 14. A. I. Keto and M. D. Doherty, BBA 151, 721 (1968). 15. B. T. Hooton, Biochemistry 7, 2063 (1968). 16. H. M. Eppenberger, D. M. Dawson, and N. 0. Kaplan, JBC 242, 204 (1967). 17. H . J. Keutel, H. K. Jacobs, K . Okabe, R. H. Yue, and S. A. Kuby, Biochemistry 7, 4283 (1968). 18. T. Wood, BJ 89, 210 (1963). 19. R. J. Sullivan, 0. N. Miller, and 0. Z . Sellinger, J . Neuroclzem. 15, 115 (1968). 20. H. Jacobs, H. W. Heldt, and M. Klingenberg, BBRC 16, 516 (1964). 21. E. K. Brownlow and D. B. Gammack, BJ 103, 47P (1967). 22. P. D . Swsnson, J . Neurochem. 14, 343 (1967). 23. J. H. Ottaway, Nature (London) 215, 521 (1967). 24. A. I. Keto and M. D. Doherty, BBA 151, 721 (1968). 25. R. J. Baskin and D. W. Deamer, JBC 245, 1345 (1970).

386

D. C. WATTS

It. Structure

A. ISOENZYMES, INTERSPECIFIC HYBRIDS,CONFORMERS, AND GENETICVARIANTS In vertebrates the soluble creatine kinase occurs as three forms readily distinguishable by their electrophoretic mobility (26-28) , and studies of the changing pattern during development suggested that the form with an intermediate mobility was a hybrid of the electrophoretically fast and slow enzymes (26, 29). This was confirmed by dissociating the subunits of a mixture of the purified fast and slow forms by treatment with 6.5 M guanidine hydrochloride or by freezing in the presence of salt and sodium phosphate. After reactivation, by dilution or dialysis, electrophoresis revealed the presence of the hybrid with an appropriate mobility intermediate between the two parental forms (SO). The threeband pattern also indicated that the enzyme existed as a dimer. Accordingly the three forms were called muscle type (MM), hybrid (MB), and brain type (BB) isoenzyme, in order of increasing mobility toward the anode a t pH values above neutrality, to indicate their major tissue of origin. During development the brain-type isoeneyme appeared first in all the tissues studied. This form remains constant as development proceeds in the brain. In chicken heart and in mammalian smooth muscle, the brain-type isoenzyme remains the predominant form accompanied by a trace of the hybrid, while in mammalian heart and red and white striated muscle fibers the muscle-type isoeneyme gradually predominates and only a trace of the hybrid remains (26, 29, 3 1 ) . Slight variations may occur in these patterns with more or less of the minor components. Atrophy of the white muscle fibers, whether caused by denervation, vitamin E deficiency, or a hereditary dystrophy, is accompan26. A. Burger, M. Eppenberger, and U. Wiesmann, Helv. Physiol. Actu 21, C6 (1983). 27. A. L. Sherwin, G. R. Siber, and M. M. Elhilali, Clin. Chim. Actn 17, 245 (1967). 28. S. Rosalki, Nature (London) 207, 414 (1965). 29. H. M. Eppenberger, M. Eppenberger, R. Richterich, and H. Aebi, Develop. BWZ. 10, 1 (1964). 30. D. M. Dawson, H. M. Eppenberger, and N. 0. Kaplan, BBRC 21, 346 (1965). 31. I. Kumudavalli and D. C . Watts, BJ 108, 547 (1968).

12.

CREATINE KINASE

387

ied by a loss of muscle-type isoenzyme and reappearance of the brain form-the so-called reversion to the fetal state (Sla, S l b ) . Formation of the hybrid can readily be achieved using crude tissue supernatants and, using such preparations, the ready formation of interspecific MB hybrids between extracts from cat, human, and chicken h a s been demonstrated ($9).With these species all the M M isoenzymes are similar but slow. A different situation exists with the muscle enzyme of the dogfish, Scylliorhinus canicula, which has a sufficiently fast mobility to allow detection of a MM’ hybrid, which may be formed by the same procedures as used for the MB hybrids, between it and the rabbit muscle enzyme (33). But perhaps most interesting so far has been the formation of hybrids between the BB forms of rabbit or ox with the dimeric arginine kinase from muscles of the sea cucumber, Holothuria jorslcali (34).Bearing in mind that no hybrids will form between the kinase.and any other protein in a crude tissue extract, these findings demonstrate that among the phosphagen kinases conservation of the essential features for subunit interaction over long periods of evolution from invertebrates to vertebrates is just as important as those for the catalytic site itself. Passeriform birds were thought to be unusual in containing two braintype isoenzymes, which upon hybridization with a muscle enzyme form two hybrid bands (36). Electrophoresis a t p H 6.5, rather than a t the usual pH of 8.6 has now revealed that 15 of the 28 known avian orders have BB enzymes that behave in this way, the Psittaciformes being the only exception found so far (35~). The available evidence (36) indicates that the two brain forms have identical primary structures folded into two slightly different, but nevertheless active, conformations. I n support of this hypothesis, it has been found that submitting the brain-type enzymes of OX smooth muscle (34) or the chicken (32) to the hybridization procedure may result in the formation of an additional band of enzymic activity 31a. F. Schapira, Compt Rend. 262, 2291 (1966). 31b. F. Schapira, in “Homologous Enzymes and Biochemical Evolution” ( N . Van Thoai and J. Roche, eds.), p. 151. Gordon & Breach, New York, 1968. 32. D. M. Dawson, H. M. Eppenberger, and N. 0. Kaplan, JBC 242, 210 (1967). 33. B. A. Simonarson and D. C. Watts, BJ 128, 1241 (1972). 34. D. C. Watts, B. Focant, B. Moreland, and R. L. Watts, Nature (London), New Biol. 237, 51 (1972). 35. M. E. Eppenberger, H. M. Eppenberger, and N. 0. Kaplan, Nature (London) 214, 239 (1967). 35a. A. Scholl and H. M. Eppenberger, Ezperientiu 25, 794 (1969). 36. H. M. Eppenberger, in “Homologous Enzymes and Biochemical Evolution” (N. Van Thoai and J. Roche, eds.), p. 231. Gordon & Breach, New York, 1968.

388

D. C. WATTS

suggesting that the ability to adopt more than one kinetically stable conformation is inherent to some degree in all brain-type creatine kinases. Electrophoresis in 8 M urea gels of the purified chicken BB enzyme from heart muscle revealed two protein components from which the existence of two different creatine kinase subunits was inferred (16). Apart from the difficulty that random combination of two different subunits might be expected to give three bands of activity upon electrophoresis, this observation suggests alternative cxplanatioiis of conformer formation such as limited proteolysis by trace contaminants or chemical modification of an amino acid side chain (see Section VI1,D). Creatine kinases of fish tissues may also show more complex electrophoretic patterns than can readily be equated with the two-subunit, threeband pattern ($624, 3%). Extracts of the stomach and small intestine of trout, for example, were found to contain up to seven bands. Skeletal muscle contained only one MM band but submitting the extract to the hybridization procedure caused the formation of additional bands with similar mobilities to those found in the other tissues. Oxidation and adventitious ligand-binding were discounted as possible causes of this effect so that, again, conformational isomers seem the only explanation. More recently (36c), other fish have been found to contain additional slowly migrating bands, particularly in extracts of stomach and testis. It is suggested from hybridization experiments (364 that these may reflect the presence of additional genes, resulting from the polyploid nature of these species, as has been found with other enzymes. Out of keeping with its apparent occurrence as an enzyme from the cell cytoplasm, creatine kinase does not commonly show electrophorctic variation attributable to a genetic origin. No genetic variants of the mammalian enzymes are known, and while patterns of either two bands or a single band in carp (Cyprinus carpio) muscle extracts werc inferred to represent the heterozygote and one of the homozygote forms (37) this was not supported by formal genetic analysis. Herring muscle also shows a double-banded pattern, and of 250 individuals examined only one was found with a single band. Sixteen other species of fish examined gave only one band of creatine kinase activity (38).From the absence of the characteristic three-banded pattern expected for the 36a. C. Gosselin-Rey, G. Hamoir, and R. K. Scopes, J . Fish. Res. Bd. Can. 25, 2711 (1968). 36b. H. M. Eppenberger, A. Scholl, and H. Ursprung, FEBS Lett. 14, 317 (1971). 36c. A. Scholl and H. M. Eppenberger, Comp. Biochem. Physiol. 42B, 221 (1972). 36d. J. C. Pcrriard. A. School, and H. M. Eppenberger J. Ezp. 2001.182, 119 (1972). 37. R. K. Scopes and C. Gosselin-Rey, J . Fish. Res. Bd. Can. 25, 2715 (1968). 38. B. Simonarson and D. C. Watts, unpublished data (1972).

B

TABLE I AMINOACIDCOMPOSITIONS OF MUSCLE-TYPE (MM) CREATINE KINASES Rabbit Amiio acid

(Ref. 41)

(Ref. 16)

(Ref. 42)

Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valiie Methionine Isoleucine Leucine Tyrosine Phenylalanine Tryptophan Total Amide

65 33 34 83 34 41 74 37 62 25 8 53 19 25 70 19 30 8 720 52

61 32 33 83 34 44 78 35 64 26

66 34 35 85 35 42 76 37 64 26 8 54 19 26 71 19 31 7 735 55

From 16.

49 17 22 72 20 31 701

ox (Ref. 43)

Chicken (Ref. 16)

Human (Ref. 8)

Monkey (Ref. 8)

Dogfish (Ref. SS)

Carp (Ref. 44)

65 29 34 81 30 35 79 35 65 38 7 54 18 30 68 16 32

65 34 40 73 29 32 83 41 60 33 7a 49 20 24 71 16 33

55

728

710

66 28 32 73 31 41 75 45 66 48 7 46 19 25 64 20 26 6 718 53

61 36 38 83 33 32 73 36 63 30 6 48 20 32 70 19 26 8 714

61 26 32 91 38 41 72 31 63 27 8 50 21 33 64 15 31 6 710 44

32 66 28

45 75 42 64 35 47 18 24 62 17 28

2

2% M

390

D. C. WATTS

heterozygote of a dimeric enzyme and the known occurrence of conformers, it must be concluded that a positive demonstration of genetic variation in creatine kinase is still awaited. The creatine kinases from invertebrates may show an electrophoretic pattern with single or multiple banding (39, 40). The cause of the latter is unknown.

B. AMINOACID COMPOSITION This has now been determined for the MM and BB creatine kinases from a number of species (Table I) (8, 16, 16, 33, 41-44) (Table 11) ( I S , 15, 16, 46). I n general, the compositions are all very similar and TABLE I T AMINOACIDCOMPOSITIONS OF BRAIN-TYPE (BB) CREATINEKINASES

ox Amino acid Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cys tine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Tryptophan Total Amide

Rabbit (Ref. 16)

Brain (Ref. 63)

Stomach (Ref. 13)

Chicken (Ref. 16)

52 30 22 82 34 32 74 40 65 35 47 20 30 81 21 41

44 23 40 83 34 38 85 48 66 52 10 48 18 30 73 19 29

46 26 44 89 34 39 81 39 59 39 10 41 17 28 76 17 30

49 21 40 87 35 35 75 31 68 36 9a 48 19 28 82 21 30

706

740

715 61

714

* From 16. 39. R. Virden and D. C. Watts, Comp. Biochem. Physiol. 13, 161 (1964). 40. J. A. E. Fitzsimmons and M. D. Doherty, Comp. Biochem. Physiol. 36, 1 (1970). 41. E. A. Noltmann, T. A. Mahowald, and S. A. Kuhy, JBC 237, 1146 (1962).

12. CREATINE

391

KINASE

the estimates for the rabbit MM enzyme by different workers show good agreement. Thoai ( 4 S ) , comparing phosphagen kinases, noticed that the proportion of amino acids with similar types of side chain were comparable in the different enzymes. Using his analysis (Table 111) (IS, 16, 41, 4S1 45) it can be seen that the BB enzymes contain significantly less of the basic amino acids than the MM forms, explaining the greater electrophoretic mobility, as well as rather more of cysteine and the aromatic amino acids. For a given species the amino acid composition of the MB hybrid is intermediate between that of the parent forms (16). An interesting problem arises with the brain-type enzymes from ox brain and stomach muscle. There are quite significant differences between the two amino acid compositions, particularly in the numbers of proline, glycine, valine, and alanine residues, suggesting that these two forms COMPARISON

TABLE 111 COMPOSITION OF GROUPSOF AMINO SOMECREATINE KINASES

O F THE PERCENTAGE: MOLAR

ACIDSIN

~

Amino acid

+ + + + + + + +

Asp G l ~ i Lys His Arg T y r + Phe (+ Trp.) Met Cys Ser Thr Pro Gly Ala Val Ile Leu Reference

+

ox

Rabbit

Chicken

Skeletal Stomach muscle Brain muscle MM BB (BB?)

Skeletal muscle Brain MM BB

Skeletal muscle Brain MM BB

22.2 17.7 6.7 3.6b 9.0 5.4 35.4

24.7 14.5 6.5 3.8 9.7 6.5 36.4

23.8 16.3 6.5 3.8 10.2 5.4 34.0

21.8 18.4 7.8 3.8 9.4 5.1 32.7

21.7 14.6 8.7 4.2= 9.2 5.6 36.0

22.0 22.7 19.5 15.4 6.9 7.1 3.gb 4 . 1 ~ 8.4 9.8 5.7 4.3 3 3 . 4 36.7

43

63

13

41

16

16

16

Values for tryptophan were not included in this analysis. A value of 8 half-cyst,inea per mole was assumed for this calculation. c A vrtlue of 10 half-cystines per mole was assumed for this calculation. a

~~

42. R. H. Yue, R. H. Palmieri, 0. E. Olson, and S. A. Kuby, Biochemistry 6, 3204 (1967). 43. A. R. Thomson, J. W. Eveleigh, J. F. Laws, and B. J. Miles, in “Homologous Enzymes and Biochemical Evolution” (N. Van Thoai and J. Roche, eds.), p. 255. Gordon & Breach, New York, 1968. 44. C. Gosselin-Rey and C. Gerday, BBA 221, 241 (1970). 45. A. R. Thomson, J. Eveleigh, and J. B. Miles, Nature (London) 203, 267 (1964). 46. N. Van Thoai, in “Homologous Enzymes and Biochemical Evolution” (N. Van Thoai and J. Roche, eds.), p. 220. Gordon & Breach, New York, 1968.

392

D. C. WATTS

could be separate isoenzymes coded by separate structural genes. However, a decision on this point must await further evidence since an improved preparation of the stomach muscle enzyme has since been obtained (47)and the proline and alanine values for the brain enzyme appear atypically high. The partial specific volume calculated from the amino acid composition for the rabbit muscle enzyme (41) is 0.735 cms/g, in good agreement with that obtained by physical measurements (42, 48).

C. PRIMARY STRUCTURE Of the 360 or so amino acids comprising each subunit only a few short lengths representing less than 10% of the total sequence are known (Table IV) (46, 49-52). Sequences I and I1 are those concerned with the essential cysteine residue which is readily and specifically labeled with alkylating agents. With one or possibly two differences, they are identical in the brain- and muscle-type enzymes and similar sequences have been reported for other phosphagen kinases (see Chapter 13, this volume). The reaction of 2 moles of alkylating agent per mole of enzyme yielded only a single sequence in accord with the evidence from TABLE IV KNOWNPEPTIDESEQUENCES IN CREATINE KINASE Sequence No.

Source

Sequence"

lief.

I

Rabbit (MM) Val-Leu-Thr-CYS-Pro-Ser-Asn-Leu-Gly-Thr-Gly-LeL~-Arg 45,

I1 I11 IV V VI

Rabbit (BB) Ile-Leu-Thr-CYS-Pro-Ser-Asx-Leu-Gly-Thr-Gl~-Leu-Arg Rabbit (MM) Asp-Ser-Pro-Val-LYS-Leu-Leu-Phe Rabbit (MM) Asp-Ser-Pro-Val-LYS-Leu-Leu-Phe-Leu Rabbit (MM) Asp-Val-Val-Gly-Gly-Glu-Glu-CYS-Ala-Ser Rabbit (MM) Gln-Lys-COOH

49 50 51 51 51 52

The amino acid residue shown in capitals is that to which the identifying label waa attached. For details see the discussion in the text. 47. B. Focant and D. C. Watts, unpublished data, 1972. 48. L. Noda, S. A. Kuby, and H. A. Lardy. JBC 209, 203 (1954). 49. T. A. Mahowald, Biochemistry 4, 732 (1965). 50. R. S. Atherton, J. F. Laws, B. J. Miles, and A. R . Thornson. BJ 117, 30P (1970). 51. T. A. Mahowuld, Fed. Proc., Fed. Amer. Soc. Exp. Biol. 28, 601 (1969). 52. 0. E. Olson and S. A. Kuby, JBC 239, 460 (1964).

12.

CREATINE ICINASE

393

isoenzyme studies (Section II,A) that crestine kinase has two subunits. Sequences I11 through V were obtained by using the bifunctional reagent, 1,5-difluoro 2,4-dinitrobenzene, to link the essential thiol of sequence I to other reactive groups on the same or another subunit. Sequence V presents difficulties in that thiols other than the essential cysteine group do not react to a significant extent until the tertiary structure of the enzyme has become unfolded to some degree. Hence, some cross-linking may have occurred during denaturation prior to proteolytic digestion. Sequence VI is the C-terminus determined by the use of carboxypeptidase. Removal of the C-terminal dipeptide is without effect on catalytic activity. A more quantitative assault on the C-terminus using hydrazinolysis and carboxypeptidase B yielded 2 moles of valine per 82,600g protein by each method showing that each of the two subunits is composed of a single polypeptide chain (4.2). No N-terminus could be detected in the rabbit muscle enzyme by using fluorodinitrobenzene or phenylisothiocyanate either on the native protein or after denaturation with urea (54). Autoradiography of electrophoretograms of acid digests of creatine kinases labeled with 14C-iodoacetate (43) has confirmed that the sequences around the essential thiols of the ox M M and BB and the rabbit MM isoenzymes are very similar. On the other hand, peptide mapping of the denatured and fully alkylated MM and BB isoenzymes reveals significant differences in the sequences around some of the buried thiols, a t least in the content of charged amino acids. This is in accord with conventional fingerprints studies ( 5 , 8, 9, IS) which indicates that the MM isoenzymes of different species are more similar in their primary structures than are the M M and BB isoenzymes within a single species. As an extreme example of species difference, the muscle enzymes of carp and rabbit, after tryptic digestion, have about 65% of their peptides in common (6) while for the BB and MM chicken enzymes the figure is only about 55% (16).

D. SECONDARY AND TERTIARY STRUCTURE The usual parameters from optical rotatory dispersion studies (55-57) indicate a compact globular structure containing 25-30% a-helix and less 53. A . R. Thompson, quoted in reference 13,.

54. T. A. Mahowald and S. A. Kuby, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 19, 46 (1960). 55. A. Samuels, Ann. N . Y. Acad. Sci. 103, 858 (1963). 56. J. H.Kiigi and T. K. Li, Fed. Proc., Fed. Amer. Soe. E z p . B i d . 24, 285 (1965). 57. C. Oriol-Audit, M. Landon, Y. Robin, and N. Van Thoai, BBA 188, 132 (1969).

394

D. C. WATTS

than 15% /?-pleated sheet. It is of interest that two other dimeric phosphagen kinases yield similar values for the content of a-helix but monomer arginine kinases give higher values of 3540% (57). The resistance of the native protein to tryptic digestion (4% of the rate observed after denaturation) and to the exchange of more than 60% of the amide hydrogens for deuterium (58) supports the view of the molecule as a compact structure. Immunological evidence is in accord with that from studies of the primary structure in indicating that the tertiary organization of the same isoenzyme from different species is more similar than that of the MM and BB isoeneymes of a single species. Antisera raised against one isoeneyme will not cross react with the other although it will with the same isoeneyme from other species (59, 60). This does not extend beyond the class of animals since although antisera to rabbit MM will cross react with the muscle enzymes of ox, mouse, and rat there is no reaction with those of pigeon, goldfish, or several invertebrates (60).

E. SUBUNIT SHAPEAND ORGANIZATION The previous sections have shown that creatine kinase consists of two freely dissociable subunits, each consisting of a single polypeptide chain containing no disulfide bridges (60a). Sedimentation studies under various conditions confirm this picture (42). The calculated frictional ratio (f/fo) was found to be only 1.21 and the axial ratio 4.4 (for an assumed anhydrous prolate ellipsoid), indicating a compact, rather cigarshaped molecule. A useful finding was that while guanidinium chloride dissociates and unfolds the subunits into the random coil configuration, sodium lauryl sulfate causes them to dissociate without any apparent loss of structural organization. The subunits were found to have the same frictional ratio as the native molecule and the same calculated axial ratio of 4.4 for an assumed anhydrous prolate ellipsoid (4%).Hence the molecule can be envisaged as consisting of two cigar-shaped subunits lying side by side rather than as two egg-shaped subunits joined end to end. The calf brain enzyme appears to have an identical shape to that from rabbit muscle (61), and from the findings that a dimeric 58. N. S. T. Lui and L. Cunningham, Biochemistry 5, 144 (1966). 59. J. A. Bulcke and A. L. Sherwin, Zmmunochemistiy 6, 681 (1969). 60. B. Viala, Y. Robin, and N. Van Thoai, Comp. Biochem. Physiol. 32, 401 (1970). 60a. P. M. Bayley and A. R. Thomson, BJ 104, 33C (1967). 61. R. H. Yue, H. K. Jacobs, K. Okabe, H. J. Keutel, and S. A. Kuby, Biochemistry 7, 4291 (1968).

12.

CREATINE KINASE

395

arginine kinase will form a hybrid with creatine kinase (S4) it would seem that this shape has been strongly conserved throughout evolution.

F. MOLECULAR WEIGHT The original investigation of the physical properties of creatine kinase (48) yielded a molecular weight of 81,000, and until a detailed reinvestigation produced a new figure of 82,600 (62) this value had been accepted for practical purposes although not always quite in accord with the experimental data. Fortunately, the correction required is less than 2%. Experimentally determined values for other purified creatine kinases fall within the range 78,500-85,100 ( I S , 32,3.9, S7, 61,62) with the latter, somewhat high value being for the carp muscle enzyme (S7). Molecular weight determinations of the individual subunits indicate that these are half those for the intact molecule (42, 61,6%). 111. Purification, Assay, and Enzyme Stability

A. CREATINE KINASEFROM RABBIT MUSCLE 1. Purification

The first described isolation procedure using alcohol fractionation (method B) (3) with only slight modifications (63) is generally used to isolate the enzyme in good yield and with a high specific activity. The use of phosphocellulose to further improve the purity has recently been described (6Sa). Further traces of impurities may be removed by an improved recrystallization procedure (BSa, 6 4 ) . 2. Assay and Specific Activity

For the forward reaction (phosphocreatine synthesis) a specific activity of 75 units (often referred to as “Kuby units”) may be obtained by assaying for acid-labile phosphocreatine (3). One unit of enzyme is defined as that amount per milliliter of reaction mixture which will catalyze the transphosphorylation reaction between 1 mM ATP and 24 mM 62. B. Moreland, D. C. Watts, and R. Virden, Nature (London) 214, 458 (1967). 62a. M. C. Grant-Greene and F. Friedberg, Znt. J. Protein Res. 2, 235 (1970). 63. L. Noda, T. Nihei, and M. F. Morales, JBC 235, 2830 (1960). 63a. H. J. Keutel, K. Okabe, H. K. Jacobs, F . Ziter, L. Maland, and S. A. Kuby, ABB 150, 648 (1972). 64. T. A. Mahowald, E. A. Noltmann, and S. A. Kuby, JBC 237, 1535 (1962).

396

D. C. WATTS

creatine in the presence of 6 mM MgSO, at pH 9.0 (glycine buffer) and 30°, and yield an apparent second-order velocity constant (Ic’) equal to 1 ml/pmole/min; k’ = kE in the relation dx/dt = kE[ATP]2[Cr]o. The conditions are such that the reaction is apparently second order with respect to ATP and the apparent second-order rate constant is proportional to enzyme concentration. These are not V,,, conditions, and the value of the assay has been in providing a convenient method for comparing the preparations from different laboratories. Although the phosphate assay is still widely used, the continuous trace afforded by a pH stat makes it the method of choice. In the presence of albumin as stabilizing agent, an approximately equivalent specific activity is 240 pEq/mg protein/min (64).The role of the albumin appears to be simply to buffer against denaturation caused by the alkali titrant producing local fluctuations in pH; 1 m M cysteine fulfills this role equally well and provides additional protection against oxidation ( 6 6 ) . For spectrophotometric assay, suitable coupling systems employ pyruvate kinase and lactate dehydrogenase for the forward reaction (66) and hexokinase and glucose-6-phosphate dehydrogenase for the back reaction (67). The pH-drift method of assay, used for rapid reaction kinetics, either by direct measurement with a pH meter or spectrophotometrically using a dye indicator, also offers possibilities that have so far only been exploited in temperature-jump studies ( 6 8 ) .The reverse reaction may also be readily followed by the colorimetric determination of creatine (69). Methods exploiting the fluorescence of creatine in the presence of ninhydrin have been described (70, 71) but not widely employed. I n general, the assay system should avoid buffers that chelate (not always easy to achieve), and Mgz+ions, as the acetate, should be added in 1 mM excess over the ATP concentration to ensure that all the ATP exists as the MgATP complex (72). Excess Mg2+ions are inhibitory (73). The inclusion of other small anions such as C1-, NO,-, HCO,-, or SO!should normally be avoided (see Section VI,E) . The presence of 0.01-0.05 mM EDTA stabilizes and activates the enzyme, apparently by chelating traces of heavy metals added with other components of the system (79). 65. E. J. Milner-White and D. C. Watts, BJ 122, 727 (1971). 66. M. L. Tanzer and C. Gilvary, JBC 234, 3201 (1959). 67. I. T. Oliver, BBA 14, 587 (1958). 68. G. G. Hammes and J. K. Hurst, Biochemistry 8, 1083 (1969). 69. J. F. Morrison, W. J. O’Sullivan, and A. G. Ogston, BBA 52, 82 (1961). 70. R. B. Conn and V. Anido, Amer. J . Clin. Pathol. 46, 177 (1966). 71. J. C. Koedam, Clin. Chim. Acta 23, 63 (1969). 72. W. W. Cleland, Annu. Rev. Biochem. 36, 77 (1967). 73. S. A. Kuby, L. Noda, and H. A. Lardy, JBC 210, 65 (1954).

12.

397

CREATINE KINASE

-

5

6

7

8

9

n

I

-

5

6

7

-. 8

9

100 -

90 -

<>

'= B

8

80 70 6050-

30 -

40

20 10 I

I

I

I

FIG.1. pH-activity curves for creatine kinase. (A) For the rabbit muscle (65) measured in the forward direction at (0) lo", (0) 30°, and ( 0 )38", using ATP, 1 m M ; MgAc*, 6 m M ; creatine, 24 mM; and histidine-acetate, 50 mM, in the pH range 5.5-8.5 and glycine, 50 mM, from p H 8.5 to 9.0.(B) For the rabbit muscle 30", using ADP, enzyme (86) measured in the reverse direction a t ( X ) 20" and (0) 0.5 mM; MgAc,, 15 mM; phosphocreatine, 5 m M ; and the same buffers as for (A). The solid line is the 30" curve from (A) drawn in for comparison. (C) For the ( A ) chicken muscle and (0) brain enzymes (16) measured in the reverse direction a t 25" using ADP, 0.5 m M ; MgCL, 3.3 m M ; phosphocreatine, 8 m M ; and trisacetate, 100 mM; in the p H range 5.5-6.2; tris-histidine, 100 m M; in the pH range 5.9-7.3 and tris-HC1, 100 m M ; from pH 7.5-8.8.

398

D. C. WATTS

Thiols (1 mM cysteine or mercaptoethanol or 0.1 mM dithiothreitol) have been found to activate or stabilize some (usually poor) preparations, particularly after prolonged storage under less than ideal conditions. The pH-activity curve shows a broad optimum between 7.5 and 9.5 for the forward direction, depending somewhat on the conditions used (Figs. 1 and 2). That for the back reaction shows a lower optimum, as might be expected since a proton is taken up, and a t physiological pH the enzyme is only about 20% active. It is a neat physiological adaptation in muscle that the sudden fall in pH that accompanies contraction and ATP splitting moves the enzyme sharply toward the optimum thereby facilitating the rephosphorylation process. 3. Stability

After prolonged dialysis against several changes of a nonvolatile (glycine) low ionic strength (0.001-0.005 I ) buffer, p H 9.0, rabbit muscle creatine kinase shows a remarkable degree of stability. When stored as a 3-576 w/v sterile solution a t 0 ' 4 " specific activity may actually increase over a 6-month period as a result of the precipitation of less stable trace contaminants and, possibly, reactivation of partially denatured enzyme. The freeze-dried powder, stored at - 18" has been found to lose only a small amount. of activity even after 2 years with

i$

100

-

80

-

$

e -

0

c 2.

-

w

-

-

60-

.-V

5

40-

w

20 0

I

I

1

FIG.2. Effect of pH on the reactivity of the essential thiol groups with iodoncetamide and the stability of creatine kinase from blue hare skeletal muscle. The enzyme was allowed to stand a t the p H shown for 10 min at 25" (0) in the absence or ( 0 )in the presence of 0.1 rnM iodoacetamide and then assayed for residual enzymic activity. The lack of change in the ionization state of the essential thiol groups is shown by the constancy of the residual activity which does not decrease until the enzyme denatures.

12.

CREATINE KINASE

399

frequent warming and refreezing to remove samples. Crystalline suspensions in alcohol stored a t - 12°C appear equally stable (@). Shorter term investigations show the enzyme to be unstable above 50" but to be fairly stable a t 37" between p H 6.5 and 9.5 (73). Under some conditions stability in the high p H range may extend to p H 10.5 or above for short periods of time but, as shown with the muscle enzyme from the closely related hare (Fig. 2), once the limit is reached denaturation is rapid and irreversible although the enzyme does not precipitate. At 20" the enzyme will also withstand 10 min a t pH 3.0. Physical measurements show that while the titration curve for enzyme tyrosine groups is reversible beyond p H 11.6, precipitation occurs when the enzyme is returned to pH 7.2. Without such treatment the enzyme shows only slight turbidity a t the isoelectric point (pH 6.1) for the conditions used (74, 7 6 ) .The sedimentation constant also decreases from s20,w 5.6 to 3.0 and the relative viscosity increases from 1.02 to 1.14 around the upper renaturation point ( 7 4 ) . A similar sharp transition point for enzymic inactivation occurs a t low pH (76). Incubation for 1 hr a t 30" at p H 5.0 causes 95% loss of activity but only 5% loss a t p H 5.8. A shift of only 0.31 p H units was found to cause a tenfold change in denaturation rate. At p H 5.0 activation energies for denaturation of 31 and 86 kcal/mole were found for temperatures below 30" and above 40", respectively, with both modes of denaturation being initiated by 3.4 hydrogen ions per molecule. Enzyme that had been alkylated with iodoacetate was denatured a t the same rate as the native enzyme, and the denaturation initiation point was inferred not to be part of the catalytic site (76).The enzyme is stabilized to some extent by an equilibrium mixture of substrates (73),but not by inorganic phosphate (76)as once thought. This problem of acid denaturation is further discussed by Keutel et al. (6%). Physical measurements a t low p H indicate rather complex changes in structure. The sedimentation coefficient does not change significantly until below pH 5.0 while the relative viscosity shows a gradual increase between pH 6.0 and 4.2 from 1.02 to 1.06, and a sharp increase in viscosity does not begin until pH 3.4 ( 7 4 ) .As with alkaline denaturation these changes are irreversible and although the enzyme remains soluble it precipitates when returned to neutrality. The titration curve for carboxyl and imidazole residues is reversible over a limited p H range below the precipitation region (77). 74. 75. 76. 77.

W. A. DaCosta and F. Friedberg, JBC 235, 3134 (1960). J. R. Sparks, W. A. DaCosta, and F. Friedberg, ABB 95, 371 (1961). R. K. Scopes, ABB 110, 320 (1965). B. F. Floyd and F. Friedberg, JBC 242, 1027 (1967).

400

D. C. WATTS

B. OTHERMUSCLE-TYPE CREATINE KINASES 1. Purification For the different sourccs used so far (SCC Scctioii I) purification prcsents few problems. It is generally achieved by either alcohol fractionation as used for rabbit muscle (3) or acetone fractionation (5, 33) followed by chromatography on diethylaminoethyl cellulose ion exchanger (5,9,13,33,63a)or electrophoresis (8,33).Acetone fractionation (78) has proved particularly suitable for the fish enzymes, and ammonium sulfate has been used for the enzymes from chicken and monkey muscle ( 7 9 ) . 2. Assay and Specific Activity

Using the procedures described for the rabbit cnzymc, it is found that all the mammalian enzymes investigated have similar specific activities. Similar specific activities were also found with the enzyme from chicken muscle where a value of 70-84 Kuby units a t 30" (12) may be compared with a value of 720 pmoles/min/mg protein a t 25" obtained using the spectrophotometric assay in the reverse direction (16). The fish enzymes, on the other hand, have low specific activities, one-third to onefifth that of the rabbit enzyme. A complication in carrying out the comparison with the dogfish enzyme (33) was that the Michaelis constant for creatine was more than twice that for the rabbit enzyme under the standard assay conditions a t 30". But even correction for this difference left the dogfish enzyme less than one quarter as active as the rabbit enzyme. A partial explanation appears to be that the dogfish enzyme evolved to function a t temperatures nearer 12". Decreasing the temperature from 30°, a t which the specific activity was measured, to 12" decreascs tlic activity of the dogfish enzyme by half (33) but that of the rabbit enzyme is decreased fourfold (73). Hence, a t the physiological temperature of the dogfish, the two enzymes are much less different suggesting that the higher activity of the mammalian and bird enzymes may be considered as an evolutionary adaptation to a much warmer environment. This change is also indicated by the activation energy for the forward reaction being about 6,500 cal/n~olefor the fish enzyine(s) and 11,000 cal/mole for the rabbit enzyme (73).An additional compensating factor may be that the fish enzymes represent 16-20% of the sarcoplasmic proteins (6, 33), nearly twice that found in rabbit muscle. 78. B. A. Aslronns, BJ 48, 42 (1951). 79. J. A. Underhill and D. C. Rat,ts, unpublislicd data (1972).

12.

CREATINE KINASE

401

3. Stability The stability of various mammalian muscle-type enzymes has recently been investigated (6Sa); the only enzyme that appears a t all unstable is that from primate muscle where the thiol groups are sensitive to oxidation more in the manner of the brain-type enzymes (q.v.).

KINASEFROM O x BRAIN C. CREATINE 1. Purification Creatine kinase constitutes less than 1% of brain cytoplasmic proteins (17).Consequently, the purification procedure is correspondingly more complex (17, 18), involving two rounds of fractionation using ammonium sulfate and either acetone (18) or ethanol a t -10" ( 1 7 ) . An important improvement is the use of phosphocellulose (17) to remove hemoglobin-like proteins and any muscle-type creatine kinase present in the extract. Unlike most muscle enzymes, brain enzymes require thiols to maintain them in an active state, and enzyme solutions are maintained in 1 m M 2-mercaptoethanol after the initial fractionation stages. The enzyme crystallizes from ammonium sulfate as relatively large octahedrons (17) in contrast to thc fine needles obtained with the muscle enzyme by crystallization from ethanol (3). 2. Assay and Specific Activity Using the samc phosphate assay as for thc muscle enzyme (3) specific activities of 73 and 130-140 Kuby units (references 18 and 17, respectively) were claimed for purifications of 50- and 200-fold1 respectively. Thc latter spccific activity was found to equal 240-250 pEq/mg per protein a t 30" when assayed by the pH-stat assay (64). This is the same as found for the rabbit muscle enzyme although the specific activity in Kuby units is nearly double. This discrepancy, as stated above, results from the fact that the phosphate assay procedure does not use V,,,, conditions ; thus, the comparison of enzymes from different sources becomes meaningless if there are significant differences in the substrate and product binding properties of the enzymes being compared. The pHstat assay more closely approaches V,,,, conditions.

3. Stability I n the presence of 2-mercaptoethanol the enzyme shows a similar range of stability to the muscle enzyme at 0" but is much less capable

402

D. C. WATTS

O°C with mercaptoethanol

FIQ.3. The effect of p H on the stability of calf brain creatine kinase in the presence and absence of 2-mercaptoethanol, measured a t 0" and 35" ( 1 7 ) . The enzyme was incubated for 30 min under each set of conditions and then assayed 35", for residual enzymic activity. ( A ) O", (A)0" plus 2-mercaptoethanol, (0) and (a) 35" plus 2-mercaptoethanol. The buffers used were for p H 4.0 and 4.5, sodium acetate; for pH 5.0-6.5, sodium succinate; for p H 7.0-8.5, tris-HCI; and for pH 9.0-10.5, sodium glycinate; all a t 0.1M.

of prolonged treatment a t elevated temperatures (Fig. 3). At 35" inactivation begins a t pH 8.0 even in the presence of thiol. Without thiol, 4U% of the initial activity is lost after 30 min a t 0" and the enzyme becomes slightly more sensitive to denaturation a t acid pH values although not noticeably so in the alkaline range. At 35" the range of maximum stability is limited to pH 6.3-7.5, and even here 50% of the activity is lost in 30 min (17').

D. OTHERBRAIN-TYPE CREATINE KINASES The brain-type enzyme from chicken heart muscle has been purified (16,16) and found to have a similar specific activity to thc ox brain enzyme (15).The native enzyme appears to be rather more stable than the ox brain enzyme (15). A brain-type enzyme has also been isolated from ox smooth muscle by salt fractionation and electrophoresis on blocks of Sephadex G-100. Although the product was essentially homogcneous the specific activity measured spectrophotometrically in the forward direction was 100

12.

CREATINE KINASE

403

pmoles/min/mg protein (52). Stability properties were similar to the. ox brain enzyme. A rabbit brain enzyme has been isolated with a specific activity of 530 pmoles/min/mg protein measured spectrophotometrically in the reverse direction (16). This compares with a value of 720 pmoles/min/ mg protein for the chicken muscle enzyme isolated by the same workers. A more detailed purification procedure recently described for the rabbit brain enzyme gave a 70% yield and a specific activity of 122 Kuby-units and 224 pmoles/min/mg protein when measured spectrophotometrically (63a). A similar purification procedure for the human brain enzyme gave a 21% recovery a t the first recrystallization stage and a slightly lower specific activity.

E. HYBRIDCREATINE KINASES Preparation of the MB hybrid creatine kinase from the purified muscle and brain creatine kinases of ox and human has been carried out in vitro by using approximately 4 M guanidine, pH 7.5, as the subunit dissociating agent. After recombination of the subunits by dilution and dialysis, the three isoenzymes are separated, each in about 20% yield by DEAE cellulose chromatography (6%). The specific activities obtained are similar to those of the parental forms.

IV. Substrate Specificity

A. GUANIDINE SUBSTRATES AND THE ORGANIZATION OF THE CREATINE BINDING SITE

Of the seven known naturally occurring substrates for guanidine phosphotransferases only glycocyamine (Fig. 4,II) will also act as a substrate for rabbit muscle creatine kinase. The only other phosphoryl group acceptor known until recently was N-ethylglycocyamine (7.9). Phosphoarginine and phosphotaurocyamine bind to the enzyme as inhibitors but are not substrates (80). The development of new synthetic procedures (81) has led to the list of guanidinc substrates being greatly expanded. The formulas of these compounds are shown in Fig. 4, and the reaction rates 80. E. James and J. F. Morrison, BBA 128, 327 (1966). I,. Greenleaf, and G. L. Kenyon, JACS 93,5542 (1971).

81. G. I,. Rowley, A.

404

D. C. WATTS

7% -+/.mi,

-0,C -C€&-N-C< t (1) R = CH, (creatine) (n) R = H (III) R = C,H, (IV) R = CSH,

I 110 ‘H O

(IA) (phosphocreatine)

CHS .sNH, - O-P-CH,-N-=X! 11 + 0

-o,c-cH,-N--C:

+

NH,

BNH,

I

R

(VIII) R = H (JX)R = CH,O (X)R = HO

-0,C-CH-N-C I

CH3 g N+H ,

cHs

+NH,

(XI)

- 0,C(XW

H37H3c>NH CH,-N=C:/+ *N%

(xv)

FIG.4. Guanidine substrates of rabbit muscle creatine kinase and some phosphorylatcd products (62). Names of the compounds are given in Table V. relative to creatine are listed in Table V. The position of phosphorylation of compound (V) is the free amino group as shown in compound (VI). This made possible the inference (81) that creatine is phosphorylated on the guanidino nitrogen atom trans to the methyl group (Fig. 4, IA). Increasing the length of the side chain on the secondary amino nitrogen atom of the guanidino group [compounds (111) and (IV) J results in a progressive decrease in activity suggesting that steric hindrance may be involved. However, glycocyamine (11),bearing only a hydrogen atom at this position, is also less active as a substrate, supporting the

TABLE V RELATIVE

Substrate No. in Fig. 4 (11 (V) (VIII) (111) (XI) (11) (IV) (XW (XIV) (IX) (VW (XIII) (X) (XV)

ANALOGSOF CREATINE B Y RABBIT MUSCLEKINASEA N D KINETICCONSTANTS OBTAINED FROM ENZYME A N D NMR MEASUREMENTS~

1tATES O F PHOSPHORYLATION OF

Substrate Creatine (N-methyl-N-amidinoglycine) 1-Carbox ymethyl-2-iminoimidazolidme N-Methyl-N-amidinoaminomethylphosphinicacid Negmine (N-ethyl-N-amidinoglycine) D ,cN-Methyl-N-amidinoalanine Glycocyamine (N-amidinoglycine) N-Propyl-N-amidinogly cine D-N-hidmoproline N-Methyl-N-amidino-p-alanine Methyl hydrogen N-methyl-N-amidinoaminomethylphosphate 1-Carboxymethyl-24minohexahydropyrimiduie L-N-hidmoproline N-Methyl-N-amidinoaminomethylphosphonicacid N-Methylamidino-N-methylglycine

yoof initial KD [NOS-]+ (1/Tl)Q velocity with Relative K , creatine vmaX (mM) (mM) ( m ~ ) ( 1 / ~ 1 ) T (Ref. 83) (Ref. 83) (Ref. 83) (Ref. 83) (Ref. 83) (Ref. 81) (100) 31 13 7.1 4.9 1.0 0.68 0.65 0.19 0.092

(100) 90 27 32 24 10 1.0 0.9 0.3 1.2

5 25 50 47 71 72 53 100 230 500

0.08 1.2 2.2 0.85 1.54 3.4 5.6 2.3 10.5 200

0.5 0.5

-

0.32 0.33 0.57 0.57 0.52 0.6 0.64 0.4 0.7 -

0.078 0.006 0.003

n.d.b n.d.b

-

-

4.0 100

-

5.0

0.56

n.d.b

-

100

-

-

1.0 0.8 4.0 5.0 5.0 10.0

2 w

$

-

a The data of reference 81 were obtained at pH 9.0 (forward reaction), 30" using the pH-stat assay (64); those for reference 83 were obtained spectrophotometrically (66)at pH 9.0,l". Vmx and K,,, were obtained from double reciprocal plots of measurements made with M@+as the activating ion. The dissociation constant ( K D )was obtained by analogous plots of the decrease in the longtitudmal relaxation time of water protons obtained from pulsed NMR messurements m a function of guanidine concentration in the presence of enzymeMnADP with and without added nitrate ions. TIT and TIQare the proton relaxation r a t e measured in the presence of the enryme~ MnADP and the guanidine-enzyme-MnADP complexes, respectively, both in the presence of nitrate ions. Hence, ( l / T i ) ~ / ( l / Z ' i )is a measure of the change in the state of organization around the bound metal ion in the enzyme-MnADP-nitrate complex upon adding ) ~ is a rough guanidine. [NOS-]i is the concentration of nitrate ions required to induce half the maximum effect on ( l / T l ) ~ / ( l / T ~and measure of the binding constant for the nitrate ion in each guanidine-enzyme-MP-nitrate complex. b Here n.d. stands for not detectable.

s

406

D. C. WA'M'S

suggestion (82) that interaction of the methyl group with the enzyme plays a specific role in the catalytic process. More detailed kinetic and nuclear magnetic resonance studies with these guanidine compounds have confirmed and extended these findings (83,Table V). The importance of the planarity and correct orientation of the guanidino group for catalysis is emphasized by the decrease in catalytic activity with the puckered ring structure (VII) as compared with (V). The additional kinetic data (Table V) show that in the presence of nitrate ions (VII) binds to the enzyme as well as the more active acyclic analogs, N-propyl-N-amidinoglycine (IV) and glycocyamine (11),and indicate that distortion of the primary nitrogen atom out of planarity with the rest of the guanidino group greatly decreases the possibility for interaction with the terminal phosphoryl group of ATP (83). I n contrast, the complete loss of activity that occurs in going from (V) to (XV) simply by cleaving the ring between the adjacent methylene groups may be interpreted solely in terms of steric hindrance within the iV-methylamidino-N-methylglycine molecule resulting in a negligible amount of productive binding to the enzyme (83). Similarly, as the bulk of the molecule is increased a t the carboxyl end in compounds (VIII), ( I X ) , and (X) the ability to act as a phosphoryl acceptor decreases rapidly. The difference in rate between (IX) and (X) is not thought to result from a difference in inductive effect of the altered substituent because phosphinic and phosphonic acid monoesters have similar pK, values and both are much stronger acids than are carboxylic acids. The low reactivity of (VIII) may be partly explained by the fact that the molecule would exist largely as the dianionic form under assay conditions a t pH 9.0 (81). Lengthening the carbon chain as in (XIV) also increases K , and decreases V,,,, by large factors. From this work a picture of the creatine binding site emerges as a narrow slot in the surface of the enzyme into which the slim creatine molecule is bound with its planar guanidino group orientated such that the primary amino group trans to the N-methyl group is very precisely aligned toward the incoming phosphoryl group of the nucleotide substrate. The finding that l-carboxymethyl-2-iminoimidazolidine (V) has a V,,,,, only slightly less than that of creatine (Table V) suggests that the positions and angles of the atoms fixed by the ring structure are very close to those adopted by creatine in the enzyme-substrate complex (83). The D- and L-proline derivatives, (XII) and (XIII), tell us a little about the probable orientation of the carboxyl group relative to the planar part of the molecule; the arrangement corresponding to the L configuration 82. E. James and J. F. Morrison, BBA 128, 327 (1966). 83. A. C. McLaughlin and M. Cohn, JBC 247, 4382 (1972).

12.

407

CREATINE KINASE

results in a substrate, albeit a poor one (Table V) , while that corresponding to the D configuration does not. The role of the creatine methyl group is partly to aid the orientation of the guanido group and, simultaneously, by its specific interaction with the enzyme-MgATP complex to trigger the conformational change that initiates the transphosphorylation process. The nature of this conformational change will be discussed in detail in Sections VI and VII.

B. NUCLEOTIDE SUBSTRATES AND RELATED INHIBITORS Although the true substrate for the enzyme is the metal-nucleotide complex, the limited evidence available on the effects of Mg2+,Mn2+, or no metal ion (63, 84, 85) suggests that while the metal ion may affect the extent of binding i t does not alter the specificity for the nucleotide substrate. The nucleotide substrate may be considered as having three parts, base, sugar, and phosphate chain. The nucleoside triphosphate is essential for the forward reaction and the diphosphate for the back reaction (73). However, AMP, adenosine, tripolyphosphate, pyrophosphate, and orthophosphate all bind to the enzyme. Some inhibitor constants are listed in Table VI.

INHIBITOR

TABLE VI CONSTANTS FOR MgADP AND RELATEDCOMPOUNDS RABBITMUSCLECREATINEKINASE

Variable substrate:

MgATP" pH 9.0 Ki (mM)

Inhibitor:

MgADPb pH 8.0

k't ( m M )

(I ~~

+ E-PCr

$ I-EPCr)

WITH

MgADPb pH 8.0 Ki (mM) (I E I-E)

+

~

ADP AMP Adenosine Phosphate Pyrophosphate Tripolyphosphate Reference

0.27 7.0 7.0 13.0 11.0 8.0 86

11.12 f 3.34 29.6 f 1 . 4 3.97 f 1 . 2 86

2 . 0 8 f 0.32 49.7 f 22.0

-

1 . 5 9 f 0.25 86

a Measured in the presence of 24 mM creatine, 50 mM glycine, and excess magnesium acetate at 30OC. b Measured in the presence of 10 mM phosphocreatine, 100 mM N-ethylmorpholine HCl and 1 mM excess of Mge+ over the ADP concentration a t 30°C.

84. W. J. O'Sullivan and M. Cohn, JBC 241, 3116 (1966). 85. T. Nihei, L. Noda, and M. F. Morales, JBC 236, 3203 (1961).

408

D.

C. WATTS

TABLE VII KINETICCONSTANTS FOR VARIOUS NUCLEOSIDE DIPHOSPHATES AND FOR PHOSPHOCREATINE WITH RABBITMUSCLECREATINE KINASEAT 3O"a.b

Substrate

Michaelii constant K , (mM)

MgADP MgdADP MgIDP MgCDP MgUDP MgGDP

0.05 0.08 6.3 8.0 10.7 1.4

f f f f

Phosphocreatine in the presence of MgADP MgdADP MgIDP MgCDP MgUDP MgGDP

2.9 4.9 61.9 53.2 37.8 17.0

f f f f f f

0.01 0.01 1.3 1.8 f 6.3 f 0.5

0.3 0.5 11.3 10.9 22.8 5.1

Dissociation constan t Ki (mM) 0.17 0.21 1.1 1.9 3.9 1.4

8.6 13.4 10.0 12.7 13.7 19.2

f f f f f f

0.02 0.04 0.1 0.2 1.1 0.3

Ir,,, relative to ADP = 100 (100) 39.9 f 0 . 1 29.3 f 0 . 4 9.1 f 0.1 9.6 f 0.4 3.85 f 0.05

f 1.3 f 3.4 f 1.4 L- 1 . 5 f 3.6 f 5.1

Data from James and Morrison (86). Measurements were carried out in the reverse direction in 100 mM N-ethylmorpholine HC1, pH 8.0 and with a 1 mM excess of Mg2+ over the nucleotide concentration. a

Limited variation of the sugar has a relatively small effect on either enzyme velocity or dissociation constant of the metal-nucleoside diphosphate complex, but when the nature of the base is varied the affinity of the complex for the enzyme decreases by an order of magnitude as compared with MgADP. Kinetic constants for a series of diphosphates are listed in Table VII showing that specificity for the base is very broad (86). Since these were determined using the back reaction in which the creatine concentration remains low relative to its K , the measured values are unlikely to be significantly affected by the presence of a chloride containing buffer (see Section V1,E). Other data suggesting that the maximum enzyme velocity is independent of the type of nucleotide used (86) are probably wrong. The nucleotide specificities of creatine kinases from other sources are compared in Table VIII (IS,47, 87). The data are only semiquantitative since measurements were made with a single nucleotide concentration. The BB enzyme from ox smooth muscle and the monkey M M en86. E. James and J. F. Morrison, JBC 241, 4758 (1966). 87. J. A. Underhill and D. C. Watts, unpublished results (1972).

12.

409

CREATINE KINASE

TABLE VIII RELATIVE NUCLEOTIDE SPECIFICITIES OF SOMECREATINE KINASES

ox

Chicken Substrate

Rabbit brain (BB)

Brain (BB)

Muscle (MM)

MgADP MgdADP MgIDP MgCDP MgUDP MgGDP MgdGDP MgXDP Nucleotide concn. (mM) Reference

(100) 82 17 6 5 0.5 16

(100) 15 1.5 0 0.5 16

(100) 59 4

-

1 4 1.5 0.5 16

smooth muscle (BB)

Monkey muscle (MM)

(100) 50 0 0 -

(100) 0 -

0 -

-

0 10

47

87

zymes are notably different in being absolutely specific for adenine as the base. The molecular organization of the nucleotide binding site is discussed in Sections VI1,B and C.

V. The Activating Metal Ion

For the rabbit muscle enzyme MgZ+,Mn”, Caz+,and Coz+have been found to act as activators while Ba2+, Srz+, BeZ+,Ni2+, Cr2+, Cd2+, and Zn2+were either inactive or inhibitory (88). Low levels of activity with Srz+and Ba2+ have recently been reported (89). For the enzyme from blue hare skeletal muscle Fez+was also found to activate while Ba2+,Ni2+, A12+,Cdz+,and Zn2+were either inactive or inhibitory (90). I n passing it might be mentioned that some other phosphagen kinases have been found not to be activated by Ca2+, Coz+, or Fez+ (reviewed in reference 90). A comparison of the activity of rabbit muscle creatine kinase as a function of Mg2+,Mn2+,and Ca2+ion concentration as measured by the pH-stat assay (65) is shown in Fig. 5 (90a) for the forward and reverse reactions. These data indicate the complex relationship between the nu88. W. J. O’Sullivan and J. F. Morrison, BBA 77, 142 (1963). 89. J. S. Taylor, A. McLaughlin, and M. Cohn, JBC 246, 6029 (1971). 90. D . C. Watts, in “Studies in Comparative Biochemistry” (K. A. Munday, ed.), p. 162. Pergamon, Oxford, 1965. 90a. E. J. Milner-White and D. C. Watts, unpublished data (1971).

410

D.

C. WATTS

I60 I-

FIG.5. Comparison of the effects of different activating metal ions on the activity of rabbit muscle creatine kinase. (A) Forward reaction at pH 9.0 using ATP, 4 mM; creatine, 40 m M ; NaAc, 100 mM; and cysteine, 1 mM in the pH-stat assay (6.5).The metal ions, added as the acetate were ( A ) Mn", (0) Mg", and ( 0 )Ca". The ATP concentration is equal to a pMe of 2.4. (B) Reverse reaction a t pH 8.0 using ADP, 1 m M ; phosphocreatine, 10 m M ; NaAc, 100 mM; and cysteine, 1 mM with added metal ions as in (A). The ADP concentration is equal to a pMe of 3.0 (DOU).

c1eotide:metal ion ratio and the enzyme activity. It is noticeable that the activities with both Mn2+ and Mg2+ ions initially follow the low activation curve obtained with Ca2+ ions but suddenly break away to follow a much steeper course of activation, the break point occurring a t a tenfold lower concentration for MnZ+than for Mg2+ions. The situation becomes more complex as the concentration of free MnZ+rises, apparently resulting from formation of M n ( 0 H ) which can be particularly troublesome with the pH-stat assay (forward direction) in which the addition of alkali causes transient local increases in pH. This explains the sharp descending limb of the pMn2+ curve a t high metal ion concentrations and also why the activity with Mg2+is greater than that with MnZ+when high ATP concentrations are used (91). Other experiments (91, 92) suggest that Mn2+ ions may actively inhibit creatine kinase 91. D. C. Watts, BJ 89, 220 (1963). 92. A. H. Ennor and H. Rosenherg, BJ 57, 203 (1954).

12.

411

CREATINE KINASE

even when the concentration is less than that of t.he ATP, particularly if traces of oxidizing agents are present. I n a detailed comparison of t.he nature of Mg2+,MnZC,and Ca2+ ion activation (93) in which the concentration of free metal ion was held a t 1 mM as recommended by Cleland (72) (see Section III,A,2 and Volume I1 of this work, p. 1) the more erratic nature of the results with MnZ+ions was notcd but not understood. Howcver, useful information was obtained and i t was found that the maximum velocity of the reaction with thc metal-ADP complexes decreased in the order MgADP > MnADP > CaADP (Table IX and cf. Fig. 5 ) . Thus V,,,,, decreafies with increased ionic radius of the unhydrated metal ion (there is now very good evidence that the metal ion is unhydrated when bound to the enzyme as discussed in Section VI1,B) while the affinity of metal-ADP for the enzyme-phosphocreatine complex increases with increasing size of the unhydrated metal ion (Table IX). It was concluded that the role of the metal ion in the PCr-E-MeADP complex was to polarize the N-P bond of phosphocreatine in the transphosphorylation reaction. It was also recognized that binding phosphocreatine to the enzyme could alter the conformation of the metal-nucleotide binding site and, consequently, the binding of a particular metal-nucleotide complex. The finding that Mn2+ does not complex with the y-phosphoryl of ATP in the Cr-EMnATP complex or with the nitrate ion in the Cr-E-MnADP-nitrate complex (see Sections VI1,B and C) (94)diminishes the probability of the first suggestion. The observation that while the dissociation constants of the nitrate ion from the Cr-E-MnADP and Cr-E-MgADP complexes are both less than 1.0 mM,those for chloride and bromide are 4 and 11 COMP.\RISON OF VELOCITY AND

TABLE IX EFFECTS OF DIFFERENT METAL IONSON THE MAXIMUM KINETIC CONSTINTS OF R.%nnITMUSCLECREATINF. KINASP THE

Value of constant wit'h Kinetic parameter

Ki. (mill) K , (mM) Kib (mill) Kb (mM) Vmnx(amale/mg/min 1

Equilibrium

+

EMADP E MADP PcEMADP PcE + MADP PcE*Pc+ E PcEMADP PC EMADP

+

At 30" in 0.1 M N-ethylmorpholine HC1 buffer, pH 8.0. 93. J. F. Morrison and M. L. Uhr, BBA 122, 57 (1966). 94. G. H. Reed and M. Cohn, JBC 247, 3073 (1972).

Ca 0.06 0.09

6.6 1.0 33

Mn

Mg

0.07 0.11 0.016 0.07 2.5 6.5 0.7 3.4 94 207

412

D. C. WATTS

mM with the magnesium complex (65) but 10-20 mM and too high to measure, respectively, with the manganese complex (94) draws attention to the importance of the precise orientation of substrates in a catalytic mechanism known to involve substrate-dependent conformational changes. High concentrations of free Mg2+ inhibit noncompetitively with respect to both MgADP and phosphocreatine (95). On the other hand, Ca2+and Mn2+ inhibit phosphocreatine noncompetitively but MeADP uncompetitively under similar conditions (93). The only rational explanation appears to be that because of their larger ionic radii Ca*+and Mn2+are unable to slot into a site on the enzyme that will accommodate the smaller Mg2+ion until binding of the metal-nucleotide complex produces a conformation change that sufficiently opens up the site. The mechanistic role of the activating metal ion is discussed in detail in Section VII.

VI. Enzyme Kinetics

A. ACTIVEFORMS OF

THE

SUBSTRATES

1. Nucleotide

The first kinetic studies on the purified rabbit muscle enzyme (7'3) showed a striking dependence upon the Mg2+ concentration that was interpreted as indicating that MgADP and MgATP were the active substrates while the free nucleotides were inactive. Product inhibition studies suggested separate binding sites for the nucleotide and creatine or phosphocreatine. A reinvestigation of the dissociation constants for ATP and the MgATP complexes (96) gave pK, values of 3.93 and 6.97 and a pKM, value of 4.88. This means that in Scheme 1, which shows the interrelationships of the possible complexes between ATP, Mg2+,and H+, those species joined by weak arrows do not usually exist in sufficient concentration over the pH range a t which the enzyme is active (above pH 5.5) to be of significance. The important ATP species are MgATP2+,ATP4-, and HATPS-. Equilibrium dialysis and gradient sedimentation studies showed that, while MgATP*- and ATPI- were bound to similar extents in the absence of the guanidine substrate a t pH 7.9, and 3"C, free Mg" W. J. O'Sullivan, BJ 94, 221 (1965). 96. W. J. O'Sullivan and D. D. Perrin, BBA 52, 614 (1961).

95. J. F. Morrison and

12.

CREATINE KINASE

413

ions were bound only to a very slight degree indicating that the formation of an E-Mg complex as an intermediate in the formation of the E-Mg-nucleotide complex was extremely unlikely (97). The true substrate of the enzyme should be considered as the magnesium-nucleotide complex although temperature-jump studies indicate that a minor pathway in which the Mg2' ions bind to the enzyme-nucleotide complex cannot be completely excluded (68). By making measurements a t p H 9.0 so that the concentration of HATP3- was negligible, it was then possible to show conclusively that the initial enzyme velocity correlated with the calculated concentration of MgATP'- in solution when the Mg2+ concentration was varied a t several fixed concentrations of ATP and vice versa (63). Appreciable deviation from theoretical expectations occurred only if the total ATP concentration was very much greater than the total Mg" concentration, when the concentration of ATP4- became appreciable and caused slight inhibition. It is mechanistically significant and probably physiologically important that ATP4- is such a poor inhibitor of creatine kinase that it can be ignored for most experimental purposes. For this reason many studies using lower concentrations of Mg2+ ions, and frequently with Mg*+:ATP in the 1 : l ratio originally found to give optimal activity (73), remain interpretable. This situation does not apply when the pH is lowered so that the concentration of HATP3- increases, for HATP3- is a potent inhibitor with a Ki of 0.3 mM (63). Further, while the stability constant of MgATP2- is about 90,000 M-' that for MgHATP- is only 800M-' (96),thereby favoring the tendency to form HATP8- at the lower p H limits of kinase activity. Consequently, it is experimentally difficult to determine whether MgHATP- is an inhibitor, and this remains unknown. Similarly, for the back reaction, the true substrate is MgADP- but the inhibitory species is ADP3-,while HADP2- does not appear to react with the enzyme (95, 98). 97. S. A. Kuby, T. A . Mahowald, and E. A. Noltmann, Biochemistry 1, 748 (1962). 98. T. Nihei, L. Noda, and M. F. Morales, JBC 236, 3203 (1961).

414

D. C. WATTS

2. Gmnidine

Neither creatine nor phosphocreatine shows any dependence for activity on the MgZf ion concentration, and these react as the free guanidine compounds. Creatine occurs in solution as the zwitterion. Upon phosphorylation only a single proton is given up (99) as a result of the rearrangement of phosphate ionizations. Consequently, the guanidino group must retain its positive charge giving phosphocreatine a net negative charge of two [Eq. (1)1. B.

SUBSTRATE

BINDING

Under some conditions rabbit muscle creatine kinase appears to show a very simple pattern of substrate binding with both nucleotide and guanidine substrate binding to the enzyme in a manner that is independent of the concentration of the second substrate. However, careful analyses show the situation t o be more complex. To eliminate the possibility of having uncomplexed forms of ATP in the assay it has been recommended (72) that conditions be adjusted to give a constant concentration of free Mgzt ions of 1 mM (100). Under these conditions it is found that the Michaelis constant for one substrate is a function of the concentration of the second substrate. The observed Michaelis constant decreases as the concentration of the second (fixed) substrate is raised, giving a family of Lineweaver-Burk plots that intersect above the abscissa and to the left of the ordinate for all four substrates (101). These results indicate that the reaction mechanism is sequential ; thus, both substrates react with the enzyme before either product dissociates but may be either of the ordered, Theorell-Chance or the rapid-equilibrium random type of mechanism (102) since they have the same form of initial velocity equation. The rapid-equilibrium random mechanism may be depicted by Scheme 2 where A, B, P, Q, and E represent MgADP-, phosphocreatine, creatine, MgATP2-, and enzyme, respectively. The kinetic constants Ka,K b , K,, and K , are derived from 99. M. Gellert and J. M. Sturtevant, JACS 82, 1497 (1960). 100. A small excess of Mg2+ ions is probably not inhibitory for the forward reaction, and the original observation of such inhibition (73) may be explained by the

sulfate used as a counterion since no inhibition is observed when sulfate is replaced by acetate (63; see also Section V). For the back reaction a five- to tenfold excess of Mg'+ over ADP may be required to ensure that all the ADP is converted to the metal complex. However, this excess is not only inhibitory but also affects the combination of phosphocreatine with the enzyme (101). I n this circumstance a small concentration of free ADP'- is the lesser of the two evils and a 1 m M excess of Mgzt ions is a reasonable compromise.

12.

415

CREATINE KINASE EMgADP

PCr

EhQATP

CrE

PCrE

SCHEME 2

the points of intersection of the primary plots while Ki,, Kib, Kip, and Kiq, the dissociation constants of each substrate from the free enzyme, are obtained from secondary plots of l/Vmsx against l/concentration of fixed substrate used in the primary plots. The constants obtained (101) are listed in Table X, and the only reservation required is that those for the forward reaction are likely to be affected by the chloridecontaining buffer used in the assay (see Section V1,E). A correction for the uncertain value of the stability constant of MgADP used (4000M-') made no significant difference (101). Kinetic mechanisms with the same initial velocity equations may be distinguished by initial velocity measurements in the presence of one of the reaction products (100, 102, 103). A rapid-equilibrium random TABLE X KINETIC CONSTANTS FOR RABBITMUSCLECREATINE AT pH 8.0 AND 3 0'0 KINASEMEASURED Value of constant

Kinetic constant

Equilibrium

+ + + + + + +

MgATP E eEMgATP Cr EMgATP F! CrEMgATP Cr E CrE MgATP CrE F! CrEMgATP MgADP E 2 EMgADP PCr EMgADP PCrEMgADP PCr E PCrE MgADP PCrE F! PCrEMgADP

+

a

~

Data of Morrison and James (101).

101. J. F.Morrison and E. James, BJ 97, 37 (1965). 102. W.W. Cleland, BBA 67, 104 (1963). 103. R. A. Alberty, JACS 80, 1777 (1958).

(d) 1.2 f 0.3 6.1 f 1.0 15.6 f 4.9 0.48 f 0.10 0.17 f 0.02 2.9 f 0.3 8.6 f 1.3 0.05 f 0.01

416

D. C. WATTS

mechanism with two dead-end complexes, Cr-E-MgADP and PCr-EMgATP, would show a pattern of product inhibition in which, for the forward reaction, MgADP would be competitive with MgATP and noncompetitive with creatine, while phosphocreatine would be competitive with creatine and noncompetitive with MgATP. A similar situation would hold for the back reaction. This has been found to be so (73, 101) together with evidence for the two dead-end complexes (101). However, since the Cr-E-MgADP dead-end complex would have been strongly stabilized by the chloride ions in the assay system (65) precise values for the kinetic constants must await further investigation. Additional evidence for the rapid-equilibrium random mechanism comes from the finding that by using equilibrium dialysis or gradient ultracentrifugation all four substrates bind separately to the enzyme (97) and that in the absence of guanidino substrates no phosphate exchange occurs between [azP]ADP and ATP ( 6 3 ) . By following the isotope exchange in equilibrium mixtures of substrates the rate of phosphoryl transfer between creatine and phosphocreatine, and ADP and ATP were found to be approximately equal (104). Similar results were obtained when Ca2+or Mn2+were used as the activating metal ion instead of Mg2+ (105).Illustrative data for one metal ion concentration are shown in Table XI (see also Section V1,F) from which it may be concluded that changing the nature of the activating metal ion does not alter the reaction mechanism. The situation becomes somewhat more difficult to interpret when other nucleotide pairs are used instead of ATP and ADP (Table XI). The CDP-CTP pair give the same exchange rates as the guanidine pair, the UDP-UTP pair is slightly different from the guanidine pair, while the GDP-GTP pair exchange a t a markedly different rate from the corresponding guanidine pair. It is also notable that the exchange rate of the guanidine pair decreases by 2-3 orders of magnitude with these nucleotides as compared with that obtained with the adenine nucleotides. Bearing in mind the kinetic data discussed in this section and other data discussed in Section VII, it is clear that the magnitude of the exchange rate is closely allied to the conformational changes induced in the enzyme as a result of substrate binding. It would be of interest to compare exchange rates with the new range of guanidines that has recently become available (Fig. 4 ) . I n discussing their exchange data the authors (105) are disturbed that although the exchange rate with the CDP-CTP pair indicates a rapid-equilibrium random mechanism their earlier data 104. J. F. Morrison and W. W. Cleland, JBC 241, 673 (1966). 105. J. F. Morrison and A. White, Eur. J. Biochem. 3, 145 (1967).

12.

417

CREATINE KINASE

COMPARISON

TABLE XI EQUILIBRIUM EXCHANGE RATESWITH DIFFERENTMETALIONS A N D DIFFERENT NUCLEOTIDE PAIRSAFTER ENZYMIC ESTABLISHMENT OF EQUILIBRIUM^^^

OF

Exchange ratec Creatine-phosphocreatine NDP-NTP 109 x nmoles/min/pg creatine kinase

Metal ion

Nucleotide pair

Mg2+ Cazf Mn2+

ADP-ATP ADP-ATP ADP-ATP

1500 f 130 (2) 980 f 140 (1) 1700 f 120 (2)

1460 f 40 (2) 1100 f 20 (1) 2150 f 120 (2)

Mg"+ Mg Mg

CDP-CTP UDP-UTP GDP-GTP

2.7 f 0.2 (3) 3 . 5 f 0.3 (4) 1.7 f 0 . 1 (3)

2 . 5 f 0 . 1 (3) 2.3 f 0 . 2 (4) 4 . 8 f 0.2 (3)

At 30' in 0.1 M triethanolamine HC1 buffer, pH 8.0 (106). Equilibrium was established by adding various amounts of enzyme (5-600 pg/ml) to reaction mixtures containing nucleoside triphosphate, 4 pmoles; nucleoside diphosphate, 20 pmoles; phosphocreatine, 20 pmoles; and metal ion, 20 pmoles in a total volume of 4 ml. The position of equilibrium was established by assaying for creatine a t 10 min intervals. Samples were then taken for the addition of radioactive substrates, 0.5 ml for the labeled nucleotide and 1.0 ml for the labeled creatine. Weighted mean values for number of experimenta given in parentheses. a

b

0

(867,contrary to expectations, gave different values for the dissociation constant of C D P as a n inhibitor and as a substrate. This anomaly is very likely a consequence of the C1--containing buffer used stabilizing the Cr-E-MgCDP complex (see Section VI,E) in their assay mixtures. Consequently, the Ki for MgCDP (3 mM) is lower than the K , (8 mM) for dissociations from what was assumed to be the E-PCr complex, while the corresponding dissociations from the free enzyme (1.9 and 2.5 mM, respectively) were not significantly different. It was found that U D P and G D P behaved in the same way (86). Taken a t face value the exchange data for UDP and G D P indicates that with these substrates the reaction is not rapid-equilibrium random, but again chloride-containing buffers were used and where the exchange rate is relatively small the possibility remains that such adventitious influences may alter the picture. The authors (105) were inclined to blame the high enzyme concentration that was required for these requirements. At least for the normal substrates it can be said that these data provide full support for the mechanism of action of the rabbit muscle enzyme acting via Scheme 2 with all equilibria being established rapidly and randomly except for the interconversion of the central complexes, Cr-E-MgATP PCr-E-MgADP. By increasing the concentration of

418

D. C. WATTS

creatine and MgADP in the equilibrium mixture evidence was also obtained for the formation of the dead-end complex Cr-E-MgADP, this effect probably being enhanced by the stabilizing effect of chloride ions added in high concentration with the buffer. Evidence for the formation of the other dead-end complex, PCr-EMgATP, was ambiguous as a result of the difficulty of controlling the concentrations of other inhibitory species (104). Comparison of the enzymes from ox brain and ox skeletal muscle, using the pH-stat assay (64)in the presence of 1% albumin but without any inhibitory anions being present, has produced interesting results (106). The data were found to be consistent with the rapid-equilibrium random mechanism in which interconversion of the central complexes represented the rate-limiting steps (Scheme 2) although the analysis differed slightly from that used for the rabbit enzyme in that independent binding of the substrates was not assumed. Values for the derived kinetic parameters are compared in Table XII. At pH 8.8 the most notable feature is the difference between the constants for the binding of a substrate in the ternary complex (I&) as compared with that for the binary complex (&). For the muscle enzyme this is small but for the brain enzyme the difference is an order of magnitude or more. In this respect the ox muscle enzyme is, like that from rabbit muscle (Table X), in accord with the proposal that the binding of one substrate to the enzyme induces a protein conformational change that results in tighter binding of the second substrate (101). While the ox brain enzyme exhibits this behavior to a greater degree a t pH 8.8, it becomes much more like the muscle enzyme when the pH is reduced to 7.4. This decreased difference betwen K, and Ki, a t pH 7.4 has reasonably been inferred to reflect a “tightening up” of the protein three-dimensional structure at the lower pH (106). Since this is accompanied by, if anything, a further lowering of the equilibrium constants, it would seem that the organization of the substrate binding sites inherent in the three-dimensional orgsnieation of the free enzyme is close to the conformations adopted when containing bound substrate. This is indicated particularly by the constants for the reverse reaction of the brain enzyme a t pH 7.4 where K, and Ki, values are essentially the same. The decrease in pH is also accompanied by a large increase in the maximum initial velocity from 70 to 500 pmoles/min/mg, reflecting the fact that a t pH 8.8 measurements were being made well away from the optimum (cf. Fig. 1). 106. H.K.Jacobs and S. A. Kuby, JBC 245, 3305 (1970).

c

h3

E

TABLE XI1 KINETICCONSTANTS FOR CREATINE KINASESFROM Ox MUSCLEA N D BRAINAT 30"

1:

1 .,1

~~

L-J

Kinetic parameter Cleland notation

Ki,

Author's notation

K1 KZ K3

pH 8.8 Skeletal muscle (MM)

Equilibrium

+ + + + + +

EMgATP E MgATP C r E e E Cr KP CrEMgATP S Cr EMgATP KQ K4 CrEMgATP g CrE MgATP Ki, Ki EMgADP E MgADP Kib KO PCrE E PCr Kb Ks PCrEMgADP = EMgADP PCr K. K8 PCrEMgADP = PCrE MgADP V , (forward) pmole/mg/min Vmm (reverse) pmole/mg/min Reference KdP

+

+

0.97 53.0 21.0 0.78 0.17 45.0 23.0 0.09

f 0.18 f 38.0 4.0 f 0.56 f 0.02 f 4.2 f 6.3 f 0.04 220 130 106

pH 8.8 Brain (BB) 0.93 29.0 3.7 0.13 0.12 20.0 2.0 0.01

f 0.14 f 11.0 f 0.4 f 0.05 f 0.01 f 6.0 f 0.1 f 0.003 200 70 106

pH 7.4 Brain (BB)

pH 8.6 Smooth muscle (BB)

0.18 f 0.03 13.0 f 5.0 3.0 f 0.4 0.05 f 0.015 0.04 f 0.003 0.67 f 0.14 0.52 f 0.12 0.03 f 0.003 120 510

0.75 2.20 0.58 0.20

106

47

420

D. C. WATTS

The less flexible ox muscle enzyme has the higher Vg:ye of 130 a t pH 8.8 suggesting that destabilization of the protein structure may well contribute to the fall of activity a t high pH. For the hare and rabbit muscle enzymes decrease of activity in the forward direction only occurs when there is a manifest disruption in protein organization (Fig. 2). As will be seen from Section VII,B the actual movement of protein side chains needed to bring about such profound changes in substrate binding and catalytic activity need not be large and may indeed be so small as to be extremely difficult to quantify. However, other experiments on the enzymes from dogfish (33) and monkey (107)indicate that such changes are probably a feature of all creatine kinases and probably of all phosphagen kinases, the extent to which they are manifest being a feature of species variation. Some experiments on the rabbit muscle enzyme (98)suggest, in contrast to the work described above, that the binding of one substrate is not affected by the prior binding of the second substrate to the enzyme. The K,,values were also inferred not to vary with pH. This behavior remains to be checked since results with the ox brain enzyme do not support this finding (106). The question of whether the brain-type enzyme isolated from smooth muscle is a different isoenzyme from that isolated from brain also remains open. As Table XI1 shows, while the dissociation constants for MgATP are similar, those for creatine are very much lower than can be accounted for by experimental error. However, the assay systems used were different, and it is a problem with flexible enzymes that small changes in the environment may have disproportionately large effects on the kinetic constants. C. TEMPERATURE EF~FECTS The effect of temperature on the kinetic parameters of creatine kinase is poorly documented. The data are limited to a heterogeneous collection of information about the rabbit muscle enzyme and a single, fairly detailed study of the dogfish muscle enzyme from Lineweaver-Burk plots with near-saturating concentrations of the fixed substrate but using MgSO, in a 1: 1 ratio with ATP (33) as in the original study of the rabbit enzyme (7'3). The data are assembled in Table XIII. Correction for the inhibition by sulfate ion a t 30" indicates that i t is without effect on 107. J. A. Underhill and D. C. Watts, unpublished data (1972).

12.

421

CREATINE KINASE

TABLE XI11 COMPARISON OF THE EFFECTOF TEMPERATURE ON THE KINETICPARAMETERS FOR THE CREATINE KINASESFROM DOGFISHAND RABBITMUSCLE

Temp.

Dogfish0

Rabbit

Creatine MgATP Vzfozard K , (mM) K , (mM) (pmole/mg/min)

Creatine MgATP ATPa K , (mM) K , (mM) K , (mM)

0 1 3

1.2

4 7 8 16 24 30

0.8

-

-

32 38 40

-

1.5 1.7 1.5 -

2.3

-

2.3

2.9 3.6 3.5 9.7 7.7 15.0 21.0

-

23.6 52.0

Data on the dogfish from Lineweaver-Burk plots (31). By inactivation kinetics (64). By equilibrium dialysis (97). By gradient sedimentation (97). From Lineweaver-Burk plots (73). From Lineweaver-Burk plots (66). 0 From Lineweaver-Burk plots (98).

the apparent K , for creatine but lowers that for MgATP by 10-300/0 (108). The overall picture suggests that temperature has only a small effect on K , for both creatine and MgATP over the normal physiological range. 108. This correction was made assuming a Kt value for the sulfate ion of 4 mM as reported for the rabbit muscle enzyme (86). It has been suggested by Kuby and Noltmann ( 1 0 8 ~ )that this estimate may be too low because no inhibition by sulphate ions was observed in earlier experiments (73) using conditions where i t should have been obvious. A recent estimate of 30 mM for the Ki value a t pH 9.0, when MgATP was used as the variable substrate in a Lineweaver-Burk type of experiment (66) could account for the observed discrepancy. Using this value the correction obtained for sulfate inhibition when MgSOl and ATP are used in a 1 : l ratio in the assay mixture falls well within the normal range of experimental error (i,e., +5%). 108a. S. A. Kuby and E. A. Noltmann, "The Enzymes," 2nd ed., Vol. 6, p. 572, 1962.

422

D. C. WATTS

The activation energy for the dogfish enzyme was found to be about 6,400 cal/mole increasing sharply to around 20,000 cal/mole below 7" (33). This compares with a value of about 12,400 cal/mole for rabbit creatine kinase, also measured in the presence of sulfate but with neither substrate concentration near to saturating (73). The effect of temperature on the ascending limb of the pH-activity curve for the rabbit muscle enzyme is quite small, yielding a heat of ionization of about 1800 cal/mole for a group with a pK near 6.5 a t 30" in the enzyme-substrate complex ( 6 3 ) .It is speculated that this ionization is that of a phosphate group since carboxyl, histidine, sulfhydryl, or amino groups would assume this pK and AH only in very unusual circumstances. It is also pointed out, however, that the presumed "K" of the hypothetical group can actually be a complex function of many equilibrium constants describing successive steps in the enzyme reaction (109, 110).

D. ATP AND ADP

AS

INHIBITORS

As has already been mentioned (Section VI,A,l) ATP4- is said to be virtually without inhibitory effect (63, 101). This is surprising in view of the fact that the dissociation constant for E-ATP4- has been found t o be about 0.4 mM by equilibrium dialysis and gradient sedimentation studies (97), a value very similar to that of 0.3 m M reported for the manifestly inhibitory species HATP3- (63,98). A possible explanation that the dissociation constant is raised by the presence of the second substrate may be correct but is not supported by the only available data which are for the binding of ADP3- to the ox brain enzyme (105) in the reverse reaction. At pH 8.8 the dissociation constant for the binary E-ADP complex is 0.6 mM while that for the ternary PCr-E-ADP complex is 0.15 mM. At pH 7.4 the difference is in the other direction giving 0.03 and 0.08 mM, respectively. These values are only tentative but sufficiently accurate to indicate the absence of any marked effect of phosphocreatine on the binding of ADP"-. For the rabbit enzyme, assuming no interaction between substrates, values for the dissociation constants of ADP3- of 0.2-0.3 mM have been reported (63,95,98).However, ADP3is an inhibitory nucleotide species for the reverse reaction and nothing is known about the behavior of HADP2-, the species more comparable to ATP"- in the forward reaction. 109. I,. Peller rind R. A. Alberty, JACS 80, 5907 (1959). 110. T. C. Bruice and G . L. Schmir, JACS 81, 4552 (1959).

12.

CREATINE KINASE

423

E. EFFECTS OF ANIONS The first attempts to study the effect of anions on creatine kinase led to the conclusion that SO:-, C1-, and PO:- ions all acted as competitive inhibitors of phosphocreatine and noncompetitive inhibitors of MgADP (98). Hence, interaction of the anion with a site on the enzyme that normally bound the transferable phosphoryl group of the substrate was inferred. Other data indicated that the presence of chloride ions in the assay mixture could be inhibitory, but only recently has this problem been given serious attention. As it has turned out this negligence has proved particularly unfortunate because so much of the research on creatine kinase has been carried out using chloride buffers or solutions containing other anions that materially affect the interpretation of the results. One attempt (111) to investigate the effect of chloride ions gave results that were difficult to interpret because the experiments were carried out against a background concentration of 40 mM chloride used in the buffer system. It was concluded, probably correctly, that NaCl is a noncompetitive inhibitor of the reaction with respect to all four substrates and Ki values in the range 50-200 m M were reported. Difficulties arose when comparing the binding of chloride ions in the binary and ternary complexes and the Ki values obtained for chloride ions with the latter, 250-500 mM, suggested that inhibition of the ternary complexes was weak. This may be correct for the Cr-E-MgATP and PCr-EMgADP complexes, but because of the high background concentration of chloride ions the important effect on the ternary dead-end complexes was not detected. It is now realized that the readiness of creatine kinase t o form the nonphosphorylated dead-end complex, Cr-E-MgADP, permits the forination of a further complex in which certain anions sit on the site of the transferable phosphoryl group between creatine and MgADP to form an extremely stable quaternary complex ( 6 5 ) . The inhibitory effect on chloride ions in forming this complex far outweighs any other inhibitory effects that the anion may have with respect to other enzyme-substrate combinations. Small, planar anions such as NOa- are particularly effective in this respect, and the hypothesis has been presented that the anion acts by simulating a planar phosphoryl group in a stable intermediate complex inferred to occur in the course of the reaction (see Sections VI1,B and D ) . The inhibitory effects of other anions vary according to the extent to which they can act in this way. Figure 6A and B shows how the product inhibition of the forward reaction by MgADP or the back 111. E. Heyde and J. F. Morrison, BBA 212, 288 (1970).

424

D.

C. WATTS

I/[MgADP] (rnM-')

- 2 - 1

0

1 2 l/[MqATP] (rnM-')

-0.1

3

0 0.1 0.2 I/[ Creatine] (rnM-')

o 0.1 0.2 0.30.40.5 i/[Creotine phosphate1(rnM-')

-0.3-a2 0.1

FIQ.6. Comparison of the effects of anions on the product inhibition of rabbit muscle creatine kinase in the forward (pH 9.0) and reverse (pH 8.0) reactions a t 30" using the p H state assay (66).(A) Inhibition by MgADP. Double reciprocal plots obtained in the absence, solid symbols, and presence, open symbols, of 0.161 . , 0 )100 mM NaAc or (&A) 100 mM NaCI. The mM MgADP plus either ( fixed concentration of creatine was 40 mM. (B) Inhibition by phosphocreatine. Double reciprocal plots obtained in the absence, solid symbols, and presence, open . , 0 ) 100 mM NaAc, or &A) symbols, of 6 mM phosphocreatine plus either ( 100 mM NaCI. The fixed concentration of MgATP was 4 mM. (C) Inhibition by MgATP. Double reciprocal plots obtained in the absence, solid symbols, and . , 0 ) 100 mM NaAr or presence, open symbols, of 1 mM MgATP plus ether ( (A,A) 100 mM NaC1. The fixed concentration of phosphocreatine was 10 mM. (D) Inhibition by creatine. Double reciprocal plots obtained in the absence, solid . , 0 ) 100 symbols, and presence, open symbols, of 20 mM creatine plus either ( mM NaAc or (&A) 100 mM NaCl. The fixed concentration of MgADP was 0.5 mM. I n all experiments the free Mg" ion concentration was 1 mM.

reaction by creatine is greatly enhanced by C1- ions as compared with acetate ions. On the other hand, C1- has very little effect in enhancing inhibition by the products phosphocreatine or MgATP (Fig. 6C and D) . Chloride has only a slight effect on the K , for any substrate, and the Ki values for the chloride ion varies between 140 and 230 mM according to the substrate being varied in the double reciprocal plot. This is in fair agreement with the previous findings (63, 111). When the inhibitory effects of products are investigated it is found that addition of chloride,

12.

425

CREATINE KINASE

as compared with acetate, causes a profound lowering of the K i value for creatine and MgADP but not phosphocreatine or MgATP (Table XIV). These data show that the anion stabilizes the formation of the nonphosphorylated dead-end complex but not of the phosphorylated dead-end complex PCr-E-MgATP. The value of Ki for C1- in the nonphosphorylated dead-end complex is 4 mM, a 35-50-fold decrease as compared with the value toward single substrates. The corresponding value for NO,- is 0.95 mM (66). Additional information comes from an investigation of the ability of anions to facilitate protection of creatine kinase against inhibition by iodoacetamide in the presence of creatine plus MgADP, this substrate combination alone being without effect, Anions that cause protection are, in order of effectiveness, NO,- > NO,- > C1- > HCOO- > H C 0 , - 3 Br- > F- with NO,- giving almost complete protection and F- very little. They are all either planar ions or halides smaller than iodide and, for convenience, were called Class I1 anions to distinguish them from other anions with different properties ( 6 5 ) . The general order of effectiveness of the Class I1 anions in stabilizing the dead-end complex was confirmed

TABLE XIV AND INHIBITOR CONSTANTS EFFECTSOF ANIONSON THE MICHAELIS FOR CREATINE KINASEAT 3Ooa8* ~~

PH

Variable substrate

!I .0 9.0 9.0 11.0 8.0 8.0 8.0 8.0 9.0 9.0 9.0 9.0 8.0 8.0 8.0 8.0 9.0

MgATP MgATP Creatine Creatine MgADP MgADP Creatine phosphate Creatine phosphate MgATP MgATP Creatine Creatine MgADP MgADP Creatine phosphate Creatine phosphate MgATP

a

~~~~~

~~

Inhibitor -

-

MgADP MgADP Creatine phosphate Creatine phosphate MgATP MgATP Creatine Creatine MgADP

Anion (0.1M)

K, (mM)

Ki (mM)

Acetate Chloride Acetate Chloride Acetate Chloride Acetate Chloride Acetate Chloride Acetate Chloride Acetate Chloride Acetate Chloride Acetate

0.4 0.65 10.7 9.7 0.14 0.19 3.3 4.1

-

-

-

0.36 0.05 46.0 50.0 1.2 1.4 40.0 6.0 0.34

Data from Milner-White and Watta (66). The assay conditions and substrate concentrations were as described in Fig. 6.

426

D. C. WATTS

by proton relaxation rate (PRR) and electron paramagnetic resonance measurements (94). This gave, in decreasing order of PRR change NO,- > NO2-, SCN- > C1- > HCOO-. These measurements were made at 0" using Mn2+as the activating ion, whereas the previous data were obtained a t 30" using Mg2+.This difference may account for the failure to detect any PRR change with the less effective anions, HC0,-, C0,-, Br-, and F-. Electron paramagnetic resonance (EPR) measurements showed that NO,-, NO2-, and SCN- had dissociation constants of less than 10 mM with NO,- having a dissociation contant of less than 1 mM and being much more effective than the other two. If creatine was omitted from the reaction mixture the anion had no effect, confirming the action was one of specifically stabilizing the dead-end complex. The tetrahedral anions such as SO:-, HPOi-, ClO,-, As0,-, and BF,and the iodide ion form another group, the Class I11 anions, on the basis of their action on creatine kinase. They do not affect the rate of inhibition by iodoacetamide in the presence of creatine plus MgADP but may decrease the protection afforded by Class I1 anions under these conditions. Their mode of action appears to be that originally envisaged (98) by binding to the site on the enzyme occupied by the transferable phosphoryl group in its tetrahedral configuration. This might be expected to give inhibition that is competitive with respect to the phosphorylated substrates and noncompetitive with respect to the nonphosphorylated substrates. However, there appears to be a noncompetitive element in the inhibition with respect to all substrates. This is probably because of an additional effect on the conformational changes that link the two substrate binding sites when the substrates bind to the enzyme (Section VI1,B). The Class I11 anions are precluded from participating in the dead-end complex simply because they are too large to fit the gap that is left between creatine and MgADP after the substrates have bound to the enzyme, and the decrease in protection by C!ass I1 anions is caused by decreasing the binding of these substrates. Investigation of the effects of iodide, borate, vanadate and hypophosphite ions on the Cr-E-MnADP complex by PRR measurements failed to show any change when the anion was added (94). Although il? agreement with the above findings the results may not be significant because, under the conditions used, no effects could be detected with some of the less effective Class I1 anions. Other investigations of the effects of these anions that shed light on the mechanism of action of creatine kinase will be discussed in Section VII. Class I anions behave unlike Class I1 or Class I11 anions and appear

12.

CREATINE KINASE

427

to act at a different site on the enzyme. They may activate or inhibit catalytic activity. The characteristic anion of this group is acetate which was found not to affect the rate of inhibition by iodoacetamide in the presence of creatine plus MgADP as do the Class I1 anions, nor did it behave like a Class 111 anion in that even a t high concentrations it did not affect the behavior of Class I1 anions or inhibit normal enzymic activity in the forward direction of the reaction. Instead it activated the reaction by 1030% (65). Work on the enzyme from monkey muscle (112) has increased the members of this class in order of effectiveness as activators to acetate > propionate > butyrate > chloroacetate > fumarate when used a t 0.1 M concentration. Activation is associated with an enhancement of the binding of both creatine and MgATP (106).Two other anions, fluoroacetate and glycollate, have no effect a t all so that the possibility of a simple ionic strength effect is eliminated. Likewise, an earlier suggestion that acetate simply activated by chelating traces of heavy metals no longer appears tenable. These Class I anions are not competitive with either C1- or SO,- and clearly bind elsewhere on the enzyme than a t the catalytic site. The effect of Class I anions on the reverse reaction has not been widely studied, but acetate is reported to be slightly inhibitory with an apparent Ki of 350-400 mM (98, 111, 1 1 2 ~ ) . Further work on monkey muscle creatine kinase (112) revealed another group of anions which fall into Class I because they appear to act a t the same site on the enzyme but which inhibit in the forward direction of assay rather than activate. The most effective of these found so far is salicylate which inhibits the enzyme by 70% a t 50 mM concentration and is more effective than acetylsalicylate or p-amino salicylate which cause 51 and 38% inhibition, respectively. Other inhibitory anions in this class are benzoate, benzene sulfonate, and trichloroacetate. The inhibition by chloride and salicylate is additive indicating that these two inhibitory anions act in quite different ways, but acetate protects against inhibition by salicylate, suggesting a common binding site and justifying the inclusion of these organic inhibitors as a special section of Class I . The possibility that salicylate might act by chelating the activating Mg2+ion was excluded. 112. R. Chegwidden and D. C . Watts, unpublished results (1972). 112a. Other, preliminary data from the same laboratory indicate that with the rabbit muscle enzyme, for the forward reaction, acetate, butyrate, and propionate all activate, while lactate, glycollate, P-hydroxybutyrate, malonate, and nL-isoaminobutyrate all inhibit at 100 mM. For the back reaction all the anions were found to be inhibitory.

428

D. C. WATTS

F. EQUILIBRIUM OF THE REACTION This aspect of creatine kinase has not attracted much attention following the detailed reanalysis (113) of the original equilibrium studies (114). That which has been carried out, in connection with the measurement of isotope exchange rates at equilibrium (Section V1,B) supports and extends the earlier work. The reader is referred to reference 113 for the original full discussion, a little of which is summarized here to provide a link with the recent work. The original analysis (114) defined an apparent equilibrium constant

K'

(CrP)o(ADP)o (ATPI o(Cr)0 expressed in terms of the total concentration of each substrate in the reaction mixture. However, K' was found to vary with pH and the Mg2+ or Mn2+ ion concentration a t constant temperature and hence did not rigorously satisfy the requirements of a true thermodynamic equilibrium constant. Although it was appreciated a t that time that metal-nucleotide complexes were likely to be involved in the catalytic reaction this problem had not been formally resolved, and the reanalysis (113) considered three possible reactions that might be involved. These were, with Mg2+ as the activating metal ion: ATP- + Cr* MgA'WCr* MgzATP C P

=

+

+

(4 (b) + + H+ (c> It is now appreciated that only reaction (b) is enzyme catalyzed with an equilibrium constant K'a represented by

+ +

ADPOCrP*- H+ MgADPCrP*- H+ MgADPCrPz- M$+

+ +

+

(MgADP-) (free phosphocreatine) (MgATP2-) (creatine) and the pH independent equilibrium constant being given from the relationship Ka = K'a (H+).A list of Ka values determined under various conditions of temperature and pH is given in Table XV. In the more recent study (106)measurements were made only a t pH 8.0 and 30";K' (called by the authors Keq') was determined from the total concentration of each substrate a t equilibrium and K't, (called by the authors Keq) was calculated using appropriate values for the stability constants of the various complexing species (Table XVI) and assuming that over the range of metal ions used only the 1 : l metal-nu-

K'a =

113. S. A. Kuby and E. A. Noltmann, "The Enzymes," 2nd ed., Vol. 6, p. 515, 1962. 114. L. Noda, S. A. Kuby, and H. A. Lardy, JBC 210, 83 (1954).

12.

429

CREATINE KINASE

TABLE XV EQUILIBRIUM CONSTANTS OF THE MAGNESIUM-ACTIVATED CREATINEKINASE REACTION^.^

PH

Temp. ("C)

K'b (H') X 10''

7.4 8.0 8.9 9.0 8.8 9.8

30 30 30 20 38 30

3.08 f 0.38 3.02 f 0.18 2.45 f 0.27 2.52 f 0.50 2 . 4 3 f 0.38 1.28 f 0.31

Data from Kuby and Noltmann (113). Each value is the mean of 12-16 determinations. The average value of pH 7.4, 8.0 and 8.9 at 30" was 2.81 f 0.41 X lo-''. a

b

K'b

(H+) for

cleotide was formed and that Ca2+did not complex with phosphocreatine. As shown by Fig. 7, the apparent equilibrium constant decreases to a n essentially constant level as the concentration of activating metal ion is raised for Mg2+,Mn2+,and Ca2+.The corresponding K b (Keq) values are listed in Table XVII and should, if in accord with theory, be constant, irrespective of metal ion concentration, a t constant temperature and pH. This was found to be true for Mg2+as the activating metal ion, but with Ca2+values tended to rise and with Mn2+to fall as the metal ion concentration was raised. Best values for the stability constants were then chosen (listed in Table XVI) (96,116,116) so that K'b became constant. The calculated values are shown in parentheses in Table XVII. The data

::;/JL; 8 0.2

0.2

g 0.2

0. I 2

4

6

8

1

0

Total magnesium (mM)

2

4

6

8

1

Totol calcium ( m M )

0

2

4

6

8

1

0

Total manganese ( m M )

FIG.7. Effect of the total concentration of various activating metal ions on the apparent equilibrium constant (Keq') of the rabbit muscle creatine kinme reaction at, pH 8.0 (triethanolamine HCI buffer, 100 mM) a t 30". Keq' is defined in the text. The initial concentrations of ATP, ADP, and phosphocreatine were 1.0, 5.0, and 5.0 mM, respectively (106). 115. D. D. Perrin and V. S. Sharma, BBA 127, 35 (1966). 116. W. J. O'Sullivan and M. Cohn, JBC 241, 3104 (1966).

430

D. C. WATTS

TABLE XVI AND ASSUMEDSTABILITY CONSTANTS USED IN DETERMINING THE CALCULATED EQUILIBRIUM CONSTANTS LISTEDIN TABLE XVII Determined value Constant

Of-1)

MgADPMgATP2MgPhosphocreatine CaADPCaATP2MnADPMnATP" MnPhosphocreatine

4 000 70,000 40 2,200 32,000 25,000 155,000 140

Ref.

Assumed value for Table XVII ( A 4 - I )

96 96 96 96 96

96 116

3,000 20,000 200 000 ]

116

for Mgz+ are in good agreement with those determined earlier (cf. Table XV). Computation of A F O from the average equilibrium constants a t 20", 30°, and 38" obtained a t pH values of 9.0, 8.9, and 8.8, respectively, yielded values for A F O of 12.9, 13.3, and 13.7 kcal/mole, respectively (113). These values did not vary by more than kO.1 kcal/mole within the uncertainties set by the original determinations. The secondarily derived values of AH" (-0.5 kcal/mole) and AS" (-46 cal/mole/deg) VALUES

TABLE XVII EQUILIBRIUM CONSTANT OF THE CREATINE KINASEREACTION BY DIFFERENT METALIONS AT pH 8.0 A N D 3Ooasb ACTIVATED

FOR THE

Total conc. of metal ion

Keg X 1Oa

(mM) 1 2 3 4 5 6 7 8 9 10 Mean value ~~~

~

Magnesium

Calcium

Manganese

30 30 27 29 30 30 30 30 30 30 30 f 1

35 (47) 33 (44) 32 (41) 39 (48) 39 (44) 47 (52) 46 (49) 47 (49) 46 (47) 49 (50) 41f7 47f3

73 (47) 67 (44) 67 (47) 66 (51) 55 (47) 47 (45) 38 (37) 34 (34) 28 (34) 27 (27)

~~

Data from Morrison and White (106). Values for the stability constants, used are shown in Table XV. Values for Keg in parentheses were calculated using the assumed values for stability constants listed in Table XV. Keg = K'b as defined in the text. a

12. CREATINE

KINASE

431

were subject to greater variation, almost +.5 cal and + l o entropy units, respectively. The position of equilibrium of the creatine kinase reaction would be expected to be independent of the source of the enzyme. This has now been formally demonstrated in a comparison of the chicken MM and rabbit BB enzymes a t pH values of 7.5 and 8.0 with a small difference occurring in what is presumably K' (since the authors do not make this clear) a t pH 7.0 (16). Hence, it is also reasonable to conclude, as might be expected on evolutionary grounds, that the same ionic species and complexed forms of the substrates are involved in the reaction catalyzed by the enzyme from different sources.

VII. Chemical Investigations of the Enzyme Mechanism

A. REACTIVE GROUPS IN THE NATIVEENZYME ESSENTIAL FOR CATALYTIC ACTIVITY The application of group-specific reagents has made possible the detection of several amino acid side chains of creatine kinase that are essential for catalytic activity. Except for the essential cysteine residues only the rabbit muscle enzyme has been investigated. 1. Cysteine Residues and the Number of Catalytic Sites per Molecule

Classic inhibitor studies with a variety of thiol reagents rapidly proved the initial suggestion that creatine kinase was a thiol enzyme (117) to be correct, and the results with British Anti-Lewisite (2,3-dimercaptopropan-1-01) excluded the possible involvement of a dithiol (118).Isolation of the enzyme in pure state made quantitative measurements possible and the two rapidly reacting groups per molecule (64, 119-161) were soon identified as cysteine side chains by analysis of the products after reaction with iodoacetate and 2,4-dinitro-l-fluorobenzene(64). Loss of catalytic activity was found to closely parallel the rate of alkylation by iodoacetamide. Two thiol groups per molecule reacted a t exactly the same rather slow rate, suggesting that they were in 117. K. Bailey and S. V. Perry, BBA 1, 506 (1947). 118. H. Rosenberg and A. H. Ennor, BBA 17, 261 (1955). 119. R. E. Benesch, H. A. Lardy, and R. Benesch, JBC 216, 663 (1955). 120. D. C. Watts, B. R. Rabin, and E. M. Crook, BBA 48, 380 (1961). 121. R. Kassab, L. A. Pradel, E. der Terrosian, and N. Van Thoai, BBA 132, 347 (1967).

432

D. C. WATTS

identical environments which permitted the inference that there were two catalytic sites on the enzyme with one cysteine residue a t each (122), This agreed with the number of molecules of each substrate that will bind to the enzyme as determined by equilibrium dialysis ( 9 7 ) , and the picture was completed by the discovery that peptide mapping gave only half the number of spots expected from tryptic digestion, indicating that the enzyme was composed of two subunits (123). The subunits are presumed to be identical with one catalytic site on each, and the finding of only one alkylated peptide sequence in the dialkylated enzyme (Table IV) supports this view. Although the possible existence in the rabbit of muscle-type isoenzymes that have sequence differences too small for easy detection has still to be excluded, and in other animals multiple forms of the muscle enzyme may occur (see Section I,A), the features of the molecule associated with these essential thiol groups are unlikely to be affected to any significant extent since the same amino acid sequence, with only minor differences, occurs in both the MM and BB enzymes and in lobster muscle arginine kinase (Table IV and reference 124). A detailed discussion of the role of the essential thiol groups follows in Section VI1,B-E.

2. Lysine Study of the reaction of creatine kinase with p-nitrophenyl acetate suggested that a t pH 6.8 inactivation could be explained by modification of the essential thiol groups. As the p H was raised the rate of reaction with the thiols remained unchanged, but the ionization of additional groups with a pK near 9.0 caused a progressively increasing rate of inhibition of the enzyme (125). From the irreversibility of the reaction, a comparison of the second-order rate constant with those of known amino acids in synthetic peptides and the lack of an ionic strength effect, suggesting a cationic-neutral type of dissociation, the involvement of lysine side chains was inferred. Further study of the reaction (126) showed that even a t pH 7.0 the inhibition observed could be interpreted as being caused by the acetylation of lysine side chains, and the irreversible involvement of the essential thiol groups was positively excluded. The stoichiometry of the reaction with l'C-labeled reagent indicated the reaction of 2 moles/mole enzyme, and the electrophoretic 122. D. C. Watts, B. R. Rabin, and E. M. Crook, BJ 82, 412 (1962). 123. N. Dance and D. C. Watts, BJ 84, 114P (1962). 124. E. der Terrossian, L. A . Pradel, R . Kassab, and N. Van Thoai. E w . J. Biochem. 11, 482 (1969). 125. D. C. Watts, BJ 89, 220 (1963). 126. J. R. Clark and L. W. Cunningham, Biochemistry 4, 2637 (1965).

12.

CREATINE KINASE

433

mobility of radioactive peptides obtained by use of pronase, trypsin, or pepsin was unaffected by prior reaction with iodoacetate showing that the essential cysteine and lysine residues were not in the same peptide. It is presumably these lysines that are linked to the essential thiols by the bifunctional reagent described in Section II,C (cf. Table IV). Final confirmation of the existence of two lysine groups per molecule, essential for catalytic activity, was elegantly obtained by first reversibly blocking the essential thiol groups with tetrathionate and then reacting the enzyme with l-dimethyl-aminonaphthalene-5-sulfonylchloride (127). c-DNS-lysine was identified by high voltage electrophoresis and thin layer chromatography of the pronase-digested protein as the only fluorescent derivative. The presence of an equilibrium mixture of substrates (the effective component of which was the Cr-E-MgADP-NO,- complex, see Section VI1,B) appeared to protect the lysine residue from reaction with p nitrophenyl acetate but a pH-reactivity curve showed that this was because the pK of the lysine had been raised by 0.85 p H units, inferred to result from a conformational change in the enzyme, and no physical protection by the substrates had actually occurred ( 9 1 ) . Single substrates did not affect the reaction. In contrast, the reaction with dansyl chloride (127) is enhanced by nucleotide substrates but not by creatine. An equilibrium mixture of substrates gave no more enhancement than expected for a mixture of MgADP and MgATP. Unfortunately, this experiment was only carried out a t one pH so that interpretation is difficult. Kassab et al. (127)pointed out the similarity to the enhancement of the alkylation of the essential thiol by iodoacetate in the presence of MgADP and suggested that the lysine may be close to the thiol group. The latter reaction is discussed in Section VI1,D; it could be that, as with iodoacetate, the binding of nucleotides creates a new site for interaction with the dansyl chloride so that it is more favorably orientated to react with the lysine amino group. Dansylated creatine kinase gives the same difference spectrum with MgATP as the native enzyme, but that with MgADP is greatly modified in that two maxima seen a t 278 and 290 nm have disappeared. The minimum a t 254 nm remained unaltered. It is significant that dansylated arginine kinase, which behaves in the same way, does not interact with arginine as measured by difference spectroscopy (128). Unfortunately, creatine does not give a difference spectrum with either native or modified creatine kinase. 127. R. Kaasab, C. Roustan, and L. A. Pradel, BBA 167, 308 (1968). 128. C. Roustan, L. A. Pradel, R. Kassab, A. Fattoum, and N. Van Thoai, BBA 206, 369 (1970).

434

D. C. WATTS

These experiments were carried out using a 25-fold excess of dansyl chloride over enzyme. When only a twofold excess is used, the lysine residues are not labeled and fluorescent label is not incorporated into the enzyme although the essential thiols are oxidiacd, probably to thc sulfcnic acid (see Section VI1,D).

3. Histidine Diethylpyrocarbonate reacts with two histidine residues per molecule of creatine kinase, leaving the essential thiol groups unaffected (129). Enzymic activity is inhibited by 95-1000/0. Evidence for the reaction of histidine came from the known specificity of the reagent, the characteristic spectrum of N-carbethoxyimidazole and the observation that in 6 M urea 34 equivalents of reagent per mole reacted, in good agreement with the known histidine content of rabbit muscle creatine kinase (cf. Table I). Similar results were obtained if the essential thiol groups were first reversibly blocked with tetrathionate (129) and the addition of substrates was also without effect (130).The substrate-binding properties of the modified enzyme and the associated conformational changes as indicated by difference spectroscopy appear essentially unaltered (1.28). 4. Tyrosine

Addition of arginine to arginine kinase produces a characteristic difference spectrum with peaks having minima a t 239, 280, and 287 nm, indicating the perturbation of a tyrosine residue in the enzyme upon binding the guanidine substrate (128).After reversibly blocking the essential thiol group with tetrathionate (a monomer arginine kinase was used), it is possible t o react one tyrosine per mole with tetranitromethane. A difference spectrum is no longer obtained upon adding arginine, the enzyme is inhibited by 90% (131), and a marked conformational change occurs resulting in an apparent loss of a-helix content (132). I n contrast to this behavior, thiol-protected creatine kinase does not react with tetranitromethane nor does it give ii difference spectrum upon adding creatine (131).Since the two enzymes are believed to be homologous (reviewed in reference 133) and the tyrosine content per subunit 129. L. A. Pradel and R. Kassab, BBA 167, 317 (1968). 130. The experimental data of reference 189 shows 20-25% protection of rreatine

kinase by nucleotide substrates, with or without creatine, against inhibition by dirthylpyrocarbonate. However, the authors conclude that this effect is not significant. 131. R. Kassab, A. Fattoum, and L. A. Pradel, Eur. J . Biochem. 12, 264 (1970). 132. M. F. Landon, C. Oriol, and N. Van Thoai, BBA 214, 168 (1970). 133. D. C. Watts, in “Biochemical Evolution and the Origin of Life” (E. Schoffeniels, ed.), p. 150. North-Holland Publ., Amsterdam, 1971.

12. CREATINE

435

KINASE

is identical, this is a surprising finding. A speculative explanation is that the tyrosine is present in creatine kinase but masked in such a way that access by the reagent is prevented, and the difference spectrum does not appear until guanidine and magnesium nucleotide substrates are bound to the enzyme. Figure 8 shows that, in the presence of creatine, MgADP, and nitrate ions, creatine kinase gives a difference spectrum with minima a t 286 and 296 nm that could be interpreted in this way. It should prove interesting to study the crcatine kinases from other sources than rabbit muscle in this respect.

240

I

I

I

I

260

200

300

320

FIG.8. Effect of chloride ions on the difference spectrum of the Cr-E-Mg nucleotide complex with either (A) ADP or (B) IDP. The front section of a double sector cell (2 x 4.6 mm pathlength) contained, in 12 ml, rabbit muscle creatine kinase, 2.75 X W M ; in 50 mM tris-amtatk buffer, pH 8.5. The rear section contained, in 12 ml, creatine, 29 m M ; MgAa, 3.6 mM; and either (A) 0.18 mM ADP or (B) 0.18 mM I D P in the same buffer. Test and reference cells set up as above were scanned to establish the base line (curve 1) and the test cell then mixed and rescanned to give the difference spectrum of the dead-end complex without n Class I1 anion (curve 2); 0.025 ml of 4M-LiCI was then added to each compartment and the test cell remixed and then scanned to give anionstabilized dead-end complex (curve 3). N o correction has been made for the slight dilution caused by adding the anion. Measurements were made in a Cary 1 6 s spectmphotometer ( 1 4 0 ~ ) .

436

D. C. WATTS

The bifunctional reagent 1,5-difluoro-2,4-dinitrobenzenc is reported to cross-link between the essential thiol group and a nearby tyrosine residue (134). I n light of the above evidence it seems probable that, as with the cross-linking to a second thiol (see Section II,C), this occurs after partial unfolding of the enzyme.

B. SUBSTRATE-INDUCED CONFORMATIONAL CHANGES There is now compelling evidence to believe that the transphosphorylation process involves a conformational change in creatine kinase. Further, similar changes occur when the Cr-E-MgADP complex forms and in a particularly exaggerated form when the dead-end complex is stabilized by a Class I1 anion such as C1- or NO,- (see Section V1,E). Lack of awareness of the high degree of stability of the anion-stabilized deadend complex resulted in many earlier effects being erroneously associated with other enzyme-substrate combinations. The first evidence for a conformational change came from immunological experiments in which it was found that inhibition of creatine kinase by its antibody was prevented by creatine plus MgATP but not by either substrate alone (136). It was correctly inferred that the working combination of substrates altered the conformation of the enzyme so that the antibody would not bind. Evidence has been presented (Section V1,B) that the binding of one substrate to the enzyme alters the affinity of the enzyme for the second substrate. This might result from direct interactions of the two substrates or via a structural rearrangement of the protein. If the first possibility were correct then the dissociation constant for a substrate from the nonphosphorylated dead-end complex, where interaction between substrates should be minimal, should be similar to that for the same substrate from the free enzyme. This appears to be supported by kinetic measurements (101). Thus, the dissociation constant for creatine from the Cr-EMgADP complex (8-17 mM) was found to be about the same as that from the Cr-E complex (16-19 mM), while for MgADP the values were 0.12 mM and 0.14-0.17 mM, respectively (101). On the other hand, measurements of the dissociation of creatine from the Cr-E and Cr-E-MgADP complexes by means of the protection afforded against inhibition by iodoacetate gave values of 50 and 10 mM, respectively, 134. H. L. Aanning and T.A. Mnhonald, Fed. Proc., Fed. Amer. Soc. 29, 854 (1970). 135. A. J. Samuels, Biophys. b. 1, 437 (1961).

Eq.Bkl.

12.

CREATINE KINASE

437

from which a conformationally mediated effect of one substrate on the binding of the second substrate might be inferred (136). However, in both sets of experiments, chloride-containing buffers were used so that stabilization of the dead-end complex would be expected ; hence, direct interaction between the nonphosphorylated substrates cannot be excluded. The apparent lack of interaction suggested by the conventional kinetic data (101) must result from m y difference falling within thc experimental error of this method. Temperature-jump studies indicated that binding of ATP, ADP, or the metal-nucleotide complex to the enzyme produced a conformational change but that the binding of creatine alone did not (68). Recent EPR measurements, made with an awareness of the anion effect, support and extend this finding. The binding of creatine alone to the enzyme cannot be measured by the EPR method, but addition of creatine to the E-MnADP complex produced a change in the spectral line shape indicating that either an increase in the asymmetry of the Mn2+ ion environment had occurred (i.e., a direct substrate influence) or that creatine induces a conformational change which distorts the environment of the Mn2+ion (94).The latter interpretation is favored by the finding that the effect is lost if the essential thiol groups are first blocked with iodoacetate although both substrates still bind to the enzyme (94).The binding of MnADP to the free enzyme also produces changes in the E P R spectrum but these are not affected by first alkylating the enzyme, compatible with the finding that the metal-nucleotide complex does not protect the enzyme against iodoacetamide (Section VI1,D). These changes could be explained solely on the basis of immobilization of MnADP upon binding to the enzyme (94), but taken in conjunction with other data suggest that the correct interpretation involves a contribution from conformational changes. In summary, the present picture is that creatine produces a conformational change upon binding to the enzyme-metal nucleotide complex but not on binding to the free enzyme, while the binding of the metal-nucleotide complex produces a conformational change that differs according to whether creatine is present or not. The EPR measurements showed that when both creatine and MnADP were bound to the enzyme a conformational change occurred ( 9 4 ) .AS with the free metal-nucleotide complex, this substrate combination does not protect the enzyme against inhibition by the uncharged alkylating agent, iodoacetsmide, until a Class I1 anion is added (65, Section V1,E). 136. W. J. O’Sullivan, H. Diefenbach, and M.Cohn, Biochemistry 5, 2666 (1966).

438

D. C. WATTS

Electron paramagnetic resonance measurements then indicate further marked structural rearrangements in the enzyme ( 9 4 ) .Earlier magnetic resonance measurements, assumed to be on the Cr-EMnADP complex (116, 137) were, in fact, made in the presence of N-ethylmorpholine HCl buffer so that these data should be reinterpreted in terms of the anion-stabilized dead-end complex. The same substrate combination plus either C1- or NO3- ions was also the effective agent causing the conformational change inferred to reduce the susceptibility of creatine kinase to tryptic digestion (138) and inactivation by p-nitrophenyl acetate (61,126) and iodoacetate (137,139). Addition of an equilibrium mixture of substrates to creatine kinase causes changes in the EPR spectra that cannot be attributed to the dead-end complex either in the presence or absence of anion and so were tentatively ascribed to conformational effects in the Cr-E-MnATP complex (94).This agrees with the finding that in the presence of Mg” only an equilibrium mixture of substrates affords significant protection to the enzyme against inhibition by iodoacetamide (65). However, in these experiments, perturbing the equilibrium with phosphocreatine, which was present only in low concentration in the original equilibrium mixture, markedly increased the degree of protection, suggesting that the PCr-E-MgADP complex also contributes to the conformational effects observed. I n spite of the profound effects of the conformational changes in creatine kinase upon the binding of substrates, which may bring about almost complete protection against antibody binding or inhibition by alkylating agents, the actual magnitude of the conformational change appears to be quite small. Thus, a small increase in the rate of deuterium exchange occurred upon binding MgADP to the enzyme, but this was not affected by the further addition of creatine even though the reaction was carried out in tris-HC1 buffer (68). Lack of evidence for a conformational change of any magnitude was also obtained from measurements of sedimentation constants and reduced viscosity (58).Similarly, although the binding of MgADP or MgATP produced an extrinsic Cotton effect, no changes in optical rotation occurred that could be related to a conformational change in the protein, and the situation was not altered by the simultaneous addition of MgADP and creatine (I@). 137. W. J. O’Sullivan and M. Colin, JBC 243, 2737 (1968). 138. G . Jacobs and L. W. Cunningham, Biochernkstry 7, 143 (1968). 139. D. C. Watts and B. R. Rabin, BJ SS, 507 (1962). 140. J. H. R. Kiigi and T. K. Li, Fed. Prac., Fed. Amer. Sac. Em. B b l . 24, 285 ( 1965).

12. CREATINE

KINASE

439

C. FORMATION AND TOPOGRAPHY OF THE CATALYTIC SITE As has already been discussed (Section IV,A), the use of a series of creatine analogs gives a picture of the creatine binding site as a narrow slot in the surface of the enzyme into which the creatine molecule binds, with a specific binding site for the N-methyl group which helps orientate the planar guanidino group so that it is precisely aligned to receive the incoming phosphoryl group in the trans position. For the nucleotide binding site, Table VII indicates that the organization has to be such that not only is the nucleotide firmly bound but also an appropriate conformational change is induced to refine the organization of the catalytic site, perhaps to aid in the orientation effects mentioned above. The decreased ability of nucleotides other than ADP and ATP to act as substrates appears to stem more from their inability to induce the appropriate conformational change than from any inability simply to bind to the enzyme. For example, the pronounced difference spectrum produced by the Cr-E-MgADP-anion complex (Fig. 8) (l4Oa) almost disappears when ADP is replaced by IDP. The importance of the 6-amino group of adenine has often been stressed but the precise alignment required of the tri- or diphosphate chain, a corollary implicit in the precise alignment of the guanidino group, indicates the need for complete integrity of the ribose and adenine parts of the molecule (cf. Tables VII and VIII) . The point is further illustrated by the finding that the same order of effectiveness occurs for a series of metal nucleoside diphosphates with respect to the maximum velocity measured in the presence of Mg" ion and the rate of inhibition of catalytic activity by iodoacetate (i.e., reactivity of the essential thiol group) also measured in the presence of the Mn2+ion (Fig. 9). Alteration to either the sugar or the base component of the nucleotide causes a sharp and similar fall in all three parameters. The role of the metal ion is seen in aiding the steric arrangement of the phosphate chain and, by withdrawing some of the negative charge, thereby rendering the transferable phosphoryl group more susceptible to nucleophilic attack (139).Although thc metal ion does not form a bridge between nucleotide and enzyme (69,137, lQl),the environment of the metal ion appears to be severely restricted since proton relaxation rate measurements on the Cr-E-MnADP-N03- complex show that in the first coordination sphere of the Mn2+ion the number of water ligands 140:~D. C. Watts and E. J. Milner-White, unpublished data (1972). 141. M. Cohn and J. S. Leigh, Jr., Nature (London) 193, 1037 (1962).

440

D. C. WATTS Reaction velocity

Enhancement of PRR

ABCDE

ABCDE

Increase in -SH reactivity

roo50 -

A

CDE

FIG. 9. Comparison of the effect of various nucleoside diphosphates on the relative maximum velocity of creatine kinase, the enhancement of proton relaxation rate, and the increase in rate of inhibition by iodoacetate (increase in -SH reactivity). A, MeADP; B, Me3'dADP; C, Me2'dADP; D, MeIDP; and E. MeGDP. Maximum velocity measurements were made using Mg'+ ions, P R R enhancement using Mn'+ ions, and the increase in S H reactivity using Mg2' ions except for IDP which was made using MnZt ions. Data from reference 84.

is less than one-half (142). Changes in the water proton relaxation rate as a function of frequency and temperature point to the involvement of only outer sphere relaxation in this complex, and a conformational change was inferred to render the Mn2+ ion inaccessible to the solvent water. Thus, as with the guanidino group, a picture emerges of a conformational change squeezing the roughly aligned metal-nucleotide complex into the precise orientation required for transphosphorylation t o occur. Attaching a nitroxide spin label [iV (1-oxyl-2,2,5,5-tetrainethyl-3-pyrrolidinyl) iodoacetamide] to the essential cysteine residue in each subunit of creatine kinase results in loss of catalytic activity but retention of the ability to bind the metal-nucleotide substrate (89). The resonance changes that accompany the subsequent binding of creatine are also lost as with the alkylation by iodoacetamide (137) and, by inference, the accompanying conformational changes. However, as discussed in Section VII,B, the actual movements of the polypeptide chains of creatine kinase appear to be very small so that the overall topography of the catalytic site may reasonably be assumed to have changed very little. This is very fortunate since the finding (143) that the Mn2+ion in the spin-labeled E-Mn nucleotide complex is sufficiently close to the spin label covalently attacked to the enzyme to give an easily measurable dipole-dipole interaction reflected in the spin label EPR spectrum makes 142. G. H. Reed, H. Diefenbach, and M. Colin, JBC 247, 3066 (1972). 143. If. Colin, H. Diefenbach, and J. S. Taylor, JBC 246, 6037 (1971).

12. CREATINE

441

KINASE

TABLE XVZII BICTWISICN THP; sUuSTlL%TES BOUND TO CIllCATINl
1)ISTANCES

Enzyme system

+

Creatine kinase ADP (rabbit muscle) Creatine kinase (spin labeled) (chicken heart, BB)

b

Paramagnetic center

Ligand

Mn2+

Creatine

N.4

(Distance) IL

Mn-CH2 9.8 Mn-CHa 10.3 MnATP N-Mn 11.5 & 0 . 6 MnADP N-Mn 7.5 & 1.5 MgADP N.-H, 9.3 N*-Hs 8.3 Nm-Hi, 9.7 ADP N--H2 7.9 N-Hs 7.8 N--Hlt 9.9 Creatine N-CH, 10.6 N--CHa 10.5

As determined by NMR measurements (143, 144). For location of the adenosine protons see Fig. 10.

it possible to estimate the distance between the two paramagnetic species. The value obtained (Table XVIII), taken in conjunction with other data (89, 14S), show that the nitroxide radical is more immobilized and closer to the metal ion in the ADP complex than it is in the ATP complex. Other resonance measurements (144) make it possible to calculate the distance between the spin label and selected protons of creatine and the nucleotide. The distance between the creatine protons and the metal ion can only be measured in the dead-end complex, Cr-E-MnADP (Table XVIII). These values, also listed in Table XVIII, indicate that the presence of the metal ion causes the adenine to move away from the spin label while the relative position of the ribose is not very much altered. But perhaps the most surprising finding (cited in reference 94) is that, contrary to what has been believed for the last ten years or so, the metal ion does not bind to the y-phosphoryl group of ATP but more probably forms a ligand between the p- and a-phosphoryl groups. A schematic representation of the Cr-EMnADP-N03- complex based on these measurements is shown in Fig. 10. Because the unpaired electron of the spin label is some 7 A from the thiol group to which it is covalently bound, precise location of the thiol is not possible, The measurements suggest that it is roughly equidistant from the adenosine and creatine 144. A. S. Mildvun and M. Cohn, Advan. Enzymol. 33, 1 (1970).

442

D. C. WATTS

ADP

Nitrate

Creatine

FIU.10. Perspective drawing of the three-dimensional organization of the Cr-EMnADP-nitrate complex based on the measurements given in Table XVIII (94).

protons but nearer to creatine than the nucleotide-bound metal ion. Whether it is sufficiently close to interact with the guanidino group and participate in the catalytic mechanism as has been suggested (139) must remain conjectural for the moment.

D. THE“ESSENTIAL” THIOL GROUP 1. Structural Involvement The role of the single reactive cysteine group per subunit, now shown to be part of the catalytic site (see previous section), has commanded much attention. It possesses the unusual feature that the thiol group exists in a partially ionized state that does not change significantly over the whole range of p H in which the enzyme is structurally stable (139,and see Fig. 2). Comparison of the rate of alkylation above neutral pH with that of a freely ionizing thiol suggests that the nucleophilic power, and hence degree of ionization, is not great. In contrast, blocking the essential thiol with iodoacetate does not affect the electrophoretic mobility of creatine kinase on starch gels a t p H 8.4, while blocking with iodoacetamide reduces the electrophoretic mobility by about 1 unit of charge per subunit and, indeed, the half-alkylated enzyme may

12. CREATINE KINASE

443

readily be detected by this means (16, 1.455).Further evidence suggesting a high degree of ionization of the essential thiol has come from determinations of the effect on enzyme stability of blocking the essential thiol with a neutral or a negatively charged reagent using differential thermal analysis measurements (146). Under conditions where the free enzyme is denatured a t 61.7", blocking with iodoacetate reduces the denaturation temperature to 58.5", but blocking with iodoacetamide reduces it much further to 53.5". This finding, suggesting a role for the essential thiol in the stabilization of enzyme tertiary structure, has been confirmed by quantitative measurements using differential scanning calorimetry (147). Reaction of the thiol with N-ethylmaleimide (147) appears to be rapidly followed by opening of the maleimide ring to give the N-ethylsuccinamate derivative since this inhibitor behaves exactly like iodoacetate in causing no change in electrophoretic mobility and only reducing the thermal stability by 3-4". Thus, it would seem that the negative charge on the essential thiol plays an important role in stabilizing the enzyme structure and the bulkiness of a particular alkylating agent is not very important. This is further emphasized by the fact that blocking the thiol with 2,4-fluorodinitrobenzene (FDNB) has a highly destabilizing effect and reduces the enzyme denaturation temperature by about 12" (147). Another feature of the FDNB reaction is its extreme rapidity which is too fast to measure even when the enzyme concentration exceeds that of the reagent (64),suggesting that there is a specific, perhaps hydrophobic, area on the enzyme close to the essential thiol group with which the dinitrobenzene part of the inhibitor undergoes additional interactions. The pH-independent reactivity of the thiol has been explained by specific hydrogen bonding to a nearby histidine (129, and see Fig. 11). Location of the thiol in a hydrophobic area might aid this process since a hydrogen bond involving a thiol group would be expected to be very weak. 2. Importance for Catalytic Activity In a proposed mechanism for the catalytic reaction (139),a role was suggested for the essential thiol in which it, withdrew a proton from the creatine guanidino group and initiated a circular flow of electrons from the guanidino group via the 7 - and p-phosphoryl groups of ATP back to the thiol; thus, transfer of the phosphoryl group from ATP to creatine was greatly facilitated (Fig. 11A). This mechanism was based on the 145. D.C. Watts, Abstr. 1st Meet. Fed. Eur. Biochem. SOC.A13 (1964). 146. D.C. Watts, BJ 100, 13C (1966). 147. B. Moreland, personal communication (1972).

444

D. C. WATTS

I

h-

t

(B)

FIG.11. Representations of the mechanism of action of creatine kinase inrorporating recent findings. Only the initial stage of the trmsphosphorylation process is shown in each case since the reaction pathways have not been altered. (A) Modifid from reference 139; (B) modified from reference 144. Thc relative merits of the two schemes are discussed in the text.

observation that an equilibrium mixture of substrates (now known to be the Cr-E-MgADP-NOa- complex) protected the essential thiol against alkylation by iodoacetamide but single substrates did not. It was appreciated that a conformational change in the enzyme was also likely to be involved in this process. Such is the involvement of the thiol in this mechanism that its chemical modification might be expected to destroy all catalytic activity. I n practice, to demonstrate unambiguously either this or the converse, t.hat the chemically modified enzyme retains some activity, has proved remarkably difficult. The reason for this

12. CREATINE

KINASE

4.45

appears to be that alkylation of the thiol groups on one subunit of the enzyme induces a conformational change that is transmitted to the second subunit; thus, its essential thiol is no longer reactive although the remaining catalytic activity is not lost. Evidence for this curious behavior first emerged from an attempt to titrate to completion the essential thiols of the rabbit brain and chicken muscle and brain creatine kinases. A slight departure from linearity was seen with the muscle enzyme and became very marked with the brain enzyme (32). This finding has since been amply confirmed (8, 15, SS, 148) for the enzymes from chicken and ox brain and dogfish and primate muscle. For the ox brain enzyme (148), the level of iodoacetamide-insensitive residual catalytic activity was found to depend on the absolute enzyme concentration and the ratio of inhibitor (lo) to enzyme (E,) . With 3 @ enzyme complete inhibition could not be achieved even when Io/E0 = 100 a t either 0" or 30". When the enzyme concentration was raised to 100 $If, essentially complete inhibition was achieved a t 30" with Io/Eo= 2. Thus, there would appear to be an interaction between the protein molecules a t the higher concentration (100 I/JM = 0.8% w/v) that effectively prevents the protective conformational change from taking place. If the partially inactivated enzyme was diluted 400-fold in dithiothreitol it was found to have become markedly stabilized to storage at 3" as compared with the native enzyme. Stability was maximal when the enzyme was half-inactivated (148) as might be expected if there were interaction between subunits. This phenomenon of subunit interdependence is not obvious with the rabbit muscle enzyme, and the reaction is apparently first order in enzyme and inhibitor over a wide range of concentrations yielding an apparent second-order rate constant with iodoacetate of about 300 M-l min-' a t 30", varying with the actual pH and ionic strength used (64, 12%). Complete inhibition can be achieved a t 0" with 62 p M enzyme concentration and I,/E, = 100 (64). However, the reaction is slow, taking about 80 min and, as the authors pointed out, is quite unsuitable for a titration type of experiment where the inhibitor is used a t a series of concentrations less than that of the enzyme Concentration. Here it is the extent of the reaction that is important, and from the secondorder relationship completion of the reaction would be expected to take a long time. Increasing the enzyme concentration tenfold makes possible a corresponding increase in the range of inhibitor concentrations used, and the reaction time may be decreased by a factor of a hundred. Hence, for rapid and complete reaction high concentrations of enzyme and 148.

R. S. Atherton. J. F.Laws, and A. R. Thomson, BJ 118, 903 (1970).

446

D. C. WATTS

inhibitor are recommended (6‘4). With prolonged reaction times (2 hr or more) a t low inhibitor concentrations errors in stoichiometry may creep in from such effects as spontaneous oxidation of the enzyme and partial denaturation of the native or alkylated enzyme exposing more groups for reaction with the inhibitor. These are normal problems. Abnormal complications may arise because some enzyme preparations (although apparently not that used in reference 64) show a residual activity after alkylation of about 10% of the initial level. This has been ascribed to a partial oxidation of a fraction of the enzyme that makes it unavailable to the inhibitor but able to be reactivated by the addition of a thiol used to stop the alkylation reaction (120). Even apparently fully akylated enzyme has been found to contain a residual activity of a few percent of the initial value, and this is not destroyed until the enzyme is completely denatured (149)as indicated by reaction of the buried thiol groups. The two “essential” thiol groups may also be oxidized to what is inferred to be the sulfenic acid (-SOH) by treatment with four equivalents of either iodine (149~) or dansyl chloride (149b) per mole of enzyme, the reaction being reversed by treatment with thiols such as dithiothreitol. Again, although complete loss of activity was claimed, the data show that this was not actually achieved. Thus, rabbit muscle creatine kinase could show to a small extent the phenomenon of subunit interdependence so obvious with the brain-type .enzymes. This could be explained if the relaxation time of the protein transitions is slow, of the order of many minutes. The reaction with iodoacetate when carried out over 3 hr has been reported to deviate from the linear pseudo-firstorder relationship when the reaction has gone to more than 70% (64) and, although not a very reliable region of the curve, it might be taken to indicate the occurrence of such a secondary event. With the BB creatine kinase of chicken heart, reaction a t 25°C of an 0.7% enzyme solution with Io/Eo = 100 resulted in rapid reaction with a variety of alkylating agents but after this had gone to completion 2 5 3 0 % of the initial catalytic activity remained (15).The K , values for creatine and MgATP with this remaining activity of the akylated 149. L. Noda, T. Nihei, and E. Moore, Proc. Znt. Congr. Biochem., 6th, 1961 p. 118 (1963).

149a. D. Trundle and L. W. Cunningham, Biochemistry 8, 1919 (1969). 149b. C. S. Brown and L. W. Cunningham, Biochemistry 9, 3878 (1970). Thc reaction using dansyl chloride reported here should be contrasted with that described in reference Id7 where much higher concentrations were used to label fluorescently two Iysine residues. With the low concentration of reagent, no fluorescent label is incorporated into the protein and, while the sulfenic acid obtained at pH 6.1 appears relatively stable, at pH 8.5 to 9.0 oxidation proceeds further to the sulfinic and sulfonic acids which are not readily reduced by thiols.

12. CREATINE

KINASE

447

enzyme were the same as those found with the native enzyme. An attempt was made to determine whether or not full alkylation of both essential thiols in each molecule had occurred. As with the rabbit enzyme, destruction of the residual activity required complete unfolding of the enzyme, but in this writers’ opinion the results remained unfortunately ambiguous because, despite the author’s conviction that both essential thiols per molecule had been fully alkylated, this has not been adequately demonstrated. A study of the thiol groups of the creatine kinases from ox muscle and brain failed to provide any support for the hypothesis that the fully alkylated enzyme may retain activity (150).After reaction of 1.95 thiol groups per molecule of muscle enzyme with 5,5’-dithiobis(2-nitrobenzoic acid), less than 0.5% of the original activity remained. Using similar conditions ( l o / E o= 500), the ox brain enzyme was 99.4% inactivated with the reaction of 2.1 thiol groups per molecule. The ox brain enzyme was found to be particularly susceptible to oxidation even a t 0” and a steady decrease in the number of essential thiol groups from 2.1 to 1.4 over 5 days was accompanied by a similar decrease in catalytic activity (150).Over this period the reaction with DTNB became progressively more complex and multiphasic, suggesting a slow unwinding or rearrangement of the protein structure. This, in turn, may result in disulfide bond formation and intra- and interchain disulfide bond interchange reactions. Under these conditions, attempts to correlate the stoichiometry of the reactive thiol groups with the catalytic activity becomes a meaningless exercise since it is no longer possible to distinguish the original essential thiol groups from those which were originally buried and have become exposed as a result of conformational rearrangements (150).Similarly, with the primate muscle enzymes, which are also relatively unstable, the number of thiol groups that react rapidly with DTNB (and might be inferred to be the two essential thiol groups) remain at approximately two per molecule in preparations where the enzyme had lost a significant measure of activity (151).This would be explained if the oxidation of the essential thiol group produced a conformational change that exposed a buried thiol group, the level of “essential” thiol groups thus being kept constant ( 1 5 1 ~ )However, . this phenomenon, although it may explain some 150. K. Okabe, H. K. Jacobs, and S. A. Kubg, JBC 245, 6498 (1970). 151. I. Kumudavalli end D. C. Watts, unpublished observations (1971). 151a. This effect might also explain the unusual behavior of the “essential” thiol groups of primate rreatine kinrtse in showing an apparently normal ionization curve in the presence of Cr + MgADP as reported in reference 8. At least with some preparations of monkey (Mncnca mulutta) creatine kinase, the essential thiols behave exactly like those of the rabbit enzyme in showing no significant change in reactivity toward iodoacetamide over the pH range 7-10 (Chegwidden and Watts, unpublished data, 1972).

448

D. C. WATTS

of the previously reported findings concerning the reactivity of the essential thiol groups, is quite separate from that in which after maximum reaction of the available thiol groups the enzyme retains activity which is resistant to the inhibitor. If modification of the essential thiol is accompanied by activity loss, as is firmly concluded from the ox brain experiments (160),then the residual activity must be associated with essential thiol groups that have in some way become protected against the alkylating agent. For an enzyme with initially only one reactive thiol group on each subunit this can only reasonably result from the reaction of one thiol group causing a conformational change that is transmitted to the second subunit and, in some way, protects the second essential thiol group, thereby preventing further loss of activity. In summary, it remains to be demonstrated that creatine kinase from any species may retain activity after the two essential thiol groups per molecule have been totally blocked. However, its manifest involvement in the conformational changes that accompany transphosphorylation render this possibility unlikely whether or not the essential thiol group reacts with the creatine guanidino group in the catalytic reaction. Finally, mention must be made of the fish creatine kinases which have more than two rapidly reacting thiol groups per molecule. With the dogfish enzyme there are four, but catalytic activity extrapolated to zero with reaction of the first two (33). With a low enzyme concentration (0.014%),the rate of activity loss with iodoacetamide progressively deviated from a pseudo-first-order reaction and complete inhibition was not achieved. This may explain why the carp enzyme appeared to have only three reactive thiol groups when titrated with DTNB ( 5 ) . Unless the subunits are dissimilar, the equivalent of one thiol group per molecule must have become protected. The total number of thiol groups per molecule, six in the dogfish and eight in the carp enzyme (Table I ) , suggests that these have the same distribution in the molecule as the thiols in other kinases and that two of them must lie very close to the enzyme surface, perhaps becoming exposed as a result of the investigations being carried out well above the normal physiological temperature for these animals.

3. Effects of Substrates Bearing in mind the concentration-dependent behavior of creatine kinase toward alkylating agents, the importance of closely controlling pH and ionic strength in the reaction with iodoacetate (139),and the more recently revealed adventitious effects of anions (65) it is perhaps not surprising that investigations of the effects of substrates on the in-

12.

CREATINE KINASE

449

hibition of enzymic activity by thiol reagents yielded somewhat different results in the hands of different workers. The qualitative picture that emerges is that, with iodoacetamide, single substrates in the presence or absence of Afg2+ ions are essentially without effect on the rate of alkylation. Protection is observed only with an equilibrium mixture of substrates or with creatine plus MeADP in the presence of a Class I1 anion (58, 65, 136, 139). With iodoacetate the situation is more complex. ATP and MgATP both protect (64, 84, 139), but while ADP protects MgADP enhances the rate of inactivation (83, 136, 139). Creatine and phosphocreatine protect to a small extent (64, 136). The dead-end complex protects strongly in the presence of a Class I1 anion and, although the effect of a n equilibrium mixture of substrates is not known, protection would be expected on the basis of the behavior of single substrates. From a study of the effects of a series of nucleotides on the enzyme velocity, protein relaxation rate, and reactivity of the thiol toward iodoacetate (84, see also Fig. 9) it was concluded that binding different nucleotides affected the thiol reactivity to different extents via a conformational change induced in the protein. I n accord with the kinetic data for creatine kinase (Section VI,B), showing an interaction between substrate binding sites, the inference of a conformational change upon the binding of a nucleotide appears correct. It cannot, however, influence the activity of the essential thiol group since the rate of reaction with iodoacetamide remains unaltered. Explanation of the change in rate with iodoacetate must be sought in terms of a variable degree of electrostatic interaction between the negatively charged carboxyl group of iodoacetate and other charged groups in the enzyme-substrate complex appropriately located near the essential thiol. A simple solution in keeping with the observations (139) and made more reasonable by the subsequent findings that the essential thiol is close to the nucleotide binding site (Table XVIII) and that the metal ion does not bind to the y-phosphoryl of A T P (94) woud be for the negative charge of A D P to hinder and the positive charge of the metal ion in the MgADP complex t o aid the attack of the inhibitor. MgATP would then protect, again because of the additional negative charge contributed by the y-phosphoryl group. This explanation is readily compatible with a conformational change that more precisely aligns the phosphate chain of the nucleotide, the extent of this process depending on the particular nucleotide used (see also Sections VI1,B and C ) . The rate of reaction with iodoacetate could also be influenced by electrostatic interaction with positively charged groups located on the enzyme. The existence has been inferrcd (139) of groups that affect the reaction of iodoacctate with the essential thiol groups from the fact

450

D. C. WATTS

that the rate of inhibition, unlike that with iodoacetamide, varies with the p H and anionic strength (152).Such groups could also be affected by conformational changes in the enzyme. It is of considerable interest that the essential thiol group can be so close to the nucleotide binding site and yet remain uninfluenced by nucleotide substrate-induced conformational changes, particularly when the conformational change produced by adding a metal ion to the enzyme-nucleotide complex is found to increase the immobilization of a spin label covalently attached to the thiol group (89).This suggests that the essential thiol group is located in a part of the enzyme molecule that is structurally isolated from that part which contains the nucleotide binding site. Evidence in support of this idea comes from differential scanning calorimetry measurements that show that the essential thiol group and the nucleotide binding site separately affect enzyme stability. Thus, blocking the essential thiol group with iodoacetamide lowers the thermal denaturation temperature by about 8" while MgATP and MgADP raise the thermal denaturation temperature by 3" and 5 " , respectively, and this measure of protection by nucleotides is achieved whether the thiol group has been blocked or not (147'). Although the thiol group appears to be isolated from the nucleotide binding site, it must nevertheless be close to it since the attachment of pyrrolidinyl iodoacetamide lengthens the cysteine side chain by only about 7 A and yet brings the nitroxide radical to within a few angstroms of the nucleotide phosphate chain (Table XVIII). This isolation of the thiol group may explain the lack of correlation between the enzyme activity with Mg2+,Ca2+,Sr'+, and Ba2+ions and the EPR and PRR spectra of the spin-labeled enzyme-metal nucleotide complexes (89). For each series of ADP- and ATP-metal complexes, the range of catalytic activity varied by a factor of 100 (with the Ba2+ion being virtually inactive), but the resonance measurements gave similar values. The authors (89) concluded that the enzyme-metal nucleotide complexes all had the same conformation but that the conformations upon adding creatine to form the dead-end complex were different. Likewise, the reactivity of the thiol group toward iodoacetamide was the same with either MgADP or BaADP but upon adding creatine the magnesium complex gave protection (Clions presumably being present) but the barium complex did not. Here 152. A comparable situation exists with the two catalytic site histidine residues of ribonuclease where the positive charge on one histidine orientates bromoacetate in a favorable manner for attack on the second histidine. U'ith this enzyme, unlike creatine kinase, bromoacetamide reacts less rapidly indicating the importanc*e of n charge-charge interaction between enzyme and reagent.

12.

CREATINE KINASE

45 1

again is evidence that, with regard to the reactivity of the thiol group, conformations associated with the catalytic process affect it but those associated only with nucleotide binding do not. It would have been interesting to compare the reactivity of the thiol with iodoacetate in‘ these combinations to see if any difference in electrostatic interaction could be detected to correlate with the bulkier Ba2+ion.

E. MECHANISM OF TRANSPHOSPHORYLATION It must be stated at the outset that no particular amino acid side chain has been positively identified as making contact with either substrate a t any stage in the transphosphorylation process. Hence, any [‘paper” mechanism must be deemed highly speculative. Nevertheless, it remains true that a great deal is known about the catalytic process which it may be useful a t this stage to summarize. 1. Each subunit contains one catalytic site (Sections II,VI, and VII). 2. Each catalytic site contains separate substrate binding sites (Sections VI,A and B). 3. Both substrates bind simultaneously to the enzyme (Section V1,B)’. 4. The presence of one substrate on the enzyme enhances the binding of the second substrate but the substrate binding sites are essentially preformed, and creatine kinase does not belong to the “induced fit” group of enzymes (Section V1,B). 5 . The metal-nucleotide complex is the true substrate of the enzyme (Section V1,A). 6. When the nucleotide substrate binds to the enzyme two sorts of conformational change occur. The first accompanies binding of the nucleotide to the free enzyme and probably increases the receptivity of the creatine binding site. The second occurs upon the subsequent binding of creatine and is probably the initiation of the catalytic process (Sections VI1,B and C). The time taken for the conformational changes is less than the turnover time of the enzyme (68). 7. Enhancement of the binding of the guanidino substrate by the nucleotide requires the 6-amino group of adenine. The magnitude of the effect depends on the nature of the activating metal ion (Sections VI1,B and D, Fig. 8, Table 1x1. 8. It has not been positively demonstrated that the binding of creatine to the free enzyme produces a conformational change (although this occurs with arginine and arginine kinase). It is only known that the presence of the guanidino substrate on the enzyme enhances the binding of the nucleotide substrate (see point 4 above, Sections VI,B and VI1,B).

452

D. C. WATTS

9. Creatine is bound in the creatine binding site with the planar guanidino group orientated in a highly specific manner. The creatine methyl group plays an important role in this process. It also appears to be important in promoting the conformational changes associated with the catalytic process (the topography of the creatine binding site has been quite well defined) (Section IV,A, Table V). 10. The three-dimensional orientation of the metal-nucleotide substrate in its binding site is also well defined (Table XVIII, Fig. 10). 11. The metal ion is located across the a,P-phosphoryl groups in the ADP and ATP complexes and is not hydrated in the anion-stabilized dead-end complex (Sections VI1,B and C ) . 12. The nucleotide and guanidino substrates on the catalytic site are orientated in such a way as to allow simple transfer of a phosphoryl group from one to the other. [In the absence of the transferable phosphoryl group, a planar anion such as nitrate forms a stable dead-end complex with creatine and MeADP (Section VI,E, Fig. l o ) . ] 13. I n creatine the nitrogen atom of the guanidino group that is trans to the methyl group acts as the phosphoryl acceptor (Section IV,A). 14. Transphosphorylation involves the conversion of the site that readily binds a tetrahedral anion into one that selectively binds planar anion or, because of their size F-, C1-, and Br- (but not I-) (Sections VI,E and VI1,B). 15. Conformational changes allow (or cause) the nucleotide and guanidine to approach each other. (This is not proved, but the evidence of Table XVIII is highly suggestive.) 16. If the conformational change cannot occur then neither can transphosphorylation (Sections VI1,B and D) . 17. The reactivity of the essential thiol groups is closely linked to the conformational changes that occur when both substrates are bound to the enzyme. Blocking the thiol groups prevents these conformational changes and kinase activity is either abolished or substantially diminished (Section VI1,D). 18. The integrity of a histidine and a lysine side chain are also essential for catalytic activity. Modification of the histidine causes loss of catalytic activity without any gross impairment of substrate binding or the associated conformational changes (Section VII,A,3). Modification of the lysine causes impairment of the conformational changes that accompany formation of the E-MgADP complex and, in arginine kinase, loss of ability to produce a difference spectrum (Section VII,A,2). Thus, a detailed picture emerges of the way in which the enzyme first binds the substrates; this, in turn, initiating a series of conformational changes, causes the substrates to become more tightly bound and more

12. CREATINE KINASE

453

precisely aligned. In association with these changes the tetrahedral binding site of the transferable phosphoryl group alters so that the phosphoryl group is in some way induced to form the planar sp3d hybridized state, a n intermediate in the reaction, thereby facilitating transfer to the receptor substrate. The nucleophilic attack of the receptor substrate may be further aided by the two substrates being drawn closer together. The disposition of the substrates relative to each other (Fig. 10) and the way in which planar anions stabilize the Cr-E-MeADP complex suggests that phosphoryl transfer occurs by a simple “in-line” mechanism. In the absence of anions to stabilize the dead-end complex, the protection of the essential thiol group against alkylation by iodoacetamide that may be attributed to a working combination of substrates may be as much as 40-500/0 (65). This suggests that, when the enzyme is actively catalyzing the transphosphorylation reaction, as much as 50% of the total reaction time may be spent with the transferable phosphoryl group in the planar state. This is in accord with the rapid-equilibrium random kinetic mechanism (Section VI,B) and is supported by resonance measurements (94). Since in a reaction pathway the transition state of the reaction is considered to be associated with an energy maximum of short duration, it seems probable that the observed effects are associated with relatively long-lived intermediates in which the stabilized planar phosphoryl group is more closely linked with one or other of the substrates. We come now to the specific roles of the individual amino acids listed in Section VI1,A. It is clear that the catalytic reaction would be aided by removal of a proton from the creatine guanidino group and addition of a proton to ADP in the forward reaction and vice versa for the back reaction. These roles were suggested t o be fulfilled by a hydrogen-bonded thiol-imidazole and base pair (Fig. 11A). Subsequent investigations have indicated a clear involvement of the essential thiol group with the conformational changes that occur during catalysis (Sections VI1,B and D) but have failed to show any interaction between thiol and guanidine. On the other hand, resonance measurements of the position of a spin label on the thiol group (Table XVIII) indicates that the thiol group could be near enough to the guanidino group to interact with it; furthermore, an essential histidine group has been discovered with no known role and with little, if any, involvement in conformational changes (198). Its close association with the catalytic site is indicated by the finding that MgATP and MgADP partially protect (20-25%) against inhibition by diethylpyrocarbonate (but see reference 150). Creatine does not protect although arginine protects arginine kinase under these conditions

454

D.

C. WATTS

(128).However, this could reflect the fact that binding arginine to arginine kinase causes a conformational change while binding creatine to creatine kinase apparently does not (see Section VI1,B). Thus, it would seem that a histidine side chain is available in the catalytic side that could fulfill the function indicated in Fig. 11A. For such a histidine it might be expected that the dead-end complex or a working substrate combination would provide even greater protection against the inhibition. I n fact, the protection found was no greater than with the magnesiumnucleotide complex alone. However, the experiment was carried out in 50 mM phosphate buffer which is quite inhibitory (Table VI) and could have affected the results, although the corresponding combination with arginine kinase did give increased protection. Until more is known about the properties of the essential histidine residue this problem must remain open. An alternative mechanism (Fig. 11B) adopts the idea of a charge transfer complex well authenticated for the serine proteases (163,164) and indicated for lactate dehydrogenase from preliminary X-ray data (166) as possible roles for the essential thiol and histidine residues. This very reasonable suggestion runs into the same problems as that in Fig. 11A. The properties of the cysteine and histidine are not readily compatible with a charge transfer complex that interacts with one of the substrates. There is no evidence as to the nature of the side chain that interacts with the creatine carboxyl group although such an interaction clearly contributes toward the specificity of the creatine binding site (Fig. 4, Table V). Because the specificity is so high it may well turn out that, as now seems to be the case for lysozyme and the serine proteases, interactions with the polypeptide backbone make a major contribution. Although the essential lysine residue appears to be part of the catalytic site (197,128),it cannot be directly involved in interactions with the substrates since it is not affected by single substrates and the anionstabilized dead-end complex raises the pK value of the €-amino group without affecting its reactivity. For these reasons a role in mediating the conformational changes appears to be indicated. As discussed in Section VII,A,4, a tyrosine residue appears to be closely associated with the binding of both substrates. In arginine kinase, nitration of the tyrosine residue abolishes interaction with the substrates, 153. D. M. Blow, J. J. Birktoft, and B. S. Hartley, Nature (London) 221, 337 (1969). 154. C. S. Wright, R. A. Alden, and J. A. Kraut, Nature (London) 221, 235 (1969). 155. M. G. Rossman, M. J. Adame, M. Buehner, G . C. Ford, M. L. Hackert,

P. J. Lentz, Jr., A. McPheraon, Jr., R. W. Schevite, and I. E. Smiley, Cold Spring Harbor Symp. Quant. Bwl. 36, 179 (1971).

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but in creatine kinase, although the same sort of difference spectrum as is found in arginine kinase suggests involvement of a tyrosine residue, it is not available for reaction with tetranitromethane (188,129). To establish the precise role of these side chains in the catalytic mechanism remains a task for the future.

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Arginine Kinase and Other Invertebrate Guanidin0 K inases ~

J . F. MORRISON I . Introduction . . . . . . . . . A . Historical Background . . . . . B . Catalytic Reaction . . . . . . C . Discovery and Isolation . . . . . D . Distribution and Function . . . . I1. Determination of Enzymic Activity . . . . A . Chemical Stop Methods . . . . . B. Isotopic Methods . . . . . . C . Continuous Recording Methods . . . I11. Molecular Properties . . . . . . . A . Molecular Weight and Subunit Composition B . Stability . . . . . . . . . C . Amino Acid Composition . . . . . D . Immunological Reactions . . . . IV . Catalytic Properties . . . . . . . . A . Activation by Bivalent Metal Ions . . B . Substrate Specificity . . . . . . C . pH Optimum . . . . . . . D . Function of Amino Acid Residues . . V . Reaction Mechanism . . . . . . . VI . Equilibrium . . . . . . . . . .

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457 457 459 459 461 464 464 465 465 466 466 468 469 470 471 471 473 476 477 482 485

.

I Introduction

A . HISTORICAL BACKGROUND

It has been established that invertebrates contain a number of distinct N-phosphorylated guanidino compounds which are commonly referred 457

458

J. F. MORRISON

to as phosphagens. In addition, it has been shown that each phosphagen is synthesized by a specific enzyme whose existence was demonstrated subsequent to the isolation and identification of the phosphagen. Thus, it would seem appropriate to make brief reference to the discovery and structure of these compounds before proceeding to discuss the enzymes responsible for their biosynthesis. Following the recognition of phosphocreatine as the characteristic phosphagen of vertebrate muscle (1, 2 ), phosphoarginine was isolated from crayfish muscle in 1928 by Meyerhof and Lohmann (3) and shown to have the structure illustrated in (I). Between 1928 and 1963, extenOH I N-P=O / \ HN=C, OH N-(CHJ,-CH(NHJ-COOH H

H

Phosphoarginine (1)

sive studies were made on the distribution of these two phosphagens within the animal kingdom. The results indicated that, in addition to phosphocreatine and phosphoarginine (I), invertebrates contain five other phosphagens. In 1953, phosphotaurocyamine (11) was found to be H /N-P=O \ HN=C OH N-C Hi- C &- SO,H H

OH H i N-P=O / \ HN=C, OH N- CH,-COOH H

Phosphotaurocyamine

Phosphoglycocyamine

(11)

(111)

yH

pH

H N-P=O \ OH HN=C OH I H,- CH,-O-P-O-CH,-CH(NH,)-COOH H II 0 I

h-c

Phospholombricine

(rv) 1. P. Eggleton and G. P. Eggleton, BJ 21, 190 (1927). 2. C. H. Fiske and Y. Subbarow, JBC 81, 629 (1929). 3. 0. Meyerhof and K. Lohmann, Biochem. 2. 196, 22 (19%).

13.

459

INVERTEBRATE GUANIDINO KINASES

present in Arenicola marina ( 4 ), phosphoglycocyamine (111) in Nereis diversicolor ( 4 ), and phospholombricine (IV) in Lumbricus terrestris ( 5 , 6 ) .About a decade later, the last two of the seven known phosphagens were isolated from Phascolosoma vulgare (7) and Ophelia neglecta (8) and identified as phosphohypotaurocyamine (V) and phosphoopheline (VI), respectively. /OH N-P=O / \ HN=C OH N- CH2-C H,-SO$ H

H ,OH N-P=O \ 0 HN=C OH II N-CH2-CH2-O-P-O-CHs H I OH

H

/

Phosphohypotaurocyamine

Phosphoopheline

(VI)

(V)

B. CATALYTIC REACTION The enzymes involved in the formation of the above phosphagens (I-VI) can be regarded as ATP: guanidino phosphotransferases since they catalyze the reversible transfer of the terminal phosphoryl group of ATP to a guanidino compound according to the general reaction ( l ) , guanidino compound

+ ATP MI+ phosphagen + ADP + H+

where M2+ represents an essential bivalent .metal ion. They are also referred to by the trivial name of kinase preceded by the guanidino acceptor of the phosphoryl group and this terminology will be used throughout. C.

DISCOVERY AND ISOLATION

Arginine kinase was first found to be present in extracts of crab muscle by Lohmann (9) who also demonstrated its presence in octopus muscle (10). Subsequently, Szorenyi et al. ( 1 1 ) reported the isolation of the 4. N. Van Thoai, J. Roche, Y. Robin, and N. V. Thiem, BBA 11, 593 (1953). 5. N. Van Thoai, J. Roche, Y. Robin, and N. V. Thiem, C. R. SOC.Bwl. 147, 1670 (1953). 6. N. Van Thoai and Y. Robin, BBA 14, 76 (1954). 7. Y. Robin and N. Van Thoai, BBA 63, 481 (1962). 8. N. Van Thoai, F. Di Jeso, and Y. Robin, C. R . Acad. Sci. 258, 4525 (1963). 9. K. Lohmann, Biochem. 2. 282, 109 (1935). 10. K. Lohmann, Biochem. 2. 286, 28 (1936).

11. E. T. Szorhyi, P. D. Dvornikhova, and R. G. Degtar, Proc. Acad. Sci. USSR 67, 341 (1949).

460

J. F. MORRISON

enzyme in crystalline form from extracts of the freshwater crab, Potamobius astacus, while Elodi and SzorBnyi (12) described an improved procedure for obtaining crystalline arginine kinase from P . astacus and P. leptodactylus. More recently, arginine kinase has been obtained in either purified or homogeneous form from the hermit crab, Pagurus bernhardus, the blue crab, Callinectus sapidus, and the horseshoe crab, Limulus polyphemus (IS) ; from the European lobster, Homarus vulgaris (14-17) ; from the American lobster, Homarus americanus (18); from two species of Australian seawater crayfish, Jasus verreauxi (19)90) and Panulirus longipes (21) ; ‘from the sipunculid, Sipunculus nudus (92) ; from the marine polychaetous annelid, Sabella pavonina (23) ; and from the Italian honeybee, Apis mellifera ( 9 4 ) . Two forms of arginine kinase have been isolated in homogeneous state from the muscle of L. polyphemus and shown to differ in their charges and stability, but not in their kinetic properties or molecular weights (IS,26). The suggestion has been made that the isoenzymes are products of different genes (26). Indications have also been obtained for the presence of isoenzymes of arginine kinase in other invertebrate organisms (13, 27). The first report of the presence of taurocyamine kinase in Arenicola marina and glycocyamine kinase in Nereis diversicolor was made by Van Thoai (98) and later both enzymes were prepared in a homogeneous state. Taurocyamine kinase was isolated from A. marinu (29, 30) and 12. P. Elodi and E. T. Szor&yi, Acta Physiol. 9, 367 (1958). 13. S.L. Blethen and N. 0. Kaplan, Biochemistry 7,2123 (1968). 14. L.A. Pradel, R.Kassab, F. Regnouf, and N. Van Thoai, BBA 89, 255 (1964). 15. L. A. Pradel, R.Kassab, E. Der Terrossian, and N. Van Thoai, C. R. Acnd. Sci. 280, 3212 (1965). 16. R. Virden, D. C. Watts, and E. Baldwin, BJ 94, 536 (1965). 17. E. Der Terrossian, R. Kassab, L. A. Pradel, and N. Van Thoai, BBA 122, 462 (1966). 18. S. L.Blethen and N. 0. Kaplan, Biochemistry 6, 1413 (1967). 19. J. F. Morrison, D. E. Griffiths, and A. H.Ennor, BJ a,143 (1957). 20. M. L. Uhr, F. Marcus, and J. F. Morrison, JBC !241, 5428 (1966). 21. E. Smith and J. F. Morrison, JBC 244, 4224 (1969). 22. G. Lacombe, N.V. Thiem, and N. Van Thoai, Eur. J . Biochem. 9, 237 (1969). 23. Y.Robin, C. Klotz, and N. Van Thoai, Eur. J . Biochem. 21, 170 (1971). 24. C.W. Carlson, S. C . Fink, and R. W. Brosemer, ABB 144, 107 (1971). 25. S. L.Blethen, ABB 149, 244 (1972). 26. S. L. Blethen, Bwchem. Genet. 5, 275 (1971). 27. R. Virden and D. C. Watts, Comp. Biochem. Physiol. 13, 161 (1964). 28. N.Van Thoai, Bull. SOC.Chim. Biol. 39, 197 (1957). 29. R.Kassab, L. A. Pradel, and N.Van Thoai, BBA 99, 397 (1965). 30. N. Van Thoai, L. A. Pradel, and R. Kassab, “Methods in Enzymology,” Vol. 17A,p. 1002, 1970.

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GUANIDINO KINASES

461

glycocyamine kinase from Nephtys coeca (30,31). Homogeneous preparations of lombricine kinase from Lumbricus terrestr& (29, 30) and Megascolides cameroni (32) have also been obtained following earlier reports of the occurrence of the enzyme in these same organisms (3S,S4). Methods have been described for the purification of hypotaurocyamine kinase from Phascolosoma vulgare (35) and opheline kinase from Ophelia neglecta (36),but so far these enzymes have not been prepared in pure form.

D. DISTRIBUTION AND FUNCTION Since the discovery of the various guanidino kinases much interest has centered on the distribution of these enzymes within the invertebrate phyla and a wide range of organisms has been examined for their presence. It is not practical to list all those organisms which have been shown to contain one or more of these enzymes; therefore, a selected number of examples has been chosen for inclusion in Table I (9-24,2732, 35-41). However, these are sufficient to indicate the overall distribution picture for guanidino kinases and to draw attention to the diversity of their occurrence within a phylum, class, or genus. In particular, it should be noted that (a) the Annelida contain all seven of the known guanidino kinases, (b) arginine kinase is widely distributed among invertebrates and frequently occurs in association with creatine kinase, and ( c ) some genera possess only a single guanidino kinase while others contain up to three such enzymes. The Echinoidea are of special interest in that their muscles contain both arginine and creatine kinases while only arginine kinase is found in their unfertilized eggs and creatine kinase in their spermatozoa (41). 31. L. A. Pradel, R. Kassab, C. Conlay, and N. Van Thoai, BBA 154, 305 (1968). 32. T. J. Gaffney, H. Rosenberg, and A. H. Ennor, BJ 90, 170 (1964). 33. R. Pant, BJ 73, 30 (1959). 34. H. Rosenberg, R. J. Rossiter, T. Gaffney, and A. H. Ennor, BBA 37, 385 (1960). 35. N. Van Thoai, Y . Robin, and L. A. Pradel, BBA 73, 437 (1963). 36. N. Van Thoai, F. Di Jeso, Y . Robin, and E. Der Terrossian, BBA 113, 542 (1966). 37. A. H. Ennor and J. F. Morrison, Physiol. Rev. 38, 631 (1958). 38. N. Van Thoai, Y . Robin, F. Di Jeso, L. A. Pradel, and R. Kassab, Comp. Biochem. Physiol. 11, 387 (1964). 39. N. Van Thoai, N. V. Thiem, G. Lacombe, and J. Roche, BBA 122, 547 (1966). 40. B. Viala, Y. Robin, and N. Van Thoai, Comp. Biochem. Physiol. 32, 401 (1970). 41. B. Moreland, D. C. Watts, and R. Virden, Nature (London) 214, 458 (1967).

462

J. F. MORRISON

TABLE I DISTRIBUTION OF GUANIDINO KINASESAMONGINVERTEBRATE PHYLA^ ______

Phylum

Class

Species

Coelenterata Phtyhelminthes

An thozoa Twbellaria

Calliactis parasilica Polycelis cornztta

Annelida

Polychaet,a

Sabella pavonina Aphrodile amleata Hyalinoecia tubiwla Travisia forbesii Myxiwla infundibulum Glycera sp. Hermione hystrix Nereis fucata

Gephyrea Arthropod&

Crust acea

Mollusca Echinodermata

Insecta Arachnida Lamellibranchia Crinoidea Holothuroidea

Nephtys weca Nereis diversiwlor Areniwla marina Areniwla assimilis Branchiomma vesiculosum Lumbrims terreatris dlegaswlides cameroni Ophelia biwrnis Ophelia neglecta Sipunculus nudus Phascolosoma vulgare All genera All genera All genera All genera Antedon bi$da Comanthus japonica Cucumaria echinata C. elongata C. lactea Holothuria tubulosa H. jorskali Leptosynapta coplax Sticopus moebi S. tremulus S japonicus Ceramasler granularis Henricia sanguinolenta Other genera Arabacea punctulata Centrostephanus rodgersii

.

Asteroidea Echinoidea

Kinase AK, CK AK, GK, TK AK AK, CK AK, CK AK, CK AK, GK, TK CK CK CK, GK, TK CK, GK GK TK TK TK LK LK LK OK AK HTK AK AK AK AK AK AK AK AK AK

AK AK AK AK AK AK AK, CK CK, GK AK AK AK

Reference 27 27

23, 27 27,37 27 38,39 27,37 37 37 27,37 SO, 31,37 28,37 29, 30,37 97 67 29

32 40 :;I3

22, 39

36 9-21

37,41 13, 24 13

27,m 41

41 41 41 41 37,41 27,41

41 37,41 41

41 41

41 27, 37, 41 37, 41

37

13.

463

INVERTEBRATE GUANIDINO KINASES

TABLE I (Continued) Phylum

Chordata

Class

Species

Clypeaster japonicus Diadema setosum Echinocardium purpurea Scuphechinus mirabilus Anthocidaris crassispina Echinometra lacunter Echinus esculentus Heliocidaris mthyrogramma Hemicentrotus puleherrimus Lytechinw variegatus Mespilia globulus Paracentrotus lividus Pseudocentrotus depressus Spaerechinus granularis Temnopleurus hardwickii Toxopneustes pileolus Tripneustea esculentus Psammechinus miliaris Ophiuroidea All genera Tunicata Styella mammiculata Other genera Cephalochorda All genera

Kinase

Reference

AK AK AK AK AK, AK, AK, AK, AK, AK, AK, AK, AK, AK, AK, AK, AK, CK CK AK, CK CK

CK CK CK CK CK CK CK CK CK CK CK CK CK CK

a The references refer mainly to papers in which partial summaries of enzyme distribution have been given. These may be consulted for references to the original articles and for the names of those genera which contain only a single guanidino kinase. The abbreviations AK, CK, GK, HTK LK, OK, and TK represent arginine, creatine, glycocyamine, hypot,aurocyamine, lombricine, opheline, and taurocyamine kinase, respectively.

The data relating to the distribution of guanidino kinases, as well as phosphagens (@-44), have been used in an endeavor to gain further insight into the evolutionary transitions that invertebrates have undergone. But the complexities of the distribution patterns mitigate against any straightforward interpretation of the results. The work in this area has been reviewed by Watts (44). On the basis of what is known about the function of phosphocreatine and creatine kinases (37, @), it would appear to follow that the invertebrate guanidino kinases also play an important role in the process of muscular contraction. But little is known about their precise physiological function and the relationship, if any, between the marked structural 42. Y. Robin, Comp. Biochem. Physiol. 12, 347 (1964). 43. X. Van Thoai and J. Roche, Biol. R e v . 39, 214 (1964). 44. D. C. Watts, Advan. Comp. Physiol. Biochem. 3, 1 (1968). 45. S. A. Kuby and E. A. Roltmann, “The Enzymes,” 2nd ed., Vol. 6, p. 515, 1962.

464

J. F. MORRISON

variations of invertebrate muscle (46) and the occurrence of different guanidino kinases, as well as different forms of arginine kinase, has yet to be established.

II. Determination of Enzymic Activity

The same general procedures may be applied to the determination of the initial rates of the reactions catalyzed by each of the guanidino kinases. However, it should be stressed that accurate data are obtained only if the enzyme preparations are free of contaminating enzymes such as ATPase, phosphoamidase, and adenylate kinase. A. CHEMICAL STOPMETHODS Those methods, which involve stopping the reaction after fixed time periods and determining the amount of product formed by chemical means, have been used widely both for assaying enzymic activity and for detailed kinetic investigations. 1. Forward Reaction

Irrespective of whether phosphagen or ADP formation is to be determined, the reaction is stopped with acid. The amount of phosphagen formed is estimated by measuring the release of inorganic phosphate after hydrolysis for 1 min a t 100” in the presence of 0.1 iV HCl (31, 4 7 ) , 0.1 N trichloroacetic acid (32, 4 7 ) , or 0.1 M chloroacetate-perchlorate buffer a t pH 2.5 (16, 4 8 ) . Control experiments are necessary so that any hydrolysis of the nucleotides can be taken into account. Since arginine kinase is inactivated by the addition of acid to p H 2.0 ( d l ) , it is probably true that the other guanidino kinases are also inactivated by this treatment. Thus reaction mixtures can be readjusted to pH 7-8 and the formation of ADP measured by means of the coupled reactions catalyzed by pyruvate kinase and lactate dehydrogenase in the presence of phosphoenolpyruvate and DPNH? (21, 4 9 ) . 46. J. Hnnson and J. Lowry, in “Structure and Function of Muscle” (G. H. Bourne, ed.), 1st ed., Vol. 1, p. 265. Academic Press, New York, 1960. 47. N. Van Thoai and L. A. Pradel, Bull. SOC.Chim. Biol. 44, 641 (1962). 48. W.J. O’Sullivan, R. Virden, and S. Blethen, Eur. J . Biochem. 8, 562 (1969). 49. J. F. Morrison and E. James, BJ 97, 37 (1965).

13. INVERTEBRATE

GUANIDINO KINASES

465

2. Reverse Reaction

The conversion of phosphagen to the corresponding free guanidino compound can be followed very conveniently by stopping the reaction with alkali and adding the diacetyl-a-naphthol reagent described by Rosenberg et al. (50). The absorbance of the red-colored complex is measured a t 535 nm. This has been the preferred procedure in this laboratory for assaying enzymic activity, but its sensitivity is such that phosphagen substrates must contain only very low concentrations of free guanidino compound (20, 21). Attention has been drawn to those buffers which can interfere with color development (45).The formation of ATP is determined as described for ADP except that the coupling system consists of hexokinase and glucose-6-phosphate dehydrogenase as well as glucose and TPN (49).

B. ISOTOPIC METHODS Enzymic activity can also be determined by following the initial rate of the incorporation of [ “C] guanidino compound into the corresponding phosphagen or the incorporation of [l4C]ADP into ATP (or vice versa). The reaction is stopped by applying samples of a reaction mixture to DEAE-cellulose paper which is then developed in different solvents according to whether separation of the nucleotide (51) or guanidino (21, 52) substrates is required, The radioactive spots are excised and counted by liquid scintillation. The general technique, which was used originally in connection with isotope exchange a t equilibrium studies on creatine kinase ( 5 3 ) , has been applied so far only to measurements of initial velocities and partial exchange rates for the arginine kinase reaction (21, 52).

C. CONTINUOCS RECORDING METHODS 1. Spectrophotoinetric Methods I’sing Coupled Enzymes

The coupling systems described above for the estimation of ADP and ATP may be added to reaction mixtures and the reaction rate determined spectrophotometrically by following the absorbance change as a 50. H. Rosenberg, A. H. Ennor, and J. F. Morrison, BJ 63, 153 (1956). 51. J. F. Morrison, Anal. Biochem. 24, 106 (1968). 52. C.Roustan, L. A. Pradel, R. Kassab, and N. Van Thoai, BBA 250, 103 (1971). 53. J. F. Morrison and W. W. Cleland, JBC 241, 673 (1966).

466

J. F. MORRISON

function of time a t 340 nm (54).But it should be noted that a relatively high concentration of KCl (up to 0.1 M ) is required for pyruvate kinase to exhibit its full activity. Therefore, if these procedures are used for comparative studies of the rates of the forward and reverse reactions, allowance must be made for any effects of ionic strength on the velocity of the reverse reaction. Ionic strength effects are of importance with regard to the interaction of bivalent metal ions with the phosphorylated substrates for phosphotransferases ( 5 5 ) . 2. Po tentiometric Met hods

Since the reactions catalyzed by guanidino kinases involve the release of a proton in the forward direction and the uptake of a proton in the reverse direction, their enzymic activities can be determined in either direction by means of the pH-stat technique. This procedure has been used extensively for kinetic investigations on creatine kinase (46, 5 6 ) , but so far it has been applied to only a limited extent for studies on the other guanidino kinases (25, 4 8 ) .

111. Molecular Properties

A. MOLECULAR WEIGHTAND SUBUNIT COMPOSITION Extensive investigations have been made of the molecular weights of the arginine kinases from a variety of invertebrate species using preparations with various degrees of purity. The results, which are summarized in Table I1 (12, 13, 17, 18, 62, 23, 31, 39, 41, 57-61), indicate that these enzymes can be divided into three classes according to their molecular weights. It is interesting that arginine kinases with molecular weights in the vicinity of 40,000 have been found in the Arthropoda and Mollusca while the higher molecular weight forms occur in the Echinodermata and Annelidrt. These changes in molecular weight may well have an N. V. Thiem, G. Lacombe, and N. Van Thoai, BBA 258, 422 (19723. J. F. Morrison and E. Heyde, Annu. Rev. Biochem. 41, 29 (1972). H. K . Jacobs and 8. A. Kuby, JBC 245, 3305 (1970). C. Oriol, M. F. Landon, and N. Van Thoai, BBA 207, 514 (1970). 58. N. Van Thoai, R. Kassab, and L. A. Pradel, BBA 110, 532 (1965). 59. R. Virden, D. C . Watts, R. L. Watts, D. B. Cammack, and J. H. Raper, BJ 54. 55. 56. 57.

99, 155 (1966). 60. N. Van Thoai, in “Homologous Enzymes and Biochemical Evolution” (N. Van Thoai and J. Roche, eds.), p. 199. Gordon & Breach, New York, 1988. 61. Y. Robin, C. Klotz, and N. Van Thoai, BBA 171, 357 (1969).

13.

INVERTEBRATE GUANIDINO KINASES

MOLECULAR WEIGHTS Kinase

TABLE I1 SUBUNITS OF GUANIDINO KINASES

-4ND

Source

Callinedus sapidus Cancer pagurua Carcinus maenas chhmy8 opermlaris Dugesiella hentzi Negative Neutral Homarw, americanus

Arginine

467

Molecular weight 37, oooa 39,50@ 38,000 37,000 37,000 38,000 40,000,0

Subunits 1

Reference 13

67 41

41 13 1

13 19,18

1

17, 67-69

2

41 41 41 39 29, 39, 60

38,OOoa Homarus vulgaris

44,000,o 37,00@

Limulua polyphemus Negative Neutral Loligo sp. Melanopus h m e r i Pagurus bernhardus Palinurus vulgaris Pecten muximus Potamobiw aatacus Antedon lnfida Asteria rubens Astropecten irregularis Cummaria elongata Echinus esculentus

Glycocyamine Hypotaurocyamine Lombricine Taurocyamine

38,000. 38,000. 38,500 38,000 38 ,oooa 39,000 38,000 43,00@ 84 ,000 81 ,000 86 000 78,500 81 00094,000 Holothuria forskali 81,000 Luidia ciliaris 89,000 80,500 Marthastmias glacialis 80,000 Paracentrotus lividus Sipunculus nudus 86,500a Solaster papposa 86,000 Stichopus tremulua 89,000 Travisia forbesii 80 ,000 Sabella pavonina 150,00@ Spirographis spallanzanii 150,000 Nephtys weca 89,000 P h c o l o s m a vulgare 83,000 Lumbricus terrestris 80, oooa Arenicola marina 81, oooa ~

41

2

41 39 23, 61 61 31, 68

2 2

68 67, 68 67,68

~~

These molecular weights were determined by ultracentrifugation, gel filtration, and density gradient centrifugation using homogeneous preparations of the enzymes. The unmarked values were obtained from studies with crude extracts or partially purified enzyme preparations which were subjected to gel filtration and/or density gradient centrifugation

.

468

J . F. MORRISON

evolutionary and physiological significance which is not apparent a t the present time (44, 6 2 ) . The low molecular weight arginine kinases from crabs ( P . astacus and C. pagurus) and lobsters ( H . vulgaris and H . americanus) have been shown to consist of a single polypeptide chain. By contrast, the enzyme from the marine annelid, 8. nudus, with approximately twice the molecular weight is made up of two similar, if not identical polypeptide chains. The exact relationships between what appears to be monomeric, dimeric, and tetrameric forms of arginine kinase remain to be determined. As far as the author is aware, no X-ray diffraction studies are being undertaken on arginine kinase. The other guanidino kinases listed in Table I1 are similar t o creatine kinase in having a molecular weight around 80,000 and in being composed of two subunits. Although these enzymes appear to be a closely related group, it is too early to conclude that polymeric forms do not exist since the number of species investigated is relatively small. I n this connection it is salutary to recall that there was a time when it was believed that all arginine kinases had a molecular weight in the region of 40,000 and that it was this property which distinguished this enzyme from the other guanidino kinases.

B. STABILITY The stability of the arginine kinases appears to be related to their molecular weights. The enzymes which have been isolated from crayfish (20, 21) and lobsters (14, 16, 18) and which have molecular weights of about 40,000 are stable while the higher molecular weight forms from other sources are not. Thus the arginine kinases from S. nudus (22) and S. pavonina (23) with molecular weights of 86,500 and 150,000, respectively, are labile even in the presence of a reducing agent. No difficulties have been encountered in maintaining fully active preparations of lombricine kinase and taurocyamine kinase, but some uncertainty exists about the stability of glycocyamine kinase. The enzyme has been reported both as stable (30)and unstable (31). Arginine kinase from lobster muscle is capable of reforming a significant part of its native structure after treatment with 8 M urea as judged by the restoration of activity on dilution into buffers (18). The rate of of recovery of enzymic activity is increased in the presence of L-arginine or the magnesium complexes of ADP or ATP, but not by the addition of analogs of L-arginine. Reactivation is also promoted by thiol com62. D. C. Watts, in “Studies in Comparative Biochemistry” (K. A. Munday, ed.), p. 162. Pergamon, Oxford, 1965.

13.

INVERTEBRATE GUANIDINO IEINASES

469

pounds. It would be interesting to determine what other guanidino kinases can undergo reversible inactivation in urea and to use these enzymes for mixed hybridization studies.

C . AMINOACIDCOMPOSITION Analyses of the amino acid composition of arginine kinase from eight invertebrate sources (13, 18, 60, 63, 6 4 ) , as well as glycocyamine kinase, lombricine kinase, and taurocyamine kinase (60), have indicated that none of the enzymes contain disulfide bridges and that the number of half-cysteine residues per mole shows considerable variation (cf. Table VI). Attention has also been drawn to the similarity of the amino acid composition of all the guanidino kinases, including creatine kinase, and the relative constancy of the values for the sum of the acidic, basic, or hydrophobic amino acid residues per 1OOg of protein has been noted (60, 6 3 ) . These findings have been considered to point to the homology of the guanidino kinases, but a preferable basis for such a conclusion would be the comparison of the complete amino acid sequence for each enzyme. Such information is not yet available although there are reports of the sequences of amino acid residues in the region of the essential thiol groups of arginine kinase (65) and lombricine kinase (66). The similarity of the tryptic peptides obtained from these two enzymes and their close relationship with the corresponding peptides from rabbit muscle (67, 68) and ox brain (69) are illustrated in Table 111. However, it will be noted that the insertion of threonine (or glycine) next to proline in the invertebrate enzymes is compensated for by the deletion of glycine from near the,C-terminal end of the vertebrate creatine kinases. Other replacements of amino acid residues are of the conservative type. A further striking feature of all the sequences is the presence of proline adjacent to the essential cysteine residue. 63. A. R. Thomson, J. W. Eveleigh, J. F. Laws, and B. J. Miles, in “Homologous Enzymes and Biochemical Evolution” (N. Van Thoai and J. Roche, eds.), p. 255. Gordon & Breach, New York, 1968. 64. R. Virden and D. C. Watts, BJ 99, 159 (1966). 65. E. Der Terrossian, L. A. Pradel, R. Kassab, and N. Van Thoai, Eur. J . Biochem. 11, 482 (1969). 66. E. Der Terrossian, G . Desvages, L. A . Pradel, R. Kassab, and N. Van Thoai, Eur. J. Biochem. 22, 585 (1971). 67. A. R. Thomson, J. W. Eveleigh, and B. J. Miles, Nature (London) 203, 267 (1964). 68. T . A. Mahowald, Biochemistry 4, 732 (1965). 69. R. S. Atherton, J. F. Laws, B. J. Miles, and A. (1970).

R. Thomson, BJ 124 589

470

J. F. MORRISON

TABLE I11 IN THE REGIONOF THE REACTIVE THIOLGROUPSOF AMINOACIDSEQUENCES ARGININE, LOMBRICINE, AND CREATINE KINASES Kinase

Amino acid sequence

Arginine -Gln-Thr-CysSH-Pro-Thr-Ser-Asn-Leu-Gly-Thr-Val -Arg( H . vulgaris; ref. 66) Lombricine -Tyr-Ile--Thr-Cys.SHPro-Gly-Ser-Asn-Leu-Gly-Thr~Leu-Arg(L. te?'TeStTiS; ref. 66) Arg-Ty r-Val-Leu-Thr-Cys.SH-Pro--Ser-Asn-Leu-Gly-Thr-Gly-LeuCreatine (Rabbit muscle; refs. 67, 68) Creatine -Tyr-Ile -Leu-Thr-Cys.SH-Pro-Ser-Asn-Leu-Gly-Thr-Gly-Leu-Arg(Ox brain; ref. 69)

The carboxyl terminal sequences of the arginine kinases from H . vulgaris and S. nudus have been determined to be valylmethionine and leucyllysine, respectively (70).

D. IMMUNOLOGICAL REACTIONS From comparative studies of the immunological properties of various arginine kinases and other invertebrate guanidino kinases, it has become apparent that these enzymes may, or may not, possess common structural features ( I S , 40). Thus antiserum to the negative form of arginine kinase from L. polyphemus was found to react with the neutral form of the enzyme from the same source and with honeybee, spider, and tarantula arginine kinases, but not with any of the crustacean arginine kinases which were tested ( I S ) . On the other hand, antiserum to H . vulgaris arginine kinase was shown to react to variable extents with the same enzyme from other crustacean sources, although not with arginine kinases from echinoderms, mollusks or marine worms, or with lombricine and taurocyamine kinases (40). Further, no cross-reactions were detected between arginine kinase from 8. nudus and that from other sources with either the same or different molecular weights. While cross-reactions have been observed between three annelid guanidino kinases, vie., lombricine kinase, opheline kinase, and taurocyamine kinase, no such reac70. F. Regnouf, L. A. Pradel, R. Kassab, and N. Van Thoai, BBA 194, 540 (1969).

13.

INVERTEBRATE GUANIDINO KINASES

471

tion occurs between hypotaurocyamine kinase from several sipunculids and the antiserum to taurocyamine kinase (40). All the invertebrate guanidino kinases are inhibited, although incompletely, by their specific antisera. However, heterologous antisera do not cause inhibition and hence it has been concluded that antibodies are formed against sites other than those concerned with catalytic activity (40). The immunological data obtained with guanidino kinases have also been interpreted to indicate the conservation of common structure that has occurred in the course of phylogenetic evolution of these enzymes (40).

IV. Catalytic Properties

A. ACTIVATIONBY BIVALENT METALIONS Although it is well known that the invertebrate guanidino kinases exhibit catalytic activity only in the presence of bivalent metal ions, the elucidation of the precise function of metal ions in enzymic catalysis has attracted little attention. A limited number of qualitative investigations of the metal ion specificity of the enzymes has been made by testing the effects of fixed concentrations under standard experimental conditions. But, for the reasons elaborated below, the results of such studies are not very informative. Nevertheless, from the data listed in Table IV (16, 19, 25, 31, 32,34-36, 47, 71), it is apparent that all the invertebrate guanidino kinases are activated by Mgz+and that ions such as Cu*+,Znz+, and CdZ+do not activate arginine kinase, lombricine kinase, or taurocyamine kinase. The latter result is not surprising since these metal ions would undoubtedly react with the thiol groups that are essential for activity (cf. Table VI). Further investigations of the ability of Ca2+to activate arginine kinases would seem to be warranted. The present results indicate that the enzyme from the northern hemisphere lobster is activated by CaZ+while the effect is not observed with arginine kinase from the southern hemisphere crayfish. From the data obtained with creatine kinase and other phosphotransferases, it might be anticipated that the metal ion ( M ) activation of the invertebrate guanidino kinases would also involve the nonenzymic formation of metal-nucleotide complexes which act as substrates together with the free forms of the guanidino compounds ( 5 5 ) . This being so, the general form of reactions in the presence of adenine nucleotides 71. L. A. Pradel, R. Kaeaab, and N. Van Thoai, BBA 81, 86 (1964).

472

J. F. MORRISON

TABLE IV THESPECIFICITY OF GUANIDINO KINASESFOR BIVALENT METALIONS Metal ion Kinase

Source

H . vulgaris

Arginine

Hypo taurocyamine Lombricine

M%+, Mn2+, Co2+, Ca2+, Sns+, Fe2+ M&+, Mn2+

Ni2+, Cu*+,Ma+, Bas+, Ch+, Zns+

Reference I6

19 Bas+, Cap+, Srz+, Nil+, F@+,Calf, Cd+, Cd2+, Zn*+, Fe3+,A13+ 96 L. polyphemus Mgz+, M d + , Gas+ Zn'f, CUB+,F#+, Fe3+ N . coecu M@+, Mnz+, Gas+ 31 , 71 P . vulgare Mg2+ 36

J. verreauxi

Gly cocyamine

Inactive or inhibitory

Active

M . cameroni

Taurocyamine

A . marina

Opheliie

0 . neglecta

Mg2+,CO~+, Mn", Ca2+

Baa+, Srl+, Nip+, CU*+, 32, Fez+, F@+,Ala+,Sna+, $4 Cdz+, Zn2+, Be2+ Mgz+, Mns+, Ca2+ Nil+, Fee+,SP+, 47 Co2+, Zne+ Mg2+ 56

should be written as shown in formulation (2). On this basis, determina-

+ MATF-

+ MADP- + H+

(2) tions of metal ion specificity relate, in effect, to examinations of which metal-ATP (or metal-ADP) complexes participate in the reaction sequences. Detailed studies of this type are subject to many complications which include knowledge of the actual concentration of metal-nucleotide complex as well as the inhibitory effects of free nucleotide and excess free metal ion. Procedures for circumventing these difficulties have been outlined (66) and utilized (21, 72) to show that the kinetic data for the arginine kinase reaction in the presence of Mg2+ are consistent with formulation (2). Further support comes from the results of nuclear magnetic resonance studies which indicate that the arginine kinases from lobsters do not undergo direct reaction with Mn2+ (48). However, determinations have not been made of the relative efficiency of a range of metal ions. This information, as well as an insight into metal ion function in catalysis, can be gained from kinetic experiments which are designed to elucidate the mechanism of the reaction and to determine Guanidino compound

phosphagen

72. E. Smith and J. F. Morrison, JBC 246, 7784 (1971).

13.

INVERTEBRATE GUANIDINO KINASES

473

values for the various kinetic parameters when different metal-nucleotide complexes are used as substrates. The kinetic studies carried out with MgADP-, MnADP-, and CaADP- as substrates for creatine kinase (73) might be considered as a prototype for future investigations on the role of metal ions in the reactions catalyzed by invertebrate guanidino kinases. In this connection, it is suggested that the trivalent rare earth metal ions might be included among those to be tested as, in many respects, their properties resemble those of the alkaline earth and transition metal ions ( 7 4 ) .

B. SUBSTRATE SPECIFICITY 1. Some General Comments

The majority of studies on the substrate specificity of guanidino kinases have been qualitative, with determinations being made of whether or not reaction occurs on the addition of a compound a t a fixed concentration and under a chosen set of experimental conditions. Investigations of this type suffer from the weakness that compounds which act as poor substrates may be overlooked. They are useful as preliminary screening tests, but compounds which do not appear to be substrates should be retested a t higher concentrations using increased concentrations of enzyme, The enzyme must, of course, be of a high degree of purity. Occasionally, comparisons have been made of the rates a t which different substrates undergo reaction (35). But such results are of limited value since variations in initial velocities can result from the substrates giving rise to different maximum velocities and/or from the differences in their Michaelis constants. While it is of interest to know if substrate analogs can participate in an enzymic reaction, it is more important for the understanding of enzymic catalysis to have data relating to the strength of their binding to various enzyme forms and the way in which they influence the maximum rate of product formation. Kinetic investigations can yield such information as well as information about compounds which bind to the enzyme but do not undergo reaction. Consequently, it is possible to distinguish the structural requirements of a compound for binding from those which are required for catalysis. To date, the only invertebrate guanidino kinase to which detailed kinetic techniques have been applied is arginine kinase (21, 72). 73. J . F. Morrison and M. L. Uhr, BBA 122, 57 (1966). 74. D. W. Darnall and E. R. Birnbaum, JBC 245, 6484 (1970).

J. F. MORRISON

2. Guanidino Substrates

In Table V a comparison is made of the ability of the various invertebrate guanidino kinases to act on what can be considered as the natural substrates for each of the seven known guanidino kinases. The results show that the enzymes fall into two categories with respect to their substrate specificities. Thus arginine kinase and glycocyamine kinase exhibit a narrow specificity while the other four guanidino kinases have a somewhat broader specificity. The characteristic substrates for the latter enzymes possess some common structural features (11, IV-VI) which apparently can explain why they undergo reaction with each enzyme (60). While it has been reported that arginine kinase from S. nudus has an absolute specificity for arginine (22),this is not so for the enzymes from J . verream. (19),S. pavonina (23),and L. polyphemus (26)since they catalyze reactions with some closely related analogs such as homoarginine and canavanine. The high molecular weight arginine kinase from 8. pavonina (23)differs from the enzymes from other sources (22, 26,72) in being able to utilize D-arginine as effectively as the L-isomer. Although creatine is not a substrate for the arginine kinase from P . longipes, it does combine a t the binding site on the enzyme for arginine since it causes linear competitive inhibition in relation to that substrate (72).On the other hand, D-arginine and canavanine do not interact with the enzyme since neither compound functions as a substrate or inhibitor (72).However, these compounds do combine with other arginine kinases. Canavanine acts as a competitive inhibitor with respect to both D- and L-arginine of the S. pavonina enzyme (23)while D-arginine functions as a competitive inhibitor in relation to L-arginine of the negative and neutral enzymes from L. polyphemus (26). From kinetic studies of the taurocyamine kinase and hypotaurocyamine kinase reactions in the presence of a fixed concentration of MgATP2-, it has been found that the apparent maximum velocity for each reaction is the same whether taurocyamine or hypotaurocyamine is the substrate (36).However, for the taurocyamine kinase reaction the apparent K , for taurocyamine is less than that for hypotaurocyamine while the reverse is true for the hypotaurocyamine kinase reaction, When these results are considered in relation to the differential abilities of the two enzymes to utilize lombricine as a substrate (Table V), and to the differences in their immunological properties ( 3 9 ) ,it appears that there is justification for regarding taurocyamine kinase and hypotaurocyamine kinase as distinct entities. However, it would be advantageous to have additional information about their kinetic and physicochemical proper-

TABLE V SPECIFICITY OF GUANIDINO KINASES FOR GUANIDINO COMPOUNDS

0

9

5

8

Guanidino compound Kinase Arginine Glycocyamine Hypotaurocyamine Lombricine

Opheliie Taurocyamine

I-Arginine Creatine Glyocyamine Hypotaurocyambe Lombricine

+

-

-

-

-

+

-

-

-

-

-

+

-

-

+

-

+

-

-

Taurocyamine

Opheline

Ref.

-

19,891

-

-

60,7.9 88, 60,

-

+ + + +

36,60 39, 33, 347 60 36, 60 989 36,

+ + +

Yl

+ +

+

4r160

3 zE g ej

476

J. F. MORRISON

ties. The same comment applies to lombricine and opheline kinases which seem to have similar substrate specificities (Table V ) . Studies on opheline kinase have shown that the apparent maximum velocities of the reactions decrease in the order taurocyamine > lombricine > opheline while the apparent K , values for these substrates decrease in the same order ( 3 6 ) . Thus this enzyme and those concerned with the phosphorylation of taurocyamine and hypotaurocyamine are named after the substrate which has the lowest K , value. Both L- and D-lombricine have been shown to act as substrates for lombricine kinase ( 3 4 ) . 3. Nucleotide Substrates

The specificity of invertebrate guanidino kinases for nucleotide substrates has not been well studied, and most of the available data relates to arginine kinase. It has been claimed (23) that the enzyme from 5. pavonina is specific for ATP since no reaction was observed in the presence of CTP, GTP, ITP, or UTP under standard assay conditions. On the other hand, the enzyme from P . longipes has been shown (72) to catalyze the reaction with the magnesium complexes of ADP, dADP, GDP, IDP, and UDP, although the Michaelis and dissociation constants, as well as the maximum velocity of the reactions, differ markedly according to the identity of the nucleotide. Indeed, because of the differences in the values of these parameters, it was possible to study the inhibition of the reaction by MgCDP-, MgGDP-, and MgUDP- with MgADP- and phosphoarginine as substrates ( 7 2 ) .In addition to ATP, the two arginine kinases from L. polyphemus can utilize dATP, GTP, and ITP as phosphoryl group donors (255). The manganese complexes of dADP and dATP, as well as the corresponding ribonucleotides, have been demonstrated to function as substrates for the arginine kinases from H . wulgaris and H . amem'canus ( 4 8 ) .Lombricine kinase is also capable of phosphorylating ADP and dADP, while no significant reaction occurs with CDP, GDP, IDP, or UDP (32). Because the arginine kinases exhibit different kinetic parameters for the corresponding ribo- and deoxyribonucleotide substrates it must be concluded that the sugar moiety plays an active role in catalysis (48, 7 2 ) .

C. pH OPTIMUM The reactions catalyzed by the invertebrate guanidino kinases are characterized by having pH optima of pH 8.4-9.1 in the forward direction and pH 6.6-7.2 in the reverse direction (19, 28, 32, 34-36, 47, 71) with Mg2+as the activating metal ion. No comparisons have been made

13. INVERTEBRATE

GUANIDINO KINASES

477

of the influence on the pH optima of different bivalent metal ions which activate the enzymes. The multiplicity of factors responsible for the variation of the initial velocities of these reactions as a function of pH is not always appreciated. Therefore, it would seem worthwhile to give some general indication of these factors by referring to the involvement of ADP in the reverse reactions. This nucleotide has a pK, value in the vicinity of the pH optimum so that the relative concentrations of ADPS-, HADP'-, metal-ADP-, and metal-HADP would change over the pH range used. Thus, if the metal-HADP complex were inert, the metal-ADP- complex functioned as the nucleotide substrate, and the free nucleotide species were inhibitory, the enzymic activity would tend to decrease as the pH is lowered both because of the reduction in the concentration of the true substrate and the increase in the concentrations of the free nucleotides. The initial velocities would, of course, vary with the identity of the metal ion. Apart from effects resulting from differences in the intrinsic properties of the metal ions, the stability constants of the metalADP complexes would determine the distribution of ADP among its various species ( 5 5 ) . Because pH can also influence the values for the kinetic parameters of the reaction, the interpretation of simple pHactivity data is precluded.

D. FUNCTION OF AMINOACID RESIDUES 1. Cysteine Residues

Initial studies of the effect of thiol reagents such as iodoacetate, iodoacetamide, p-mercuribenzoate, chloroacetophenone, and N-ethylmaleimide on the activity of arginine kinase (19), glycocyamine kinase (71), hypotaurocyamine kinase (35), lombricine kinase ( 3 2 ) , taurocyamine kinase (75), and opheline kinase (36) demonstrated the importance of thiol groups for their catalytic function. More recently, considerable effort has been devoted to quantitative studies of the reaction of thiol reagents with those kinases that have been obtained in pure form. Apart from the determination of the number of reactive and total -SH groups that each of the enzymes contain, estimates have been obtained for the minimum number of these groups which must undergo reaction before complete loss of enzymic activity occurs. The results of these investigations are summarized in Table VI. When the data of Table VI are considered together with the subunit 75. N. Van Thoai and L. A. Pradel, Bull. Sac. Chim. Bwl. 44, 1089 (1962).

TABLE VI REACTIVE, TOTAL, AND ESSENTIAL THIOLGROUPS OF GUANIDINO KINASES

No. of S H groups per mole

Amino acid analysis

No. of S H groups/mole essential for activity

6

6

1

5

5 12 22 6 16

6 2 1 2

Titration with thiol reagents Kinase Arginine

Source

H.vulgaris H.amerieanua S. nudus

Glycocymine Lombricine Taurocyamhe

N . coeca L. terreslris A . marina

- urea 6 5 12

6 1 2

+ urea

12 16 (22) 6 14 (16)

Reference

18 22 60, 76 29, 60, 76 29, 60, 76

4 r

13.

INVERTEBRATE GUANIDINO KINASES

479

composition of the enzymes (Table 11), it becomes apparent that arginine kinase from H. vulgaris,glycocyamine kinase, and taurocyamine kinase possess one essential thiol group per polypeptide chain of molecular weight of about 40,000. However, arginine kinase from S. nudus and lombricine kinase do not fit this pattern. It should also be mentioned that estimates of the number of reactive thiol groups can vary according to the experimental conditions and the nature of the thiol reagent used. Thus studies of the inhibition of H. vulgaris arginine kinase by N-ethylmaleimide a t pH 7.0 (14) or DTNB [5,5’-dithiobis (2-nitrobenzoic acid)] a t pH 8.5 (76) have indicated the presence of three -SH groups a t the active center of the enzyme whereas treatment with DTNB at pH 7.0 shows that there is complete loss of activity after the reaction of only one thiol group ( 7 6 ) . Evidence for the involvement of a single -SH group in the catalytic activity of arginine kinase has been presented by Virden and Watts ( 7 7 ) . These authors showed that the reaction of iodoacetamide with the enzyme is pH independent and suggested that the -SH group is hydrogen-bonded to a histidine residue a t the active site. Glycocyamine kinase is similar to arginine kinase from H. vulgaris in that six thiol groups per mole react rapidly with DTNB ( 7 6 ) . However, stoichiometric titration of the former enzyme with DTNB has shown that the reaction of two thiol groups per mole, or one per polypeptide chain, is sufficient to cause complete inactivation. The ability of guanidino and nucleotide substrates to protect against the inactivation of guanidino kinases by thiol reagents has been investigated in an endeavor to elucidate the role of thiol groups in the catalytic mechanism. The studies have been largely qualitative although some have involved determinations of the pseudo-first-order rate constants for the reaction of thiol reagents with the kinases in the absence and presence of a substrate (48, 62, 7 7 ) . Several investigators have reported that L-arginine can protect various arginine kinases against inactivation by thiol reagents (IS, 14, 18, 22, 76-79) and concluded that the binding of this substrate to the enzymes takes place a t or near the essential -SH group. But in view of the fact that both the guanidino and nucleotide substrates of creatine kinase can protect against the inhibition by iodoacetate and still bind to the enzyme after carboxymethylation of 76. R. Kaasab, L. A. Pradel, E. Der Terrossian, and N. Van Thoai, BBA 132, 347 (1967). 77. R. Virden and D. C. Watts, BJ 99, 162 (1966). 78. C. Roustan, R. Kassab, L. A. Pradel, and N. Van Thoai, BBA 167, 326 (1968). 79. C. Roustan, E. Der Terrossian, and L. A. Pradel, Eur. J. Biochem. 17, 467 (1970).

480

J. F. MORRISON

its reactive thiol groups ( 5 5 ) , the above conclusion was not necessarily correct. It was important, therefore, to perform binding experiments with carboxymethyl-arginine kinase and these established that L-arginine is not bound to the modified enzyme ( 7 8 ) . Interestingly, it was found that isoleucine, valine, norleucine, citrulline, and ornithine, which protect against the inhibition of arginine kinase by thiol reagents, are bound to the native enzyme, but not to the carboxymethyl enzyme. Although nucleotide substrates are bound to the carboxymethyl derivative of arginine kinase ( 7 8 ) , they afford some protection against ,the inactivation by thiol reagents of the enzyme from various sources (18, 48, 7 7 ) . Thus it would seem that they exert their protective action by bringing about a conformational change which affects the reactivity of the thiol group. Less extensive investigations have been undertaken on the function of thiol groups in the reactions catalyzed by the other invertebrate guanidino kinases. A limited amount of information is available concerning their inhibition by thiol reagents in the absence and presence of substrates, but it is considered to be inconclusive (71, 7 6 ) . I n connection with experiments of the aforementioned type, it should be borne in mind that a substrate will influence the effect of an inhibitory thiol compound only if it combines with the free enzyme. Therefore, it is valuable to have a knowledge of the order in which the substrates add to the enzyme in the catalytic sequence. The second substrate to add to the enzyme that catalyzes a reaction conforming to a compulsory ordered mechanism cannot be expected to manifest any protection. Further, it must be realized that the degree of protection by a substrate which reacts with free enzyme will be determined by its concentration relative to the dissociation constant for the enzyme-substrate complex and not by its absolute concentration. Since the reactions catalyzed by creatine kinase, and arginine kinase from P . lonyipes, proceed via random mechanisms involving the formation of dead-end ternary complexes ( 5 5 ) , it is possible that other guanidino kinases exhibit a similar reaction mechanism. Consequently, it would seem to be worthwhile testing the effect, on the rate of enzyme inactivation by thiol reagents, of those substrate-product pairs which could give rise to dead-end complexes. Such studies have yielded useful data about the catalytic mechanism for the creatine kinase reaction ( 5 5 ) .

2. Other Amino Acid Residues The ease with which the thiol groups of guanidino kinases can be modified and the unavailability of specific reagcnts for the modification

13.

INVERTEBRATE GUANIDINO KINASES

48 1

of certain other amino acid residues have hampered the elucidation of the location of the latter residues and their functional role in the catalytic mechanism. The problems associated with these studies have been overcome to a large extent by initially reacting the thiol groups of the enzyme with tetrathionate to form the S-sulfenyl sulfonate derivative and then with the selected reagent before unmasking the thiol groups by treatment with dithiothreitol (80, 81). However, there are disadvantages in the use of this approach. Studies of the effect of guanidino substrates on the modification of amino acid residues are precluded because they are not bound to the S-sulfenyl sulfonate derivatives of a t least some guanidino kinases. Further, the initial modification can cause conformational changes which alter the reactivity of the derivative toward a modifying reagent as compared with that of the native enzyme (80). Nevertheless, it has become apparent that useful information about the involvement of amino acid residues in catalysis can be gained from application of the general procedure described above. Most modification studies have been carried out with arginine kinase from H . vulgaris and these have indicated the importance of lysyl, tyrosyl, and histidyl residues for its catalytic activity. It has been shown that dansyl chloride reacts with an €-amino group of a lysine residue on the enzyme to cause inhibition and that complete loss of activity occurs on the binding of one dansyl group per mole (81).Both arginine and nucleotide substrates protect against the inhibition (81), but whereas the nucleotide substrates bind to the dansylated enzyme arginine does not (82). A tyrosyl residue was implicated in the catalytic mechanism when it was found that reaction of the enzyme with iodine or N-acetylimidazole caused inhibition (80). I n the presence of lower concentrations of iodine, one of the 10-11 tyrosyl residues is converted to monoiodotyrosine with the concomitant loss of total activity. By contrast, complete loss of activity on reaction with N-acetylimidazole is associated with the formation of almost three 0-acetyltyrosine residues. Full enzymic activity can be restored by treatment of the modified enzyme with hydroxylainine which is able to remove all the 0-acetyl groups. Further evidence for the involvement of a tyrosyl residue comes from studies with tetranitromethane (83)which demonstrated that nitration of one tyrosyl residue results in complete loss of enzymic activity. I t appears that nitration of arginine kinase reduces the a-helix content 80. A. Fattoum, R. Kassab, and L. A. Pradel, Eur. J. Biochem. 22, 445 (1971). 81. R. Kassab, C. Roustan, and L. A. Pradel, BBA 167, 308 (1968). 82. C. Roustan, L. A. Pradel. R. Kassab, A. Fattoum, and N. Van Thoai, BBA 206, 369 (1970). 83. R. Kassab, A. Fattoum, and L. A. Pradel, Eur. J. Biochem. 12, 264 (1970).

482

J. F. MORRISON

of the enzyme but causes little alteration to its ternary structure (84). Solvent perturbation studies with ethylene glycol have suggested that 4 tyrosine residues are exposed on the surface of the native enzyme at neutral pH (84). While the 8-carboxymethyl (78) and S-sulfenyl sulfonate (80) derivatives of arginine kinase bind nucleotide substrates, the iodinated and nitrated enzymes, with or without free -SH groups, fail to react with any substrate (80, 8 3 ) . The latter results would seem to be related to the structural changes that occur on modification of the enzyme. At pH 6.0 diethylpyrocarbonate reacts specifically with 1-2 histidyl residues of arginine kinase to form the carbethoxyhistidyl derivative which is inactive (52, 8 5 ) . The reagent also reacts with the enzyme after modification with tetrathionate and its rate of reaction with the native enzyme is reduced in the presence of arginine, MgADP-, or h4gATPz- ( 8 5 ) . Since these substrates can bind to the carbethoxylated enzyme ( 8 d ) , it is clear that the combination of each substrate with the native enzyme must induce conformational changes which decrease the reactivity of the histidyl rksidues. In contrast to arginine kinase, taurocyamine kinase is neither inhibited nor nitrated by tetranitromethane ( 8 3 ) .Both these enzymes (14, 7 5 ) ,as well as glycocyamine kinase ( 7 1 ) , are inhibited by hydroxylamine, but inhibition has not been observed with hypotaurocyamine kinase ( 3 5 ) .

V. Reaction Mechanism

Initial velocity studies have demonstrated that the reactions catalyzed by arginine kinases from H . vulgaris (16, 5 d ) , S. pavonina ( 2 3 ) , L. polyphemus ( 2 5 ) , and P . longipes (21) conform to sequential mechanisms that allow for the addition of both substrates to the enzyme before either product is released. But it is only the enzyme from the latter source that has been subjected to detailed kinetic investigations in order to determine the sequence of substrate addition and product release. The results of product (21) and dead-end (72) inhibition studies led to the conclusion that the reaction exhibits a rapid equilibrium, random mechanism which involves the formation of two dead-end complexes, viz., enzyme-MgADP-arginine and enzyme-MgATP-phosphoarginine. However, as the isotope exchange rates a t equilibrium between like substrate-product pairs are not equal (21), the rapid equilibrium 84. M. F. Landon, C. Oriol, and N. Van Thoai, BBA 214, 168 (1970). 85. L. A. Pradel and R. Kassab, BBA 187, 317 (1968).

13.

483

INVERTEBRATE GUANIDINO KINASES

condition must be only approximated under steady-state conditions which involve net reaction. The equality of the equilibrium exchange rates with the negative form of the L. polyphemus arginine kinase suggests that the reaction conforms to a truly rapid equilibrium, random mechanism (26). The random addition of substrates to arginine kinase from H . vulgaris can be inferred from the ability of arginine and nucleotides to protect against its inactivation by thiol and other reagents (Sections IV,D,l and 2) while direct evidence comes from the binding data obtained with both types of substrate (48, 78, 82). The interaction of nucleotide substrates with arginine kinase from H. americanus (48), lombricine kinase (1 mole/mole) , and taurocyamine kinase (2 moles/mole) (82) has also been demonstrated using either magnetic resonance or difference spectrophotometric techniques. Further, the conclusion has been reached that each of these enzymes, as well as arginine kinase from H . vulgaris, undergoes conformational changes as a result of the formation of an enzyme-nucleotide complex (48, 89). Of special interest and significance are the reports that arginine kinase from both crayfish (21) and lobster (52) muscle can catalyze partial exchange reactions between arginine and phosphoarginine in the absence of a nucleotide substrate, as well as between ADP and ATP in the absence of a guanidino substrate but in the presence of Mg2+ions. Such findings have led to the suggestion that the reactions can proceed not only via a sequential mechanism but also by means of a ping-pong mechanism which involves the formation of a phosphorylated form of enzyme. However, since the exchange reactions are slow compared with the rates of the overall reactions, then a t the most, only a small proportion of the total reaction flux could proceed through the ping-pong sequence. From the kinetic data obtained with crayfish and lobster arginine kinases, it appears reasonable to propose that when both substrates are present and central ternary complexes are formed, the series of reactions which occur within these complexes involve the transfer of a phosphoryl group to and from the enzyme. Attempts have been made in this laboratory to isolate a phosphorylated form of arginine kinase, but they have not been successful and the failure may result from the intrinsic ATPase activity of the enzyme (48, 55, 86). The concept that the ability of arginine kinase to catalyze both partial exchange reactions is an inherent property of the enzyme is supported by the recent work of Roustan e t a,?. ( 5 2 ) . These authors showed that dansylation or alkylation of arginine kinase from H . vulgaris, which causes inactivation 86. C.

T.Walsh and L. B. Spector, ABB 145,

1 (1971).

484

J. F. MORRISON

and prevents the binding of arginine, eliminates the arginine-phosphoarginine exchange but not the ADP-ATP exchange. In addition, they found that modification of an essential histidyl residue with diethylpyrocarbonate, which gives rise to an inactive form of enzyme that is still capable of binding guanidino and nucleotide substrates, eliminates both partial exchange reactions. In contrast to the intersecting initial velocity patterns obtained with the aforementioned arginine kinases, those for the enzymes from J . verreauxi (20) and S. n u d w (222) appear to consist of families of parallel straight lines. Moreover, initial velocity data obtained by varying the substrates in constant ratio gave linear double reciprocal plots, and partial exchange reactions were demonstrated with the former enzyme. But because the partial exchange reactions were relatively slow and since a nonintersecting initial velocity pattern is an insufficient criterion to establish a ping-pong mechanism (55), further detailed kinetic investigations are required before definitive conclusions can be reached about the reaction mechanisms for these two enzymes. It seems likely that they will be found to exhibit mechanisms similar to those already described for the arginine kinases from H . vulgaris and P . longipes. However, the ability to catalyze partial exchange reactions cannot be a general characteristic of guanidino kinases or even of arginine kinase. Neither taurocyamine kinase (52) nor arginine kinases from S. pavonina (23) and L. polyphemus (25) are able to catalyze such reactions. Kinetic investigations on guanidino kinases have been performed under a variety of conditions with respect to pH and buffer composition and sufficient attention has often not been paid to the control of the ionic species of substrates present in the reaction mixtures ( 5 5 ) . Hence it is considered that there would be little merit in recording for comparative purposes the values that have been obtained for the kinetic constants associated with the interaction of substrates with different forms of the various enzymes and for the maximum velocities of each reaction in the forward and reverse directions. It is unfortunate that such comparisons cannot be made and this situation prompts the suggestion that, unless there are good reasons for not doing so, future kinetic and thermodynamic experiments on guanidino kinases be conducted at 30" under standard conditions. It is proposed that consideration be given to the use of the nonchelating buffer N-ethylmorpholine (0.1M) a t pH 8.0 where both ATP and ADP exist in their fully ionized forms. An added advantage of these conditions is that they have already been used for the determination of the stability constants for MgATP2- and MgADP- (87). 87. W. J. O'Sullivan and D. D. Perrin, Biochemistry 3, 18 (1964).

13.

INVERTEBRATE GUANIDINO KINASES

485

Since the forward and reverse reactions catalyzed by arginine kinase from P. longipes have been studied kinetically a t the same p H (pH 8.0) and under carefully controlled conditions, a comparison can be made of the kinetic constants for each of the four reactants (21).Such a comparison reveals that (a) M g A D P (or phosphoarginine) binds more strongly to the free enzyme than MgATP2- (or arginine) and (b) the presence of phosphoarginine (or MgADP-) on the enzyme markedly hinders the combination of MgADP- (or phosphoarginine) while MgATP2- and arginine react independently with the enzyme (cf. 25,52). In a recent report (54) the inhibition of two arginine kinases by monovalent anions and the effect of C1- ions on the kinetics of the reactions have been discussed. It has been suggested that the overall reactions catalyzed by guanidino kinases can be considered as involving a nucleophilic attack by the guanidino compound on the terminal phosphoryl group of ATP (88). But subsequently more detailed schemes have been proposed to explain in chemical terms the mechanism of the arginine kinase reaction (77, 82, 89) and these take into account the importance of various amino acid residues in the binding and catalytic processes. They envisage that following the formation of a central ternary complex, the terminal phosphoryl group of ATP is transferred initially to a histidyl residue a t the catalytic site of the enzyme and then to the guanidino acceptor. The schemes bear a close resemblance to those that have been postulated for the creatine kinase reaction (89).

VI. Equilibrium

In any discussion of the equilibria of the guanidino kinase reactions, it is important to bear in mind that the substrates for these enzymes undergo nonenzymic reactions with all the bivalent metal ions which can function as activators and that the stability constant of each metalsubstrate complex varies with the identity of the metal ion ( 6 5 ) .Therefore it follows that at fixed pH and ionic strength, the equilibrium position of the guanidino kinase reaction will depend not only on the metal ion present but also on its concentration. The effects are well illustrated by the more extensive equilibrium studies with creatine kinase (90) and 88. P. D. Boyer and W. H. Harrison, in “The Mechanism of Enzyme Action” (W. D. McElroy and B. Glass, eds.), p. 658. Johns Hopkins Press, Baltimore, Maryland, 1954. 89. A . S. Mildvan, “The Enzymes,” 3rd ed., Vol. 2, p. 445, 1970. 90. J. F. Morrison and A. White, Eur. J . Biochem. 3, 145 (1967).

486

J. F. MORRISON

to a lesser extent by those with arginine kinase in the presence of magnesium (20, 91). The latter investigations showed that the apparent equilibrium constant ( K ; ) which is defined as

K' = [phosphoarginine][ADP] [arginine][ATP] varies with the total concentration of magnesium and the experimental conditions. On the other hand, when the true equilibrium constant ( K e g ) is determined by substituting the calculated concentrations of free phosphoarginine, MgADP-, free arginine, and MgATP*- into the relationship 0,

[free phosphoarginine][MgADP] [free arginine][MgATP+] relatively constant values are obtained. Values of 0.31 (20) and 0.46 (91) have been reported for Keq, (It should be pointed out that in reference 20 the relationships given for K',, and K,, were inadvertently given as the inverse of those actually used for the calculations.) It can be expected that the K', value will approach that for K,, when the total concentration of metal is high relative to the total concentrations of the substrates (90). Under similar experimental conditions, the K,, values for the arginine kinase reaction is about ten times greater than that for the creatine kinase reaction (20, 21, 90). Thus, phosphoarginine is a less efficient compound for the storage of high energy than is phosphocreatine (91).

K,

=

91. W. W. Cleland, Annu. Rev. Biochem. 36, 77 (1967).

Glycerol and Glycerate Kinases JEREMY W. THORNER

HENRY PAULUS

I. Introduction . . . . . . . 11. Glycerol Kinases . . . A. Methods of Assay and Distribution B. Metabolic Role . . . C. Molecular Properties . . D. Catalytic Properties . . E. Regulation in Microorganisms F. Regulation in Mammals . . 111. n-Glycerate Enases . A. Methods of Assay and Distribution . B. Metabolic Role . . . C. Molecular Properties . . . D. Catalytic Properties . . .

.

.

. . .

.

. . . . . .

. . . . . .

. . . . . . . . . .

. . . . . . . . . .

. . . . . . . . . .

. . . . . . . . . .

. . . . . . . . . .

. . . . . . . . . .

. . . . . . . . . . . . . . . . . . .

487 488

488 492 493 497 502 504 504 504 505 506 607

1. Introduction

This chapter deals with the enzymes glycerol kinase and glycerate kinase, which catalyze the transfer of the terminal phosphate of A T P to glycerol and D-glycerate, respectively, according to Eqs. (1) and (2). ATP: glycerol phosphotransferase (EC 2.7.1.30) : Glycerol

+ ATP

--t

sn-glycerol 3-phosphate ( 1 )

+ ADP

(1)

ATP: D-glycerate 3-phosphotransferase (EC 2.7.1.31) : D-glycerate

+ ATP

--t

+

3-phospho-~-glycerate ADP

(2)

1. IUPAG-IUB Commission on Biochemical Nomenclature : The Nomenclature of Lipids, Tentative Rule 1.2, JBC 242, 4845 (1967). 487

488

JEREMY W. THORNER AND HENRY PAULUS

These enzymes have little in common in terms of their properties and function, but divergence of properties and function applies also to the glycerol kinases from different sources and even to the glycerate kinases existing in a single organism. The two types of enzyme will therefore be treated in separate sections and, in the case of glycerol kinase, the enzymes of microbial and mammalian origin must often be considered as quite separate entities. Emphasis will be placed on work published in the past 10 years during which considerable progress has been made in the study of the molecular properties and the regulation of these enzymes, particularly those of bacterial origin.

II. Glycerol Kinares

A. METHODS OF ASSAYAND DISTRIBUTION Since the discovery of the esterification of glycerol in kidney minces by Kalckar in 1937 ( d ) , glycerol kinase activity has been reported in a large variety of tissues and organisms (Table I) (3-24). Early studies 2. H. Kalckar, Eiizymologia 2, 47 (1937) ; BJ 33, 631 (1939). 3. P. Hahn and R. Greenberg, Experientia 24, 428 (1968). 4. S. Theil, Naturwissenschaften 53, 436 (1966). 5. R. G. Vernon and D. G. Walker, BJ 118, 531 (1970). 6. J. Robinson and E. A. Newsholme, BJ 112, 455 (1969). 7. S. C. Kampf, H. F. Seitz, and W. Tarnowski, Hoppe-Seyler’s Z . Physiol. Chem. 351, 32 (1970). 8. B. 1,. Knight and N. B. Myant, BJ 119, 103 (1970). 9. H. A. Haessler and K . J. Isselbacher, BBA 73, 427 (1963). 10. 0. Wieland and M. Suyter, Biochem. 2. 329, 320 (1957); C. Bublitz and 0. Wieland, “Methods in Enzymology,” Vol. 5, Chapter 46, 1962. 11. D. Trcble and E. G. Ball, Fed. Pioc., Fed. Amer. SOC. Exp. Biol. 22, 357 (1963). 12. C. Bublitz and E. P. Kennedy, JBC 211, 951 and 963 (1954). 13. T. Koschinsky, F. A. Gries, and L. Herberg, Diabetologia 7, 316 (1971). 14. J. Himms-Hagen, Can. J . Biochem. 46, 1107 (1968). 15. J. Lech, Comp. Biochem. Physiol. 34, 117 (1970). 16. B. Clark and G. Hubscher, Nature (London) 195, 599 (1962). 17. E. A. Newsholme and K . Taylor, BJ 112, 465 (1969). 18. 0. W. McBride and E. D. Korn, J . Lipid Res. 5, 442 (1964). 19. J. Robinson and E. A. Newsholme, BJ 104, 2C (1967). 20. D. Treble and J . Mayer, Nature (London) 200, 363 (1963). 21. R. F. Welton, R. J. Martin, Jr., and B. R. Baumgardt, Fed. Proc., Fed. Amer. Soc. Exp. Biol. 31, 698 (1972). 22. T. Koschinsky and F. A. Gries, Hoppe-Seyler’s Z . Physiol. Chem. 352, 430 (1971).

14.

489

Q L T C ~ O LAND OLTCERATE KINASES

TABLE I GLYCKROL KINAS&: ACTIVITYIN VARIOUSTISSUES Tissue Liver

Kidney

Intestine

Muscle

Mammary gland Brown adipose tissue

Organism Rat (newborn) Rat (newborn) Rat (newborn) Rat (adult female) Rat (adult female) Rat (adult male) Rat (adult male) Rat (adult) Rat (adult) Rat (adult) Rat (adult) ltat (adult) Mouse (normal) Mouse (obese) Hamster (male) Rabbit (newborn) Rabbit (adult male) Pigeon Domestic fowl Trout Rat (newborn) Rat (adult female) Rat (adult female) Rat (adult male) Rat (adult) Rat (newborn) Rat (adult female) Rat (adult male) ltat (adult) Hamster (adult male) Cat Rat (cardiac) Rat (diaphragm) Rat (thigh) Rabbit (semitendinosus) Pigeon (pectoral) Domestic fowl (pectoral) Trout (red or white) Locust (flight) Bumblebee (flight) Queen bumblebee (flight) Guinea pig Rat (newborn) Hat (adult female)

Ref. 3

4 6 6

6 6 7 8 9 10 11

12 1s 13 Y

8 14 10 10 16 6 6 6 6 10 6

6 6 9 9

16 17 17 17 17 17 17 17 17 17 17 18 6 6

Enzymic activity. Assay (temp.) 23 40 30 150 120 220 330 90 210 170 200 120 90 110 190 34 3 330 40 30 80 310 170 170 140 40 30 10 40 20 20 0.96 1.8 0.42 1.2 11

0.96 <0.04 31 140 410 10 10 20

I B (37") IA (25") 111 (30") I11 (30") 111 (26") I11 (30") I B (37") 11 (24") I B (37") IA (25") IA (25") I B (37") I11 (37") 111 (37") I B (37") 11 (24") I11 (25") IA (25") IA (25") IB (22") I11 (30") 111 (30") I11 (26") I11 (30") IA (25") 111 (30") I11 (30") I11 (30") I B (37") I B (37") IA (37") I11 (37") I11 (37") 111 (37") I11 (37") 111 (37") I11 (37") I11 (210) I11 (30") I11 (30") I11 (30") 111 (25") 111 (30") I11 (30") (Continued)

490

JEREMY W. THORNER AND HENRY PAULUS

TABLE I (Conlinued) Tissue

Organism

Ref.

Rat (adult male) Rat (adult) Rat (adult) Rabbit (newborn) Rat Mouse (normal) Mouse (normal) Mouse (obese) Mouse (obese) Domestic fowl Locust (fat body) Human (normal subcutaneous) Human (obese subcutaneous) Rabbit Domestic fowl Trout Human Bovine

6 8 11 8 1.9

~

Brown adipose tissue Whiteadipose tissue

Spermatozoa

1s 20 13 20 21

17 22 22

23 23 23 23 23

Enzymic activity. Assay (temp.) ~

9 40 140 4 0.24 0.15 0.46 1.9 3.8 3.2 14 0.01 0.03 0.3 24 0.1 0.0 8.1

I11 (30") I1 (24") IA (25") I1 (24") I11 (37") I11 (37") I11 (210) I11 (37") I11 (21") 111 (25") I11 (30") I11 (25") I11 (25") IA (37") IA (37") IA (37") IA (37") IA (37")

Maximum rate of glycerol 3-phosphate formation in micromoles per hour per gram fresh weight of tissue. Where the protein content of the tissue waa not given, the value for the corresponding rat tissue was used (24). 4

depended primarily on the enzymic estimation of reaction products. Thus, glycerol-3-P produced may be measured spectrophotometrically by the amount of NAD' reduced in the presence of glycerol-3-P dehydrogenase, either during the progress of the glycerol kinase reaction (Assay 1A;IO) or in a separate incubation after the glycerol kinase reaction has been terminated (Assay IB ;12),Assay IA permits a more convenient measurement of the rate of product formation, but it requires that the formation of glycerol-3-P be studied above pH 9. Alternatively, the progress of the reaction can be followed by coupling ADP formation to the oxidation of NADH in the presence of phosphoenolpyruvate, pyruvate kinase, and lactate dehydrogenase (Assay I1;25). Recently, several radiochemical assays have been devised which are often preferable to the spectrophotometric procedures because of their high sensitivity and greater ~~

23. H. Mohri and J. Masaki, J. Reprod. Fert. 14, 179 (1967); T. Mann, "Biochemistry of Semen and of the Male Reproductive Tract," Chapter 5. Methuen, London, 1964. 24. W. E. Knox, "Enzyme Patterns in Fetal, Adult, and Neoplastic Rat Tissues," p. 258. Karger, Basel, 1972. 25. P. B. Garland and P. 6 . Randle, Nature (London) 196, 987 (1962).

14.

GLYCEROL AND GLYCERATE KINASES

491

flexibility. The formation of glycerol-3-P from radioactive glycerol can be measured by chromatographic separation (18), by precipitation as the lead salt (26), or, more conveniently, by adsorption to discs of DEAE-filter paper (27) (Assay 111). Alternatively, glycerol-dependent conversion of [Y-~*P]ATP to an acid-stable phosphate ester can be measured (Assay IV ;28). These radiochemical methods have revealed the presence of glycerol kinase in many sources from which the enzyme had been believed to be absent. In most of the tissues described in Table I glycerol kinase is present in the soluble cytoplasmic fraction. I n some cases, however, the enzyme appears to be associated with particulate material: In homogenates of locust flight muscle, two-thirds of the glycerol kinase activity sediments a t 6000 X g (17); in bull spermatozoa, the enzyme appears associated with the mitochondria of the midpiece ($3); and in peanut seedlings, a mitochondria1 association of the enzyme is also observed (d9). Glycerol kinases have also been found in many microorganisms. Bacterial sources include Escherichia wli (SO), Shigella sonmi (31), Klebsiella aerogenes (32), Pseudomonas aeruginosa (33), Halobacterium cutirubrum (34), Acetobacter suboxydam (35), Bacillus subtilis (36), Clostridium m v y i (37), Staphylococcus aureus (38), Streptococcus faecalis (39), Nocardia asteroides (40), Mycobacterium 607 (41), M . smegmatis (@), M . butyricum ( 4 3 ) , M . tuberculosis ( 4 4 ) , and Myco26. 5.Hayashi and E. C. C. Lin, BBA 94, 479 (1965). 27. E. A. Newsholme, J. Robinson, and K. Taylor, BBA 132, 338 (1967). 28. J. W. Thorner and H. Paulus, JBC 246, 3885 (1971). 29. P. F. Stumpf, Plant Physiol. 30, 55 (1955). 30. J. P. Koch, 5.Hayashi, and E. C. C. Lin, JBC 239, 3106 (1964). 31. D. P. Richey and E. C. C. Lin, J. Bncteriol. 112, 784 (1972). 32. D. Rush, D. Karibian, M. L. Karnovsky, and B. Magasanik, JBC 226, 891 (1957); E. C. C. Lin, A. P. Levin, and B. Magasanik, &id. 235, 1824 (1960). 33. 5. 5. Tsay, Ph.D. Dissertation, Oklahoma State University, Stillwater, 1971. 34. M. K. Wassef, J. Sarner, and M. Kates, Can. J. Biochem. 4 4 63 and 69 (1970). 35. J . F. Hauge, T. E. King, and V. H. Cheldelin, JBC 214, 1 (1955). 36. L. Mindich, J. Bacteriol. 98, 565 (1968). 37. G. F. Shemanova and V. A. Blagoveshchenskyi, Biokhimiya 21, 729 (1956); C A 51, 9740 (1957). 38. D. P. Richey and E. C. C. Lin, J. Bacteriol. (1973) (in press). 39. N. J. Jacobs and P. J . Vandemark, J. Bacteriol. 79, 532 (1960). 40. R. B. Clames, Jr. and 5.J . Deal, Bacteriol. Pr.oc. 147 (1970). 41. S. V. Pande, R. Parvin, and T. A. Venkitasubramanian, Can. J. Biochem. 45, 797 (1967). 42. W. Segal and 5.Lahiri, BacterioZ. Proc. 161 (1972). 43. G. J. E. Hunter, BJ 55, 320 (1953). 44. F. G. Winder and P. J . Brennan, J. Bncteriol. 92, 1846 (1966).

492

JEREMY W. THORNER AND HENRY PAULUS

plasma naycoides ( 4 5 ) . The occurrence in fungi has been reported for C‘andida mycoderma ( 4 6 ) ,C. utilis (47), Oospora lactis ( 4 8 ) ,Hawenula anomnla (49), Saccharomyces cerevisia (10), and Neurospora crassa ( 5 0 ) .

B. METABOLIC ROLE Glycerol-3-P has diverse metabolic fates. It often is an intermediate in the catabolism of glycerol; it serves as precursor for triglycerides and complex lipids as well as for macromolecules such as teichoic acids; and it participates in a cycle of reactions by which electrons can be transferred from cytoplasm into mitochondria. Accordingly, the function of glycerol kinase may be quite different in various organisms and tissues. In microorganisms, glycerol kinase functions primarily in the utilization of glycerol as a carbon and energy source by phosphorylation and subsequent oxidation by a flavin-linked glycerol-3-P dehydrogenase (EC 1.1.99.5).This is supported by the observation that the synthesis of glycerol kinase and glycerol-3-P dehydrogenase are specifically induced during aerobic growth on glycerol (30, 32, 33, 39, 40, 42, 44, 465 0 ) . I n E . coli it has been shown that glycerol kinase, two flavin-linked glycerol-3-P dehydrogenases, a system facilitating glycerol entry, and an active transport system for glycerol-3-P are under the control of a single regulatory gene ( 5 1 ) .The synthesis of the enzymes that constitute this regulon is induced by glycerol-3-P and is subject to catabolite repression (51, 5 2 ) , further evidence for a catabolic role of glycerol kinase. Some bacterial species, however, such as Bacillus subtilis (53) and others (32, 35, 39, 54-56), can catabolize glycerol by an alternate 45. P. Plackett and A. W. Rodwell, BBA 210, 221 (1970). 46. H. U. Bergmeyer, G. Holz, E. M. Kauder, H. Mollering, and 0. Wieland, Biochem. 2. 333, 471 (1961). 47. C. Gancedo, J. M. Gancedo, and A. Sols, Eur. J . Biochem. 5, 165 (1968). 48. E. M. Kauder, Ph.D. Dissertation, University of Munich, 1960. 49. K. Otsuka and H. Masuda, Nippon Nogei Kagnku Kakhi 30, 166 (1956); C A 52, 3915 (1958). 50. J. Courtright, Bactem’ol. Proc. 153 (1971). 51. N. A. Cozzarelli, W. B. Freedberg, and E. C. C. Lin, J M B 31, 371 (1968): Y. Sanno, T. H. Wilson, and E. C. C. Lin, BBRC 32, 344 (1968); W S. Kistler and E. C. C. Lin, J. Bacteriol. 108, 1224 (1971). 52. N. Zwaig and E. C. C. Lin, BBRC 22, 414 (1966). B. DeCrombrugghe, R. L. Pcrlman, H. E. Varmus, and I. Pastan, JBC 244, 5828 (1969). 53. J. M. Wiame, S. Bourgeois, and R. Lambion, Nature (London) 174, 37 (1954). 54. I. C. Giinsalus and W. W. Umbreit, J. Bacteriol. 49, 347 (1945). 55. R. E. Asnis and A. F. Brodie, JBC 203, 153 (1953). 56. T. Ramakrishnan, P. Suryanarayana-Murthy, and K. P. Gopinathan, Bacteliol. Rev. 36, 65 (1972).

14.

GLYCEROL A N D GLYCERATE KINASES

493

route that involves direct oxidation to dihydroxyacetone and subsequent phosphorylation. Indeed, in B. subtilis the level of glycerol kinase is very low and is not induced by growth on glycerol ( 3 6 ) . In higher organisms the primary role of glycerol kinase seems to be the salvage of glycerol released upon lipolysis. This is well illustrated in mammalian intestine (9, 16) and the brown adipose tissue of rat (11) where glycerol produced by lipolysis is rephosphorylated by relatively high levels of glycerol kinase and reused for lipid synthesis. On the other hand, glycerol kinase levels in white adipose tissue are very low (Table I) ; presumably glycerol released in this tissue, as well as that produced in the capillaries by lipoprotein lipase, is reesterified in the liver and kidney which have a high content of glycerol kinase. It is of interest that tissues of genetically obese animals have abnormally high levels of the enzyme (13, 20, $ 2 ) . The content of glycerol kinase in normal animals is relatively invariant under a variety of nutritional ( 6 ) and hormonal conditions, and shows no correlation with plasma or tissue glycerol levels ( 7 ) . In tissues where reducing power is not extensively used for biosynthesis, cytoplasmic dihydroxyacetone-P is reduced by a NAD-linked glycerol-3-P dehydrogenase (EC 1.1.1.8) to glycerol-3-P, which can enter the mitochondria1 intermembrane space to be reoxidized by a flavin-linked enzyme ( 5 7 ) . The net result of this cycle is a transfer of electrons from the cytoplasm into the mitochondria but, although glycerol-3-P participates catalytically, it will occasionally have to be replenished. Insect flight muscle contains high levels of the enzymes of the “glycerol-3-P cycle” ( 5 8 ) ,and the large amount of glycerol kinase in this tissue (17) may serve primarily such an anaplerotic function.

PROPERTIES C. MOLECULAR 1 . Purification. and State of Purity

Crystalline glycerol kinase has been obtained from pigeon liver ( l o ) , Candida mycodernaa ( 4 6 ) , and Escherichia coli (69) by procedures based on the methods developed originally for the partial purification of the enzyme from rat liver (12). An important aspect of these procedures, which allowed the selective removal of many other proteins, is 57. E. Racker, “Mechanisms in Bioenergetics,” p. 100. Academic Press, New York, 1965; P. D. Holohan, C. A. Lepp, and T. P. Fondy, Fed. Proc., Fed. Amer. SOC. Ezp. B i d . 31, 421 (1972). 58. R. W. Estabrook and B. Sacktor, JBC 233, 1014 (1958). 59. S. Hayashi and E. C. C. Lin, JBC 242, 1030 (1967).

494

JEREMY W. THORNER AND HENRY PAULUS

the protection of glycerol kinase by glycerol, EDTA, and sulfhydryl compounds at elevated temperatures and during dialysis against buffers of low ionic strength and low pH. Nonetheless, these harsh treatments could produce alterations in the properties of the enzyme, and a more gentle procedure has therefore been devised for the purification of the enzyme from E . coli (28). A crystalline preparation was obtained in a yield of SO%, and electrophoresis in polyacrylamide gels and equilibrium ultracentrifugation have demonstrated its homogeneity. Partial purification of glycerol kinase has been achieved from a variety of sources such as liver of rat (6, 9, 12, 601,ox ( 6 0 ) , man (60), and hamster ( 9 ) , rat and hamster intestine ( 9 ) , Oospora Zactis (@), and Neurospora crassa (61). These preparations contain various other enzymic activities such as adenylate kinase, triose phosphate isomerase, D-triokinase, and phosphatases that might interfere with studies of these enzymes. 2. Composition

The amino acid composition has been reported only for the glycerol kinase of E. coli (28).The enzyme contains a net excess of acidic groups. The content of tryptophan, tyrosine, and cysteine is 43, 69, and 20 residues per mole of enzyme, respectively. All cysteine residues are present in the reduced form. Glutamate is the only carboxyl-terminal amino acid. The low isoionic point (PI = 4.6) of the glycerol kinase from yeast suggests that this enzyme also contains a net excess of acidic residues ( 4 6 ) . A high content of tryptophan was inferred from its ultraviolet spectrum which is very similar to that of the bacterial enzyme (28) in that it exhibits a distinct shoulder a t 290 nm. 3. Size and Subunit Structure

The partial specific volumc of the glycerol kinase from E . coli was found to be 0.732 ml/gm from the amino acid composition, and 0.724 ml/gm by differential ultracentrifugation in buffers of D20 and H,O (28). With the use of these values the molecular weight was determined by equilibrium sedimentation to be 217,000 or 210,000, respectively. [An earlier estimate of about 300,000 (59) was probably too high because of aggregate formation.] Equilibrium ultracentrifugation in the presence of 6 M guanidine hydrochloride revealed a single molecular species of molecular weight 55,000. Similarly, a single band of molecular weight 60. N. Grunnet and F. Lundquist, Eur. J . Biochem. 3, 78 (1967). 61. J. Courtright, personal communication (1972).

14.

GLYCEROL AND GLTCERATE KINASES

495

57,000 was observed upon polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate. The results suggest that glycerol kinase is a tetrameric protein composed of four similar or identical subunits, a conclusion confirmed by studies with the bifunctional cross-linking reagent, dimethylsuberimidate. Evidence that the four subunits of glycerol kinase represent a single type of polypeptide chain has come from the examination of complete tryptic digests of the carboxymethylated protein. The two-dimensional peptide map revealed 48 ninhydrinpositive spots of which 15 contained tyrosine and 10 contained tryptophan, in close agreement with the numbers predicted from the amino acid composition for a tetramer composed of four identical subunits. The recovery after hydrazinolysis of 4 moles of glutamate per mole of enzyme as the only amino acid is also consistent with this conclusion. The molecular weight of yeast glycerol kinase was calculated to be 251,000 from its sedimentation coefficient ( s & , ~= 10.87 S) and its diffusion coefficient (d,",,, = 4.2 X 10-' cmz sec-l) and an assumed partial specific volume (46).Electrophoresis of the yeast enzyme in polyacrylamide gels containing sodium dodecyl sulfate revealed a single species of molecular weight about 60,000, suggesting that this enzyme is also composed of four similar or identical subunits (62). I n contrast, preliminary studies by gel filtration and zone sedimentation in sucrose gradients with the glycerol kinase from Neurospora crassa suggest a molecular weight of only 140,000 (61). Staining of polyacrylamide gels of crude and partially purified preparations for glycerol kinase activity indicated the presence of only one form of the enzyme. 4. Stability

I n the presence of 10 mill glycerol and 1 m M EDTA, almost all the glycerol kinases can be heated for prolonged periods a t 60" with little loss of activity, a property useful for the purification of the enzymes. Exceptions are the enzymes from Neurospora crassa (61),which loses 30% of its activity in 5 min a t 50°, and from Mycobacterium 607 (4l), which is quite heat labile. At 70" and in the presence of glycerol, the glycerol kinases from rat liver (12) and from E. coli (62) have halflives of about 20 and 5 min, respectively. The yeast enzyme is remarkable in that it can be boiled for 4 min in 0.1 N HCl without precipitation or change in its ultraviolet absorption spectrum (46). The glycerol kinases from higher organisms are most stable in an acidic pH range (10,12). I n the absence of glycerol, the enzyme from 62. J. Thorner, Ph.D. Dissertation, Harvard University, Cambridge, Massachusetts, 1972.

496

JEREMY W. THORNER AND HENRY PAULUS

pigeon liver (10) loses about 90% of its activity when kept a t pH 7.0 and 0" for 6 hr, while it is completely stable at pH 5.0. Glycerol kinase from rat liver (19) has a half-life of 6.5 min a t p H 9.8 and 25", while no loss of activity is observed for several hours a t pH 7.5. On the other hand, the glycerol kinases from microorganisms are most stable a t a neutral pH. I n the absence of glycerol, the enzyme from E. coli (59)is totally inactivated when kept for 24 hr at 0" and pH 5.0, while at pH 7.0 only 50% of the activity is lost. The glycerol kinase from Mycobacterium 607 (41), in the presence of glycerol, is completely stable for 30 minutes a t 30" from pH 6.0 to 9.0, while a t pH 5.0 it loses about 75% of its activity. In every case described glycerol affords considerable stabilization a t the unfavorable pH values. The pigeon liver (10) and E. coli (59,62) glycerol kinases are insoluble in pure water or in buffers of low ionic strength. The pigeon liver enzyme is also sensitive to extreme dilution but can be stabilized by the addition of 0.01% bovine serum albumin. In crude extracts, the glycerol kinases of rat liver (12) and Pseudomonus aeruginosa (63) are stable to freezing and thawing, while more purified preparations are inactivated. The partially purified enzymes from pigeon liver (lo), rat liver (9,la), hamster liver (9), and rat and hamster intestine (9) have half-lives of several weeks to months at -20" and pH 5.0. The yeast enzyme (46) has been stored for several months and the E. coli enzyme (59,62) for several years without loss of activity RS crystallinc suspensions in saturated ammonium sulfate solutions containing 10 mM glycerol, 1 mM EDTA, and a thiol such as 2-mercaptoethanol. Glycerol kinases are susceptible to inactivation by the oxidation of sulfhydryl groups. The enzyme from pigeon liver which has lost activity upon storage can be reactivated by incubation with 20 mM cysteine or glutathione (10). The rat liver enzyme is inactivated by shaking in air and exhibits maximum activity when assayed in the presence of thiols (12). 5. Chemical Modification Since sulfhydryl groups appear to be essential for the activity of most glycerol kinases, these have been the major target for chemical modification. The rat liver enzyme is completely inactivated by treatment with 5 mM iodoacetamide at pH 7.4 for 1 hr, while an equivalent concentration of p-chloromercuribenzoate causes only an 80% loss of activity (12). In contrast, the yeast (46) and bacterial (66) enzymes are much more sensitive to mercurials than to iodoacetamide or other sulfhydryl re63. S. S. Tsag, K . K. Brown, and E. T. Gaudy,

J. Bacterial. 108,

82 (1971).

14.

497

GLYCEROL AND GLYCERATE KINASES

a),

agents. For example, p-hydroxymercuriphenylsulfonate (50 iodo, and N-ethylmaleimide (5 mM) acetate (5 mM) , iodoacetamide (5 d ) produce 100, 90, 5, and 25% inactivation, respectively, of the E. coli enzyme in 15 min at 0" (62). Not all sulfhydryl groups of the E. coli glycerol kinase are exposed to reaction with 5,5'-dithiobis(nitro-2-benzoic acid). Only 12 of the 20 thiols can be titrated with this reagent in the native enzyme (,??8), while in the presence of glycerol only two sulfhydryl groups are exposed (&%'a), suggesting that glycerol promotes a major conformational change in the enzyme.

D. CATALYTIC PROPERTIES 1. Substrate Specificity and Kinetics Glycerol kinase catalyzes the stereospecific transfer of the terminal phosphoryl moiety of ATP to one of the primary hydroxyl groups of glycerol, forming sn-glycerol 3-phosphate ( I d ) . This is a classic example of an enzyme that can distinguish between two chemically identical functional groups. The comparison of other aspects of the specificity of glycerol kinases from various sources is often difficult as a result of the variety of assay systems and conditions used in different laboratories.

a. Phosphoryl Group Acceptors. Highly purified preparations of glycerol kinase can also catalyze the phosphorylation of dihydroxyacetone TABLE I1 PHOSPHORYL GROUPACCEPTOR SPECIFICITY OF SOME GLYCEROL KINASES

Km

Relative V,,

Glycerol DHA LGA D-GA Enzymesource Eschmichia wlib Candidumycodermac Pigeon liverd Rat liver1 a

Glycerol DHA4 L-GA~ D-GA

[loo]

[loo] [loo] [loo]

187 10 138 200

81 10

-' 150

(35) (10)

-

(a) (pM) 10 35 30 10

(pM)

500 3000 5000 2000 600 -

(pM) 500 -

Abbreviations; ])HA, dihydroxyacetone; GA, glyceraldehyde.

* All values obtained by Assay I1 (69) except.K m for glycerol determined by Assay I11 (68).

All values obtained by Assay I1 (47, 60). All values obtained by Assay IA (10). Not determined. f All values obtained by a manometric assay (18)except K, for glycerol determined by Assay I1 (60).

498

JEREMY W. THORNER AND HENRY PAULUS

and L-glyceraldehyde (Table 11). The affinity of the enzyme for these compounds is very much less than that for glycerol; on the other hand, the phosphorylation of dihydroxyacetone is often more rapid. D-G~Yceraldehyde effects a greater than stoichiometric release of ADP from ATP, but orthophosphate rather than ~-glyceraldehyde-3-Pis formed as the other product of the reaction (69). Presumably, the hydrated form of this triose is phosphorylated in position 1 to yield an unstable intermediate that decomposes to D-glyceraldehyde and orthophosphate. The structures of the active substrates of glycerol kinase are illustrated in Fig. 1. No other compounds have been found to serve as phosphoryl acceptors. For the enzymes from E . coZi, yeast and mammalian liver, the apparent K , for glycerol is independent of ATP concentration. Substrate saturation curves for glycerol are hyperbolic, yielding linear double reciprocal plots (60, 6 2 ) . b. Phosphoryl Group Donors. The specificity of various glycerol kinases for phosphoryl group donors is summarized in Table 111. The enzyme from E . coli which can utilize only ATP exhibits the highest degree of specificity (69). A crude preparation of glycerol kinase from brown fat has been reported to utilize C T P even better than ATP ( I I ) , and enzyme preparations from rat heart and white fat are equally active with UTP and ATP (19). The apparent K , for ATP of the glycerol kinases from E . coZi ( 6 2 ) , yeast (GO),and liver of rat (6, 60),ox (60), and man (60) depends on the concentration of both glycerol and Mg2+.Even in the presence of an excess of Mg2+and at saturating glycerol, the response of reaction velocity to ATP concentration is not hyperbolic, and double reciprocal plots have a downward curvature (Fig. 2). This effect is especially pronounced at pH 7.0 and is reduced for some of the enzymes, hut not eliminated, at pH 9.0 (60). Such a “biphasic” nature of double reciprocal plots would be consistent with two types of sites that differ in their affinity for ATP and may account for the discrepancy between the value of

CKOH I HO -C -H I H-C-H I OH Glycerol

CH,OH I

o=c

I

H-C-H I OH Dihydroxyacetone

CHO I HO-C-H I H-C-H I OH L-Glyceraldehyde

C%OH I HO-C-H I

H-C-OH I OH D

-Glyceraldehyde hydrate

FIG.1. Structure of phosphoryl group acceptors utilized by glycerol kinase.

TABLE I11 PHOSPHORYL GROUPDONORSPECIFICITY OF SOMEGLYCEROL KINASES ~~~~

~

K,,, for MgATPa

K, for MgATPb

Relative Vmax pH 9.0 Enzyme source

ATP

UTP

ITP

Eseherichia eoli. Candida mywdemtad Pigeon liver, Rat livep

[loo] [loo] [loo] [loo]

0 14

0 55

-

70

0

GTP

CTP 0

0

22 -

TTP 0

-0

-

-

-

70

-

(do

pH 7.0

(PM)

100;500 80;300 55;200 90 2 8 35;70 1O;lOO

pH 7.0 GuM) -

160

-

170

Determined in the presence of excess MgZ+ at saturating glycerol concentration. Determined by replotting data, obtained by Assay 11, extrapolated to zero concentrations of both free Mg"+ and free ATP (60). c All values obtained by Assay IA (69) except KmaPp values for MgATP determined by Assay IV (66). d All values obtained by b a y IA (46) except K,,, for MgATP a t pH 7.0 determined by Assay I1 (60). e Not determined. Value obtained by Assay IA (10). 0 Values obtained by Assay IB (16) except K,, for MgATP at pH 9.0 determined by Assay I A (60)and K,,, for MgATP a t pH 7.0 determined by Assay I1 (60). a

500

JEREMY W. THORNER AND HENRY PAULUS

2

4

6 (MgATP, mM)-'

8

10

FIQ.2. Double reciprocal plot of effect of MgATP concentration on glycerol kinase activity. Crystalline glycerol kinase from E . coli was assayed at different ATP concentrations with a 1 mM excess of Mg*+at pH 7.0 by Assay IV [from Thorner (68)1.

the K,,, reported earlier for the E. coli enzyme (69) and that presently observed ( 6 2 ) .On the other hand, such kinetic behavior could also result from the presence of two different enzymes; this possibility is unlikely, since the unusual ATP kinetics are unchanged in the course of enzyme purification (62) and are characteristic for the glycerol kinases from many different sources. For the enzymes from yeast and from rat and ox liver, it has been shown that double reciprocal plots with respect to the MgATP complex are linear when the reaction velocities are extrapolated to zero concentration of both free ATP and free Mg2+ ( 6 0 ) . These results have been interpreted in terms of activation by free ATP; however, since free ATP and free l\lg2' are not independent variables, these observations would also be consistent with an inhibitory role of free Mgz+.

c. Divalent Metal Ions. Glycerol kinase from rat liver, even after treatment with EDTA, is stimulated only about twofold by the addition of Mgz+ (6, 1 2 ) . The optimum concentration of Mg2+ is 2-3 m M and does not depend on the concentration of ATP. Higher concentrations of Mg2+are inhibitory. In contrast, the glycerol kinases from E. co2.i (62), Mycobacterium

14. GLYCEROL AND

GLYCERATE KINASES

501

607 (41), yeast (46), and pigeon liver (10) have a n absolute requirement for added MgZ+,with the optimum depending on the concentration of ATP. At 1 mM ATP, the enzymes from E . w l i and pigeon liver exhibit maximum activity a t 2 mM Mg2+, a concentration just sufficient to convert all ATP to the MgATP complex (64). Higher concentration of Mgz+is only slightly inhibitory. With the rat (12) and pigeon liver (10) enzymes, Mn2+can fully substitute for Mg2+ a t equivalent concentrations, while with the E . coli enzyme (59) the maximum rate of the reaction with MnZ+is only 30% of that with Mgz+.The glycerol kinases of Clostridium nov.yi (37) and Mywbacterium 607 (41) are remarkably unspecific in that they can use Mn2+,CoZ+,and NiZ+as well as Mg2+. I n no case has Ca?+been found to be capable of substituting for Mgz+. In fact, in the absence of Mg2+the rat liver enzyme is inhibited by Ca2+ ( 1 2 ) . It can be restored to full activity by the addition of one-tenth the concentration of Mg2+(6).

d. Ionic Conditions. At pH 7.0, the activity of the glycerol kinase of E. coli is stimulated 30% by 0.4M KC1, while LiC1, NaCl, and NH4C1 have no effect ( 6 2 ) . I n contrast, the enzyme from Mycobacterium 607 is stimulated 25% by 0.15M NH4C1, whereas KC1 has no effect, and LiCl is strongly inhibitory (41). An extreme case, of course, is the glycerol kinase of Halobacterium cutirubrum which is active in 4 M NaCl (34). The pH optimum for almost all glycerol kinases studied lies in the range of 9.0-9.8. At pH 7.0, about half-maximal activity is observed. Exceptions are the enzymes from Neurospora crassa (61) and castor bean (65) with optimal activities between pH 8.0 and 8.4,and those of Clostridium novyi (37) and Mycobacterium 607 (41) with a p H optimum of 7.0-7.5. 2. Product Znhibition

The effect of ADP on rat liver glycerol kinase is rather complex (6).

At low concentrations of ATP, ADP inhibits noncompetitively with respect to that substrate, while a t high ATP the inhibition is of a partially competitive type. The degree of inhibition by ADP is increased by raising the Mg2+ concentration, but since Mg2+ a t high concentrations is itself inhibitory, this effect is difficult to interpret. With respect to glycerol, ADP is clearly an uncompetitive inhibitor. 64. W. W. Cleland, Annu. Rev. Biochem. 36, 77 (1967). 65. M. Yamada, Sci. Pap. Coll. Gen. Educ., Univ. Tokyo 10, 283 (1960); C A 55, 15636 (1961).

502

.JEREMY W. THORNER AND HENRY PAULUS

Glycerol-3-P is a competitive inhibitor with respect to glycerol ( K i = 0.6 mlll) and displays a mixed competitive-noncompetitive type of inhibition with respect to ATP (6). Insofar as the complexity of the ATP kinetics permit an interpretation, these results and the substrate kinetics are most consistent with an ordered mechanism for the enzyme from rat liver, with glycerol as the first substrate to bind to the enzyme. The product inhibition pattern of the glycerol kinase from E . coli suggests a similar ordered mechanism (6% * 3. Thermodynamics

The reaction catalyzed by glycerol kinase is essentially irreversible with AGO’ = - 5.1 kcal/mole, corresponding to an equilibrium constant of greater than lo3 (66). Several of the thermodynamic parameters have been estimated for the reaction catalyzed by the E . coli glycerol kinase ( 6 2 ) .Binding experiments show that the dissociation constant of the enzyme-glycerol complex is 10-5M, in good agreement with the K , for glycerol. Therefore, the free energy of glycerol binding is about -6.8 kcal/mole. The K , for glycerol is found to increase with decreasing temperature, indicating a positive enthalpy of binding (+ + 6 kcal/mole) . Consequently, the entropy of binding must also have a positive value ( 4 +45 cal mole-’ deg-’). Such a large positive entropy of substrate binding is unusual and suggests that the binding of glycerol to the enzyme is accompanied by a considerable conformational change. This conclusion is also supported by the masking of 10 additional sulfhydryl groups per enzyme molecule in the presence of glycerol and the marked stabilization of the enzyme by this substrate. The turnover number of the glycerol kinase of E. coli a t 25” is 11,600 min-’, corresponding to a free energy of activation of the enzymesubstrate complex of 14 kcal/mole (62). Its variation with temperature, when analyzed by an Arrhenius plot, is consistent with an activation energy of 5 kcal/mole. The turnover numbers a t 25” for the enzymes from pigeon liver (10) and from yeast (46) are 4,100 min-’ and 25,100 min-’, respectively.

E. REGULATION IN MICROORGANISMS The glycerol kinases of E . coli (67) and several other bacterial species (31, 38, 62) are specifically inhibited by fructose 1,6-diphosphate. Other 66. A. L. Lehninger, “Biochemistry,” p. 304. Worth, ,hew York, 1970.

14.

GLYCEROL AND GLYCERATE KINASES

503

glycolytic intermediates, members of the citric acid cycle and related compounds, have no effect on the activity of the enzyme (62, 67). The inhibition by fructose 1,g-diphosphate is noncompetitive with respect to both substrates with a Ki of about 1 mM a t pH 7 (62,67). Under conditions of optimal activity, such as in the presence of 0.4M KCl or a t pH 9.5, the enzyme is much less sensitive to inhibition, and several mutant strains of E . coli have been isolated whose glycerol kinase is unaffected by fructose 1,6-diphosphate under all conditions (67, 68). Its noncompetitive nature and its dissociation from catalytic activity under certain conditions and by mutation suggest that the inhibition of glycerol kinase by fructose 1,6-diphosphate proceeds by an allosteric mechanism. I t is of interest that a t least one of the mutant enzymes as well as the normal enzyme a t pH 9.5 have an unaltered molecular weight and can still bind fructose 1,6-diphosphate (62). Evidence that the inhibition of glycerol kinase by fructose 1,6-diphosphate operates under physiological conditions comes from the faster growth rates on glycerol of mutants with a “desensitized” enzyme (68, 69) and from the behavior of a strain that has a temperature-sensitive fructose 1,6-diphoephate aldolase (YO). The growth of the latter on glycerol is immediately inhibited by the addition of small amounts of glucose a t temperatures a t which the aldolase is inactive, presumably as a result of the inhibition of glycerol kinase by accumulated fructose 1,6-diphosphate derived from the added glucose. Normally, this control mechanism would operate to prevent the utilization of glycerol in the presence of a substrate that is a more efficient source of glycolytic intermediates. Crystalline glycerol kinase from Candida mycoderma is not inhibited hy 3 mM fructose 1,6-diphosphate (60). Assuming that the heat treatment during the purification procedure has not desensitized the enzyme to allosteric effects, this suggests a mode of regulation different from that of the bacterial enzyme. Indeed, the yeast enzyme is subject to uncompetitive inhibition (with respect to ATP) by AMP with a Ki of 0.2 mM (60).The response of enzymic activity to varying concentrations of AMP is hyperbolic which may imply that allosteric effects are not involved. The physiological significance as well as the mechanism of the AMP effect is obscure and difficult to rationalize in terms of a catabolic function of the enzyme. 67. 68. 69. 70.

N. Zwaig and E. C. C. I h , Science 153, 755 (1966). M. Berman and E. C. C. Lin, J. B a c t e k l . 105, 113 (1971). N. Zaaip, W. S. Kistler, and E. C. C . Lin, J. Bacteriol. 102, 753 (1970). A. Bock and F. C. Neidhardt, J . Bacteriol. 92, 464 and 470 (1966).

504

JEREMY W. THORNER AND HENRY PAULUS

F. REGULATION IN MAMMALS Like the yeast enzyme, partially purified preparations of glycerol kinase from rat, human, and beef liver are unaffected by fructose 1,6diphosphate but are inhibited in an uncompetitive manner by AMP with K i values of about 0.5 mM ( 6 0 ) . Again, such an effect would be unusual with a catabolic enzyme, but it would be consistent with a biosynthetic function ( 7 1 ) . Such a function is also supported by the inhibition of the enzyme in rat liver homogenates by glycerol-3-P (6, 7 2 ) . This may simply be product inhibition ; however, the observation that in partially purified preparations of the enzyme, but not in crude homogenates, the inhibition by glycerol-3-P is completely eliminated by low concentrations of phosphate or sulfate ion (72) suggests an allosteric mechanism. Nevertheless, the physiological significance of this effect is questionable for the inhibition by glycerol-3-P is competitive with glycerol with a K iof about 0.5 mM (6, 7 2 ) , while the intracellular concentration of glycerol never falls below 50 p M ( 7 3 ) , an order of magnitude above the K , for glycerol kinase.

111. o-Glycerate Kinases

A. METHODSOF ASSAYAND DISTRIBUTION D-Glycerate kinase was first identified in yeast (74) and mammalian liver ( 7 5 ) ,and the properties of the enzyme from higher organisms have been reviewed in the second edition of “The Enzymes” ( 7 6 ) . Since that time studies have been confined to the glycerate kinases of bacteria. The enzyme was detected in the Crookes strain of Escherichia coli by means of a titrimetric assay procedure (77, 7 8 ) . Subsequent work with E. coli K12 utilized a more versatile assay system which measures the D-glycerate-dependent formation of ADP by observing the reduction of NADH in the presence of excess phosphoenolpyruvate, pyruvate kinase, 71. 72. 73. 74. 75. 76. 77. 78.

D. Atkinson, “The Enzymes,” 3rd ed., Vol. 1, p. 461, 1970. N. Grunnet, BJ 119, 927 (1970). A. L. Greenbaum, K . A. Gumaa, and P. McLean, ABB 143, 617 (1971). S. Black and N. G. Wright, JBC 221, 171 (1956). A. Ichihara and D. M. Greenberg, JBC 224, 331 (1957); 225, 949 (1957) H. G. Hers, “The Enzymes,” 2nd ed., Vol. 6, p. 75, 1962. R. W. Hansen and J. A. Hayashi, J . Bacteriol. 83, 679 (1962). C. C. Doughty, J. A. Hayashi, and H . I,. Gunther, JBC 241, 568 (1966).

14.

GLYCEROL A N D GLYCERATE KINABES

505

and lactic dehydrogenase ( 7 9 ) . Alternatively, the pyruvate produced in this system in the absence of lactic dehydrogenase can be measured as the phenylhydrazone, which avoids interference by NADH oxidases present in crude bacterial extracts ( 7 9 ) .

B. METABOLIC ROLE In bacteria, glycerate kinase serves two quite different functions which depend on the nature of the substrate for growth. With D-glycerate as the carbon source, glycerate kinase plays an essential role in catabolism by converting the substrate to the glycolytic intermediate, 3-phosphoglycerate. This applies also to growth on D-glucarate which is metabolized to pyruvate and D-glycerate (80). The specific induction of a glycerate kinase in E. coli by growth on these substrates supports such a catabolic role (78, 7 9 ) . During growth on glyoxylate or on substrates that are readily converted to glyoxylate, such as glycolate, a quite different situation prevails. The catabolism of these compounds proceeds through the series of reactions known as the dicarboxylic acid cycle, illustrated in Fig. 3 (81). Since the intermediates of the cycle must also serve as precursors for biosynthesis, the operation of this catabolic scheme requires the continuous replenishment of intermediates through so-called anaplerotic reaction sequences (81). In the presence of amino acids, the preferred anaplerotic reactions are those that produce pyruvate, oxalacetate, and acetyl-CoA, but in the absence of additional carbon sources, glyoxylate itself must providc biosynthetic precursors. This is achieved through thc glycerate pathway (Fig. 3) which effects the net conversion of glyoxylate to phosphoenolpyruvate in four steps of which one is the phosphorylation of D-glycerate (81). Therefore, glycerate kinase also has an anaplerotic function, and this accounts for the observation that in E. coli K12 a glycerate kinase is induced by growth on glycolate, but not by growth on glycolate plus Casamino acids ( 7 9 ) . The dual role of glycerate kinase is supported by the discovery of two distinct enzymes in strains of E. coli K12 ( 7 9 ) .Glycerate kinase I, the catabolic enzyme, is present during growth on glycerate or D-glucarate, and its formation is not affected by the presence of Casamino acids, while glycerate kinase 11, the anaplerotic enzyme, is found only during growth on glycolate in the absence of Casamino acids. Glycerate 79. M. K. Ornston and L. N. Ornston, J . Bacterial. 97, 1227 (1969). 80. H.J. Blumenthal and D. C. Fish, BBRC 11, 239 (1963).

81. H.1,. Kornberg, Essays Biochem. 2, 1 (1966).

506

JEREMY W. THORNER AND HENRY PAULUS

I

Glyoxylate Glyoxylate

I-

semi$dehyde Tartronic

\i

I Glyoxylate 1

J

2H

Glycerate

/ I 2Hh

) Malab 4

Phosph,oglycerate

-

COA SH

J

Oxalalcetate

I

i_/ ATRE/

m-

Phosphopyruvate

/

I

Cell components

FIG.3. Routes for the provision of energy and of cell components during microbial growth on glyoxylate. The catabolic route (dicarboxylic acid cycle) is shown by light arrows; the anaplerotic sequence (glycerrite pathway) is shown by heavy arrows [from Kornberg (81) 1, kinase I is also formed-to varying extents in different strains of E . co2i K l a - d u r i n g growth on glycolate, perhaps as a result of internal induction by glycerate produced from glycolate by the glycerate pathway. It is not clear whether two types of glycerate kinase arc also formed by the Crookes strain of E . coli, but probably the enzyme studied in this organism (78) corresponds to glycerate kinase I. C. MOLECULAR PROPERTIES 1. Purification and State of Purity

A crystalline preparation of glycerate kinase was obtained from E. coli (Crookes strain) after 4000-fold purification (78). An especially effective step in the purification procedure was chromatography on DEAESephadex, which afforded a 150-fold purification. (The peak of activity that emerged from the column was followed by some trailing material that eluted a t a higher salt eonccntration, perhaps representing a small

14.

GLYCEROL A N D GLYCERATE KINASES

507

amount of glycerate kinase 11.) The crystalline preparation migrated as a single component upon electrophoresis on cellulose acetate a t several pH values. A partial purification of the glycerate kinases from E. coli K12 was effected by chromatography on DEAE-Sephadex (79). This procedure also results in the partial separation of glycerate kinases I and 11, the latter eluting a t a slightly higher salt concentration. 2. Stability

The glycerate kinase from the Crookes strain of E. coli is quite stable in 5 mM potassium phosphate buffer, pH 7.0, a t temperatures below lo", even in the presence of 45% ethanol (78). The purified enzyme has been stored for 2 years a t -40" without loss of activity. In contrast, the glycerate kinases of E. coli K12 are rapidly inactivated a t low temperatures (79). At 0", 90% of the activity is lost in 24 hr, while at room temperature, the loss in that time is only 10%. This striking cold-lability may be indicative of differences in the enzymes from the two strains of E. coli; on the other hand, it could be because the enzyme from the Crookes strain was studied in 5 mM potassium phosphate, pH 7.0, whereas the enzymes from E. coli K12 were investigated in buffers that also contained 2 mM MgC12, 1 mM EDTA, and 0 . 1 4 . 4 M NaCl. The two glycerate kinases from E . coli K12 differ in their stability a t elevated temperatures (79). At 49", glycerate kinase I and I1 have half-lives of 90 and 10 min, respectively. The glyccrate kinase from the Crookes strain of E. coli is totally inactivated by M p-hydroxymercuribenzoate or iodoacetate, suggesting that sulfhydryl groups are essential for enzymic activity (78). D. CATALYTIC PROPERTIES Glycerate kinase can convert only one-half of added Dpglycerate to products (77) and is completely inactive with L-glycerate (79), while D-glycerate is converted stoichiometrically to products (79), suggesting that it is the sole substrate. The enzyme from the Crookes strain has an t e while both enzymes apparent K , of 0.24 mM for ~ , ~ ( ? ) - g l y c e r a (78), from E. coli K12 have I<, valucs of 0.07 M for D-glycerate and 0.05 M for ATP (79). The product of the reaction has been shown to be 3-phospho-D-glycerate by paper chromatography and enzymic assay (78). Assuming a molecular weight of 100,000, the glycerate kinase from E. coli (Crookcs strain) has a turnover number of 21,000 min-l (78). Glyceratc kinase requircs tlivalcnt cations for activity, being com-

508

JEREMY W. THORNER AND HENRY PAULUS

plctely inactive in the presence of M EDTA (78). The requirement can be satisfied by lO-'M Mg?+ and, partially, by &InY+,Fez+,or Caz+. The activity of the enzyme is markedly stimulated by potassium ion ( 7 7 ) . All three bacterial enzymes studied are most active between pH 7.0 ant1 7.5, hut the pH range in which glyccrate kinase I from E. coli K12 is active is broader than that for glycerate kinase I1 (78, 79). The very limited studies on the glycerate kinases from E. coli have revealed no cvidence for any type of regulation of their activity. Of obvious interest, however, is the cold inactivation of the enzymes from E . coli K12, since this phenomenon is ordinarily confined to enzymes with subunit structure ( 8 2 ) . The relationship between the three enzymes described here, and espccially that between glycerate kinase I and I1 of E . coli K12, is also of considerable interest. It remains to be shown whether these enzymes represent different polypeptide chains or whether they are modified forms of a single type of protein.

82. J. Jarabak, A. E. Seeds, Jr., and P. Talalay, Biochemistry 5, 1269 (1966).

Microbial As-artokinases PAOLO TRUFFA-BACHI I . Introduction . . . . . . . . . . . A . Historical Background . . . . . . . B . The Reaction Catalyzed . . . . . . . C . Methods of Assay . . . . . . . . I1. Escheiichin coli Aspartolrinases . . . . . . . A . Three Isofunctional Enzymes . . . . . . B . Aspartokinase I-Homoserine Dehydrogenase I C . Aspartokinase 11-Homoserine Dehydrogenase I1 . D . Aspartokinase I11 . . . . . . . . I11. Other Coliform Bacteria . . . . . . . . . I V . Aspnrtokinase Regulated by Concerted Feedback Inhibition A . Rhodopseudomonas capsulatus . . . . . B . Othcr Sonsulfur Photosynthetic Bacteria . . . C . Bacilli . . . . . . . . . . . D . Pseudomonads . . . . . . . . . E . Other Genera . . . . . . . . . V . Rhodopseudomonas spheroides . . . . . . . 1'1. Succharomyces cerevisiae . . . . . . . .

.

. . . . . . .

. . . . .

. . . . . .

. . . . . . . . . . . . .

. *

. . .

509 509 511 512 513 513 515

540 542 544 544 544 544 546 551 552 552 553

.

I Introduction

A . HISTORICAL BACKGROUND In 1948. Teas et a1. (1) studied a mutant of Neurospora crassa which required either homoserinc or methionine plus threonine for growth . Further evidence that homoserine was a common precursor of threonine and methioninc was adduced when a methionine-requiring mutant of the 1. H . J . Teas. N . H . Horowitz. and M . Fling. J . Biol . Chem. 172, 651 (1948) . 509

5 10

PAOLO TRUFFA-BACHI

same organism was found to accumulate threonine and homoserine ( 2 ) . Several organisms, when grown on labeled acetate, had similar isotope distributions in aspartate and threonine (3-6). The results of isotopic competition studies were consistent with the status of homoserine as a precursor of threonine (6). I n 1954, Cohen and his colleagues (7-10) found that bacterial suspensions and crude extracts of a threonine-requiring mutant were able to reduce aspartate to homoserine and that homoserine was a precursor of threonine in experiments performed with cell-free extracts in the presence of homoserine, ATP, and pyridoxal phosphate (10). The sequence of events from aspartate to threonine became clear when Black and Wright (11-13)discovered two new intermediates of the pathway, namely, p-aspartyl phosphate and aspartate-p-semialdehyde. The sequence of reactions catalyzed is as follows: HOOC -CH,-CH-COOH

I

ATP paspartokinase

=

H,O,P*

&OsP- OOC-CH,-CH-COOH I

m,

OOC-CH,-CH-COOH I

m,

NH, L-aspartic acid

p-aspartyl phosphate NADPH OHC- CHa-CH-COOH

h

ASA-dehydrogenase

aspartate-p-semialdehyde (MA) OHC-CHa-CH-COOH I

NH,

NADPH or NADH

hornoserine dehydrogenase

=

CI-&,OH-CI&-CH-COOH I

m

L-hornoserine

CH,OH-C~-CH-COOH I

NH,

ATP homoserine kinase

--

H,O,WH,C -CH,-CH-COOH I

m

homoserine phosphate

&o,w&c- CH,-CH-COOH I

NH,

pyridoxal phosphate threonine synthetase

+ CHs-CHO€i-CH-

I

COOH

m L-threonine

2. M . Fling and N. H. Horowitz, J . B i d . Chem. 190, 277 (1951). 3. G. Ehrensvard, Proc. Int. Congr. Biochem., Znd, 1962, p. 72 (1952). 4. C. Cutinelli, G . Ehrensvard, L. Reio, E. Saluste, and R. Stjernholm, Actn Chem. Scand. 5, 353 (1951). 5. R. B. Roberts, P. H. Abelson, D. B. Cowie, E. T. Bolton, and R. J. Britten, Carnegie Inst. Wash. Publ. 607 (1957).

15.

511

MICROBIAL ASPARTOKINASES

Further advances in the elucidation of isoleucine (14) and lysine (15) biosynthesis established the following simplified scheme for bacterial synthesis of threonine, isoleucine, methionine, and lysine: Aspartate

---+

p-agpwl phosphate

-

semialdehyde

L homoserine

1

3

diaminopimelate

-

~-methionine

L-threonine

1

L-isoleucine

The uncovering of the main mechanisms of regulation of biosynthesis, i.e., repression and feedback inhibition, introduced a new dimension in the study of this pathway: The repression of the synthesis of an early enzyme of the biosynthetic pathway, such as aspartokinase, or its efficient feedback inhibition, by any one of the essential metabolites should create insuperable difficulties for the microbial cell because it would limit the supply of the synthesis of the common intermediate necessary for the synthesis of the other essential metabolites of this pathway. I n the course of this chapter, we shall see that several different solutions have been developed in various organisms to overcome this difficulty.

B. THEREACTIONCATALYZED Aspartokinases of different microbial sources all catalyze the reaction: COOH I

7%

CHNH, I COOH

+

Aspartic acid

ATP

WZ+

0 II I FH2

\

00

+

ATP

yNHa COOH

@-Asparty1phosphate

~~

6. A. M. Delluva, Arch. Biochem. Biophys. 45, 443 (1953). 7. G. N. Cohen and M. L. Hirsch, C. R . Acad. Sci. 236, 1302 (1953). 8. G.N. Cohen and M. L. Hirsch, J . Bacteriol. SO, 182 (1951). 9. M.I,. Hirscli and G. N. Cohen, Biochim. Biophys. Actu 15, 560 (1954). 10. G. N. Cohen, M. L. Hirsch, S. B. Wiesendanger, and B. Nisman, C. R. Acatl. Sci. 238, 1746 (1954). 11. S. Black and N. G. Wright, J . Bwl. Chem. 213, 27 (1955). 12. S. Black and N. G. Wright, J . Biol. Chem. 213, 39 (1955). 13. S. Black and N. G. Wright, J . Biol. Chem. 213, 51 (1955).

512

PAOLO TRUFFA-BACHI

Insofar as they have been tested, no other natural amino acids or D-aspartate are substrates of this reaction. At p H 8.0 and a t 15”, using the yeast enzyme, Black and Wright (11) have found the equilibrium constant of the reaction to be

a value close to that found for the analogous reaction involving ATP and 3-phosphoglyceric acid (16). Aspartokinases of all sources are activated by Mgz+ions. I n several cases, Mgz+can be replaced by MnZ+or even Fez+. C. METHODS OF ASSAY

Activity can be determined by ( 1 ) measuring the amount of aspartohydroxamate formed by incubation of the enzyme with the substrates and hydroxylamine (11, 17), (2) coupling the reaction with aspartate semialdehyde dehydrogenase (aspartokinase-free) (12), or (3) coupling the reaction with pyruvate kinase and lactic dehydrogenase (17,18). The main advantage of methods (1) and (3) are their sensitivity and rapidity. I n method (2), the aspartyl phosphate formed is reduced in the presence of NADPH to aspartate-P-semialdehyde. The rate of oxidation of NADPH in the absence of aspartate must be substracted from the total rate observed. I n the cases where an NADPH-specific homoserine dehydrogenase is present, since aspartate-P-semialdehyde can be further reduced to homoserine, this method suffers from imprecision. Method (3) cannot be used with crude extracts since it is based on the ADP requirement for the pyruvate kinase and since large amounts of ADP can be generated by aspartokinase-independentATP cleavage. With purified enzyme, it presents the advantage of keeping the concentration of ATP constant during the reaction. I n the case of Enterobacteriaceae, we shall see that there exist in the same organism several isofunctional enzymes, the first of which can be totally inhibited by threonine and the second by lysine; the activity of the third is not inhibited by methionine, but its synthesis is repressed by 14. 15. 16. 17. 1970. 18.

H. E. Umbarger and E. A. Adelberg, J. Biol. Chem. 192, 883 (1951). Y. Yugari and G. Gilvarg, J . Biol. Chem. 240, 4710 (1965). T. Bucher, Biochim. Biophys. Acta 1, 292 (1947). P. Truffa-Bachi and G. N. Cohen, “Methods in Enzymology,” VoI. 17, p. 694,

D. E. Wampler and E. W. Westhead, Biochemistry 7 , 1661 (1968).

15.

MICROBIAL ASPARTOKINASES

513

this amino acid. In consequence, whichever method of assay is chosen in the case of crude extracts, it should include an assay in the presence of lysine, threonine, and both amino acids. Any residual activity in the presence of the two inhibitors represents the methionine-repressible enzyme. In Enterobacteriaceae the threonine-sensitive aspartokinase is associated with the threonine-sensitive homoserine dehydrogenase activity, and the methionine-repressible aspartokinase is associated with the methionine-repressible homoserine dehydrogenase. The homoserine dehydrogenase activity can be measured either in the forward direction using aspartate-/I-semialdehyde and NADPH (in this case a blank without aspartate-/I-semialdehyde should be run and the NADPH oxidation by NADPH oxidase substracted) or in the reverse direction using homoserine and NADP (13,1 7 ) . The maximum velocity of the reverse reaction is only one-twelfth that of the forward reaction but has the advantage of using commercial substrates. Aspartic semialdehyde can be synthesized from allylglycine according to Black and Wright (12).

II. Escherichia coli Aspartokinares

A. THREEISOFUNCTIONAL ENZYMES Of all the amino acids known to occur naturally as protein constituents only two, L-lysine and L-threonine, were found to influence the aspartokinase activity of crude extracts (19). The unnatural isomers are inactive. Maximal inhibition is reached a t concentrations of 2 mM. The concentration required for half-maximal inhibition is approximately 0.5 mM for L-threonine and 0.3 m M for L-lysine. The data of Fig. 1 show that when lysine and threonine are added simultaneously the total inhibition is the sum of that observed for each independently. The fact that the inhibitions were independent and additive suggested that crude extracts contain two different aspartokinases, one of which is inhibited by lysine and the other by threonine. This interpretation was soon borne out by the physical separation of the two enzymic activities (Table I ) . The synthesis of the lysine-sensitive enzyme is repressed when the 19. E. R. Stadtman, G. N. Cohen, G. Le Bras, and H. De Robichon-Szulmajskr.

J. B i d . Cheni. 236, 2033 (1961).

514

PAOLO TRUFFA-BACHI

I 00

80 60 c Q) c

2

40

a" 20

0 0

1

2

3

4

Inhibitor concentration, mM

FIG. 1. Effects of L-lysine and L-threonine or both on the inhibition of aspartokinase activity (19).

organism is grown in the presence of lysine; Freundlich (20) showed later that the synthesis of the threonine-sensitive enzyme is subject t o a multivalent repression by threonine and isoleucine, a point which was confirmed later (21).When Eschem'chiu coli K12 is thus grown under one or the other repressive conditions, only one type of aspartokinase is found which can be totally inhibited by its specific feedback inhibitor (19). TABLE I SEPARATION OF LYSINE-SENSITIVE AND THREONINE-SENSITIVE ASPARTOKINASE~-~

Protein fraction Sonic extract Streptomycin supernatant NH4S0, precipitate, 0-377& Dialyzed NH4S04 precipitate, 0-377: N&S04 precipitate, 37-50%

Protein (mg)

Total aspartokinase (units)

blysine

bthreonine

1225 750 395 -

208,000 200,000 91,000 46,000

33.3 30.4 53.6 87.7

50.2 49.7 17.0 2.7

285

110,000

3.6

73.0

Inhibition (%) by

Data from Stadtman et a / . (19). Aspartokinme activity of each fraction w m measured in the presence of 10 mM dysine and L-threonine. Activities are expressed in arbitrary units. a

20. M. Freundlich, Biochem. Biophys. Res. Commun. 10, 277 (1963). 21:G. N. Cohen and J. C. Patte, Cold Spring Harbor Symp. Quant. Biol. 28, 513

(1963).

15.

515

MICROBIAL ASPARTOKINASES

In the original report, it was observed that growth on methionine did not significantly influence the level of aspartokinase; this was an unexpected result (19). The finding that a single organism may contain more than one enzyme catalyzing the same biochemical reaction was not uncommon even in 1960. The discovery, however, of the existence of two aspartokinases in E . coli was of special significance because it offered a reasonable explanation for the multiplicity of enzymes. The synthesis of multiple enzymes which catalyze the formation of the common precursor, aspartyl-P, each of which is independently subject to feedback inhibition and to repression by different end product metabolites, presented a rational solution to the question posed. Aloreover, Patte et al. (22) demonstrated in 1967 the existence of a third aspartokinase in small amounts in E. coZi K12. This enzyme is not subject to feedback inhibition by an end product metabolite, but its synthesis is repressed by methionine. The accompanying tabulation describes the control of the three activities, which haye been separated physically and characterized as distinct protein species. Enzyme Aspartokinase I Aspartokinase I1 Aspartokinase I11

Repressor

+

Threonine isoleucine Methionine Lysine

Allosteric inhibitor Threonine None Lysine

B. ASPARTOKINASE I-HOMOSERINE DEHYDROGENASE I Aspartokinase I has as an integral property another threonine-sensitive catalytic activity, namely, homoserine dehydrogenase I. After mutagenesis organisms can be selected in which the aspartokinase I and the homoserine dehydrogenase I both show modified allosteric properties. Invariably, both activities were modified in the same way. It was at first thought that the two enzymes possessed a common polypeptide chain responsible for the allosteric properties of both enzymes (23). It was then found that mutants which had lost the homoserine dehydrogenase I activity had also lost the aspartokinase I activity. Mutations which caused reversion enabled both activities to be recovered simultane22. J. C. Patte, G . Le Bras, and G . N. Cohen, Biochim. Bwphys. Acta 136, 245 (1967). 23. C. K. Cohen, J. C. Patte, and P. Truffa-Bachi, Biochem. Biophys. Res. Commun. 19, 546 (1965).

516

PAOLO TRUFFA-BACHI

ously. From this it seemed probable that both activities resided in the same protein complex. A considerable purification of the wild type enzyme failed to result in any separation of the two activities ( 2 4 ) . 1. Purification and Criteria of Honaogeneity

The purification steps used (25, involved, after sonic disruption of the bacterial suspension, precipitation of the nucleic acids by streptomycin, two-step ammonium sulfate fractionation, chromatography on DEAESephadex A-50, and chromatography on hydroxyapatite. The overall yield is about 50%. The purification must be carried in the presence of L-threonine to prevent activity loss. The ratio of the homoserine dehydrogenase activity to the kinase activity remains coiistant throughout the purification. The protein gave a single, sharp, and symmetrical peak throughout the sedimentation process in the ultracentrifuge. When a solution of the enzyme was subjected to disc gel electrophoresis, a minimum of four protein bands was detectable with variable intensities of staining by amido black. However, all thcse bands, except the one which ran fastest, showed homoserine deliydrogenase activity. Measurement of the enzymic activity in the presence or absence of 10 mM L-threonine, the allosteric effector, showed that the second and third band represented desensitized enzyme ; the purple color of reduced tetrazoliuin appeared as rapidly as in a control gel revealed without threonine. The top band shows activity only after a long lag because of the presence of the inhibitor and presumably represents native enzyme. Alkaline conditions as used in the electrophoresis dcstroys the aspartokinase activity and desensitizes the homoserine dehydrogenase activity. L-Threonine protects against this denaturation (26'). Gel electrophoresis in the presence of 10 mM L-threonine yielded one main band of protein and of activity, with only a second faint band of desensitized enzyme being still detectable, but represented at most a few percent of the total protein. The position of this minor band coincided with that of the major denscnsitized bands present in the experiment when the electrophoresis was run without threonine. The multiplicity of bands observcd under some conditions appears to have arisen from alkaline denaturation of the enzyme. 24. J. C. Patte, P. Truffa-Baclii, and G. N. Coilen, Biochim. Biophys. Acln 128, 426 (1966). 25. P. Truffa-Bachi, R. van Rapenbusch, .I. Janin, C. Gros, and G. N. Cohen, Elm. J. Biochem. 5, 73 (1967). 26. P. Truffa-Baclli, G. Le Bras, and G. S . Cohen, Biochim. Biophys. Actn 128, 450 (1966).

15. MICROBIAL

ASPARTOKINASES

517

2. Stability

The enzyme is stable a t room temperature either in the presence of 10-3M L-threonine or of 0.15M KC1. In the absence of threonine or T P N H the enzyme is very sensitive to low temperatures. The inactivation can be totally or partially reversed by rewarming the cold enzyme preparation at room temperature (27). 3. Extinction Coefficient

An extinction coefficient of 0.46 & 0.02 absorbancy unit cm?/mg had heen previously determined (281, using the biuret procedure, calibrated by measurements of the carbon and nitrogen content of the dried protein. Two independent measurements of the protein concentration of solutions of known ultraviolet absorbancy were later carried out, based on the quantitative amino acid analysis of an acid-hydrolyzed sample and on the determination of the refractive index increment. Both measurements gave a value of 0.63 k 0.03 absorbancy unit cm2/mg ( 2 9 ) . The introduction of this new value obviously leads to a reassessment of the stoichiometry of ligand binding and of the molar specific activity of aspartokinase and homoserine dehydrogenase.

4. lllolecular Weight The sedimentation coefficient obtained upon linear extrapolation to infinite dilution is s20,w = 11.5 (-1-0.2) X 10-13 sec. This agrees very well with the results of sedimentation in a sucrose gradient using catalase and alcohol dehydrogenase as markers, where the peaks of both activities sec ( 2 5 ) . have an s value of 11.0 f 0.5 X Equilibrium sedimentation runs, using interference optics, gave mean values, corresponding to the three protein concentrations used, of lIW,,,, = 357,000 f 14,000 and MW, = 350,000 f 30,000. The ratio l I W z :MW,,, is not significantly different from unity, an additional confirmation of lack of heterogeneity. The estimated molecular weight of the native enzyme is 360,000 f 20,000 ( 2 6 ) . The same molecular weight and the same sedimentation coefficient were also found 27. G. S . Cohen, J. C. Patte, P. Truffa-Bachi, C. Sawas, and M. Doudoroff, Colloq. I n t . Cent. Nnt. Rech. Sci. 243 (1965). 28. J. Janin. R. van Rapenbusch, 1’. Tniffa-Bachi, and G. N. Cohen, Eur. J . Riochem. 8, 128 (1969). 29. F. Falros-Kelly, J. Janin, J . C. Saari. M. Veron, P. Truffa-Bachi, and G. K. Cohen, Eur. J. Biochem. 28, 507 (1972).

518

PAOLO TRUFFA-BACHI

by Wampler et al. ( S O ) . Light scattering studies ($9) confirm a value for the molecular weight of 358,000 k 35,000 daltons for the native enzyme. A change in the elution pattern and sedimentation in sucrose gradients of the aspartokinase (18) and homoserine dehydrogenase (31, 32) activities, has led several groups to propose that a major effect of threonine is a polymerization of the enzyme. Cunningham et al. (31) actually concluded that the molecular weight of the native protein is in the 80,000120,000 range, with a sedimentation coefficient of 6.3 S in the absence of threonine. However, these experiments were performed in the cold and in a buffer containing no KCl. Patte et al. (33) have noted that polymerization also occurs upon addition of KC1, K+ being essential for both catalytic activities (18, 33). The polymerized form (sedimentation coefficient near 11 S) is stable in the absence of threonine, at least at room temperature and in 0.15M KC1; moreover, this form does bind NADPH (see below), the addition of threonine modified significantly neither its sedimentation coefficient nor the binding of NADPH. Dissociation of the protein was actually observed in the absence of threonine and KCl and favored by low temperature which inactivates the homoserine dehydrogenase (33). 5. Tetrametric Structure Earlier studies from the author’s laboratory have led to the conclusion that the enzyme is an oligomer of 360,000 molecular weight (25, 34). The molecular weight of the subunits of aspartokinase I-homoserine dehydrogenase I, 60,000 daltons, which had been determined by high-speed sedimentation equilibrium in 6 M guanidinium chloride ( 3 4 ) ,was used as an indication for hexameric structure for the native enzyme. However, subsequent separations by sodium dodecyl sulfatc (SDS) gel electrophoresis gave a single protein band with a molecular weight of 84,000, higher than anticipated ( 2 9 ) . Since S-carboxymethylated protein showed the same behavior on SDS gels, spurious effects resulting from sulfhydryl oxidation could be eliminated. Similar studies by Clark and 30. D. E. Wampler, M. Tnkahashi, and E. W. Westhead, Biochemistiy 9, 4210 (1970). 31. G. N. Cunningham, S. R. Maul, and W. Shive, Biochem. Biophys. Res. Cornmiin. 30, 159 (1968). 32. J. W . Ogilvie and J. H. Sightler, Biophys. J. 8, TC5 (1968) (abstr.). 33. J. C. Patte. G. Le Bras, T. Loviny, and G. N. Cohen, Biochim. Biophys. Acln 67, 16 (1963). 34. P. Truffa-Bachi, R. van Rapenbusch, J. Janin, C. Gros, and C,. N. Cohen, Ew. J. Biochem. 7, 401 (1969).

15.

MICROBIAL ASPARTOKINASES

519

Ogilvie (35) also gave an apparent molecular weight of 83,000 for the subunit. Gel filtration in 6 M guanidinium chloride, a solvent known to denature the enzyme completely ( S 6 ) , gave a value of 88,000 f 5000 ( 2 9 ) . From both results a value of 86,000 f 4,000 for the subunits of the enzyme was assumed (29, 37). A value of MW,,, from ultracentrifugation data was 66,000 f 5,000, and small deviations from linearity were interpreted as a sign of incomplete dissociation of the protein (34). The difference between this value and those derived from SDS gel electrophoresis and from Sepharose chromatography in guanidinium chloride remains unexplained. This contrasts with the agreement between the results obtained by Rosenbusch and Weber (38) who used similar procedures to revise the molecular weight of the subunits of E. coli aspartate transcarbamylase. Recent studies of the behavior of the aspartokinase I-homoserine dehydrogenase I from another strain of E . coli in the ultracentrifuge are in agreement with a subunit molecular weight of 80,000, taking into account the nonideal behavior of the protein in the presence of 6 M guanidinium chloride ( 3 9 ) .The results of Wampler et al. (SO), which suggest the existence of active dimers of a molecular weight of 122,000 daltons in “Tes” or “Hepes” buffer, are difficult to reconcile with the present data. Further support for a tetrameric structure was provided by cross-linking the subunits with dimethyl suberimidate followed by disc gel electrophoresis in SDS ( 2 9 ) . Only four bands were seen when the enzyme was cross-linked at pH 8.5 in the presence of 0 . 3 M KCl, with the mobilities expected for the monomer, dimer, trimer, and tetramer of the subunits. Since the protein carries two enzymic activities, the question of whether its subunits are identical is of particular interest. Electrofocusing in denaturing solvent gives a single peak, suggesting that the subunits are not oiily identical in molecular weight but also carry the same net charge. 6 . Kinetic Parameters

The apparent dissociation constants of the substrates of homoserine dehydrogenase I have been published on incompletely purified preparations (33).Essentially identical values were obtained with the pure protein: K,,, NADPH = 4 X M ; K,n L-aspartate-p-semialdehyde = 35. R. B. Clark and J . W. Ogilvie, Biochemistry 11, 1278 (1972). 36. H. d’A. Heck and P. Truffa-Bachi, Biochemistry 9, 2776 (1970). 37. D. E. Wampler, Biochemistry 11, 4428 (1972). 38. J. P. Rosenbusch and K. Weber, J . B i d . Chem. 246, 1644 (1971). 39. W. L. Starnes, P. Munk, S. B. Maul, G. N. Cunningham, D. J. Cox, and W. Shive, Biochemistry 11, 677 (1972).

520

PAOLO TRUFFA-BACHI

1.2 X lo-'M. For the aspartokinase I, the values found for the pure enM. M and I(, L-aspartate = 1.5 X zyme are K , ATP = 4 X The values for the kinase do not differ from those published by Wampler and Westhead (18) and are identical whether or not the assay is performed with the aspartate semialdehyde assay. With a more rational assay using pyruvate kinase and phosphoenolpyruvate in order to keep the ATP concentration constant during the assay, Wampler and Westhead found a value of 1.8 X 10-+M for K , ATP (18). The molar specific activity a t 27' for both enzymes has been calculated to be 24,000 moles of aspartate semialdehyde reduced per minute per mole of dehydrogenase and 3400 moles of aspartate phosphorylated per minute per mole of aspartokinase (29).

7. Chemical Properties a. Amino Acid Analysis. The amino acid composition of aspartokinase I-homoserine dehydrogenase I is given in Table I1 ( 2 6 ) .A redetermination using carboxymethylated protein gave results in good agreement (99).I n particular the value of 10.8 k 0.8 carboxymethylcysteine found per 86,000 daltons is in good agreement with the cysteic acid content obAMINOACIDCOMPOSITION

OF

TABLE I1 ASPARTOKINASE I-HOMOSERINE DEHYDROQENASE Io Residues per 344,000

0

Amino acid

daltons

Lysine Histidine Arginine Aspartic acid Threonine Herine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Tryptophan

126 53 178 311 122 215 346 153 245 35 1 42 254 75 172 327 69 118 16

Data from Truffa-Bachi et al. (26).

15.

MICROBIAL ASPARTOKINASES

521

tained after performic acid oxidation (10.1 -I- 0.3 recalculated for the same molecular weight). The tryptophan content of the protein, previously determined from its ultraviolet absorption in 6 M guanidinium chloride (25),is 3.9 k 0.15 residues per 86,000 daltons when recalculated for the new value of the extinction coefficient.After reaction with 2-nitrophenylsulfenyl chloride, the number of tryptophan residues was found to be 3.4 2 0.1. Since incomplete reaction resulting from partial unfolding of the protein can be expected (36), it is reasonable to assume 4 tryptophan residues per subunit of molecular weight 86,000.

b. Sulfhydryl Groups. When using the new extinction coefficient, and with preparations freshly reduced by dithiothreitol, the number of titratable sulfhydryl groups determined by DTNB titration under denaturing conditions is equal to the number of carboxymethylcysteine or cysteic residues determined by amino acid analysis. Under nondenaturing conditions, the number of titratable sulfhydryl groups is from 24 to 26 per mole of native enzyme (MW = 344,000). Variable quantities of cysteine may form a disulfide bond. Carboxymethylation with [ W]iodoacetic acid, followed by reduction and carboxymethylation with [ **C]iodoacetic acid, with subsequent autoradiograms of the tryptic digest, show that the disulfide bond is made exclusively from the two cysteinyl residues of a single tryptic peptide (T85, see Table 111).

c. Partial Sequence Information. Tryptic hydrolysis of S- [ 14C]carboxymethylated aspartokinase I-homoserine dehydrogenase I or of the carTABLE I11 TRYPTIC PEPTIDESCONTAININQ CARBOXYMETHYLCYSTEINE AND/OR TRYPTOPHAN~ Peptides

Amino acid sequence or composition

T85

Ala-Asp-Ile-CMCys-G1u-Trp-Thr-ksp-Vrtl-Asp-Gly-Val-Tyr-Thr-CMCys-

T60 T30 T10 TNa TNb T10’ TB1 TB2 Ch6W

Tyr-Val-Gly-Asx-Ile-Asx-Glx-Asp-Gly-Val-CMCys-Arg Leu(CMCys,Asx~,Thr,Ser~,Glx~,Gly,Ala~,Val~,Ile,Tyr) (Asxa,Ser~,Glx,Pro,Gly~,Ala,,Val,Ile,Leu3,His)Ile-CMCys-Arg

Asp-Pro-Arg

a

Val-CMCys-Gly-Val-Ala-Asn-Ser-Lys

Thr(Thr,Glu,Pro,Ala,Ile,Phe)Gln(CMCys,Pro,Ilea,Leu)Lys (Asx~,Thr,Glx6,Pro,Gly,Ala~,Val,Leu,His,Trp)Lys Gln-Ser-Trp-Leu-Lys Thr-Leu-Ser-Trp-Lys CMCys-Val-Pro-Glx(CMC s,Asx2,Ser,Glx,Va12)Arg

Data from Falcoe-Kelly et al. (29).

* Chymotryptic peptide from the insoluble core.

522

PAOLO TRUFFA-BACHI

boxymethylated uniformly labeled 14C enzyme yielded 50-52 soluble peptides, four of which contained tryptophan. Autoradiograms showed the presence of six peptides containing carboxymethylcysteine. An insoluble core, after digestion with chymotrypsin, gave an additional carboxymethylated peptide. Present information about sequences of unique peptides is given in Table I11 (29). Automatic sequence analysis of the carboxymethylated protein showed the following N-terminal sequence (29) : NHrMet-Arg-VaI-Leu-Lys-Gly-Gly

...

Data from carboxypeptidase A digestion indicated a carboxy terminal sequence as Leu-Gly-Val-COOH. No amino acids were released by carboxypeptidase A in the absence of SDS, suggesting that in the native conformation of the enzyme, the carboxyl terminus is buried.

d. Identity of the Four Subunits. The yields of tryptic peptides, the number of unique carboxymethylated peptide, and, in particular, the presence of four unique sequences around tryptophan residues are consistent with the presence of identical subunits. I n addition, all four subunits have the same amino and carboxy terminal sequences (29). The results can be considered as a good indication that the four subunits are indeed identical in their primary structure. The sequence studies are in harmony with a molecular weight of 86,000. A consequence of the identity of the subunits is that the two distinct catalytic activities are carried by a single polypeptide chain. A similar situation has already been shown to exist for the bifunctional enzyme carrying the activities phosphoribosyl anthranilate isomerase and indole glycerophosphate synthetase in E. coli and in Salmonella typhiinuriuw (40, 4 1 ) . Another bifunctional protein carrying two distinct enzymic activities has also been ohtaincd by fusion of the his D and his C genes of the histidine operon in S. typhimurium ( 4 2 ) .The presence of distinct catalytic sites on a single polypeptide chain has already been discussed in detail ( 4 3 ) ,in terms of structure and evolution. As noted in Section I1 the two catalytic activities are carried by two independent regions of the polypeptide chain. 40. A. J. Blume and E. Balbinder, Genetics 53, 577 (1966). 41. T. E. Creighton, Biochem. J. 120, 699 (1970). 42. J. Yourno, T. Kohno, and J. R. Roth, Nature (London) 228, 820 (1970). 43. D. M. Bonner, J. A. De Moss, and S. E. Mills, in “Evolving Genes and Proteins” (V. Bryson and H. J. Vogel, eds.), p. 305. Academic Press, New York, 1965. 44. M. E. Goldberg, J . Mol. Bid. 46, 441 (1969).

15.

MICROBIAL ASPARTOKINASES

523

8. Ligand Binding The stoichiometries have all been corrected for the new values of the molecular weight and of the extinction coefficient of the protein ( 2 9 ) , whether they originate from the author's laboratory or not. This also applies to Section II,B,9.

a. Threonine Binding. The threonine binding experiments were performed using L-threonine I4C, either by equilibrium dialysis or by gel filtration on Sephadex G-25 columns preequilibrated with the ligand (28). I n a buffer containing 0.15 M KC1, saturation of the binding sites is reached a t free ligand concentration in the 0.3-0.5 mM range, where 8.2 f 0.3 moles of thrconine are bound to 1 mole of enzyme. Essentially the same values were obtained with aged samples of protein having lost 30% of their homoserinc dehydrogenase activity. These preparations lose none of their binding capacity toward the allosteric effector and the remaining activity is normally inhibited by threonine; on the other hand, treatment with N-ethylmaleimide, as well as other -SH reagents which desensitize the homoserine dehydrogenasc activity, leads to the loss of binding capacity toward threonine. The Scatchard plot of threonine binding shows that the binding is cooperative in the presence of 0.15 M KC1 (28). When the concentration of KC1 is lowered, the shape of the curvc changes and the binding is no longer sigmoidal ( 4 5 ) . The significance of this change will be discussed later. The number of molecules of threonine bound per monomer is approximately two. Similar valucs, 1.6 molecules per monomer, are obtained by Takahashi and Westhead ( 4 6 ) . Threonine binding reduces exchange of enzyme hydrogen with water ( 4 6 ) . I n the absence of threonine during the exchange out process, there are approximately 220 nonexchangeable hydrogens per mole, while in the presence of threonine, a t saturating concentrations, the number of core hydrogens is increased to 310. The binding of threonine evidently tightens the enzyme structure and increases the core size about 40%. b. Pyridine Nucleotide Binding. The binding of NADP' has been followed by the gel filtration technique using the l4C-labeled coenzyme. The linear Scatchard plot indicates a simple Michaelis type of binding. EXtrapolation leads to a value of 3.9 f 0.2 moles of NADP bound per mole of enzyme with an affinity constant one order of magnitude lower than the apparent I < , (28, 29). 45. J. Janin and G . N. Cohen, Eur. J . Biochem. 11, 520 (1969).

46. M. Takahashi and E. W . Westhead, Biochemistry 10, 1700 (1971).

524

PAOLO TRIJFFA-BACHI

The binding of NADPH was studied using the absorption a t 340 nm of the reduced coenzyme upon elution from equilibrated Sephadex columns. At saturating concentrations of NADPH near 23", and in a buffer containing 0.15M KC1, the number of bound NADPH molecules is between 2 and 4 molar equivalents, with an uncertainty of the order of 15%. The binding does not vary significantly when 1 mM L-threonine is added or when the KCI concentration is varied from 0.15 to 0.65M. Moreover, addition of the substrate of the reverse reaction, L-homoserine, does not lead to higher values. However, in contrast to the binding of L-threonine, the binding of NADPH by aged and partially inactivated samples of protein leads to a decreased binding (28). The enhancement of the coenzyme fluorescence or the appearance of transfer fluorescence has been used to titrate a given solution of protein by increasing amounts of NADPH. A sharp titration is observed when the initial protein concentration is of the order of 2-10 rJJM, and the end point corresponds to 4 molar equivalents of added coenzyme. Other studies a t a different protein concentration allow one to calculate a value of 0.3 pM for the affinity constant, two orders of magnitude lower than the apparent K,. Here again, the presence of threonine does not influence the amount of coenzyme bound a t saturation (28). The binding of NADPH has also been studied by circular dichroism techniques (36). NADPH binding results in the appearance of an extrinsic dichroic band a t 348 nm. The change in molecular ellipticity was used to titrate the protein with NADPH. The number of sites found, 4 2 0.3 per tetramer, is in good agreement with the fluorometric titration. In conclusion, each subunit of the cnzyme possesses a binding site for the NADPH.

c. Covalent Binding of an ATP Analog. 6-Mercapto-9-P-~-ribofuranosylpurine 5'-triphosphate, an ATP analog with an -SH replacing the 6-NH, group reacts specifically with sulfhydryl groups at the adenosine triphosphatc sites of myosin (4'7). This compound is a suhstratc of aspartokinase I-homoserinc dehydrogenase I ( 4 8 ) . The disappearance of aspartokinase activity during reaction of aspartokinase I-homoserine dehydrogenase I with the ATP analog is a pseudofirst-order process to a t least 80% of reaction. During the same time period, there is no loss in homoserine dehydrogenase activity. Although the dehydrogenase activity is unaltered by reaction of the enzyme with the mercapto analog, the ability of L-threonine t o inhibit 47. A. J. Murphy and M . F. M o d e s , Biochemistry 9, 1528 (1970). 48. P. Truffa-Badii and H. d'A. Heck, Biochemistry 10, 2700 (1971).

15. MICROBIAL ASPARTOKINASES

525

the dehydrogenase activity, i.e., the sensitivity of the dehydrogenase to threonine, is lost. The selectivity of reaction with the mercapto analog adequately explains both the pseudo-first-order kinetics of aspartokinase inactivation and the constancy throughout reaction of the homoserine dehydrogenase activity and of the protein molecular weight. I n contrast to these results, when nonselective sulfhydryl reagents such as p-mercuribenzoate or DTNB are employed, complex kinetics are observed, owing to reaction at numerous sites ( 2 5 , 4 9 ) .In addition, inactivation of the dehydrogenase activity and dissociation of the protein take place following extensive reaction with p-mercuribenzoate ( 4 9 ) . Threonine, but neither aspartate nor, unexpectedly, ATP, protects the enzyme against the mercapto ATP analog inactivation. The number of molecules of the analog bound per mole of native enzyme, under a variety of experimental conditions including various times of incubation, is consistently 3.6-4.2. In conclusion, each subunit of the aspartokinase I-homoserine dehydrogenase I possesses a binding site for the mercapto ATP analog that may or may not correspond to an aspartokinase site per monomer.

9. -SH Titration and Its Effects Studies of the effects of p-mercuribenzoate on the protein had shown that thc aspartokinase activity and the inhibition of homoserine dehydrogenase activity by L-threonine were sensitive to this reagent indicating the importance of -SH groups. Moreover, the allosteric effector, L-threonine, could protect enzymic activities against the effects of p-mercuribenzoate ( 4 9 ) . Quantitative estimation of the -SH groups could be achieved on the pure protein by the use of Ellman’s reagent, DTNB ( 2 5 ) . Total -SH content was determined in the presence of SDS or urea and led to a value of 40-44 cysteine residues per 344,000.When the native protein was exposed to DTNB in the absence of denaturing agent, only 24-26 -SH residues per mole of enzyme were readily available. When the reaction was performed in the presence of 2 m M L-threonine, essentially no -SH groups could be titrated. In the presence of the allosteric effector, the protein takes a conformation such that as many as 24-26 cysteine residues become “buried.” Titration of the available -SH groups by DTNB in the absence of 49. P. Truffa-Bachi, G. Le Bras, and G. IV. Cohen, Biochim. Biophys. Acta 128, 440 (1966).

526

PAOLO TRUFFA-BACHI

E

.-e

5

0

-e 0

E

\

c 0 .c

8

2

n i5

-

No protection

-

-

15 -

E, Y

l 0 n

$

-

t 2 m M L-Threonine

-

Time (minutes)

F I ~2.. Titration of -SH groups with DTNB ($6)

L-threonine has the same qualitative effects on both activities of the protein as was previously reported for p-mercuribenzoate ( 4 9 ) . Aspartokinase inactivation and desensitization of the homoserine dehydrogenasc activity were concomitant with the titration of the reacting -SH groups (Fig. 2 ) . At any given moment, the remaining aspartokinase activity was fully sensitive to threonine inhibition. When the reaction with DTNB was performed in the presence of L-threonine, no effect on the enzymic activity was found ( 2 5 ) . 10. Conformational Changes

Conformational changes of proteins upon addition of ligands are well documented, and two major theories have been proposed for their interpretation: the “allosteric” model of Monod et nl. (50) and the sequential model of Koshland et al. ( 5 1 ) . The allosteric inhibition by L-threonine of both aspartokinase and homoserine dehydrogenase activities is based on the conformational change of the protein upon addition of threonine and can be visualized by different techniques.

a. Difference Spectra. An intense difference spectrum in the 250310-nm range was obtained by comparing similar solutions of protein in a phosphate buffer containing 0.15M KC1 with and without 1 mM 50. J. Monod. J. Wyman, and J. P. Changeux, J. Mol. B i d . 12, 450 (1965). 51. D. E. Koshlmd, G . Nemetlry, and D. Filmer, Biochemistry 5, 365 (1966).

15. MICROBIAL

527

ASPARTOKINASES

260

280 300 Wavelength (nm)

320

FIG.3. Difference spectra in the presence of threonine. Identical quartz cuvettrs placed in the sample and reference compartments of the Cary 15 spectrophotometcr contain similar solutions of protein (9 p M ) with the following additions. Base line: reference, no addition; sample, no addition. Dashed line : reference, no addition; sample, 0.5 mM t-threonine. “Aspartate” spectrum : reference, 15 mM L-aspartate ; sample, 15 mM L-aspartate and 0.5 mM L-threonine. “KCl” spectrum : reference, 0.15 I14 KCI ; sample 0.15 KCI and 0.5 mM L-threonine ( 4 4 .

L-threonine (28). KCl can be replaced by another ligand of the protein, aspartate (Fig. 3) (45). The difference spectra were used to define two conformations of the protein. The T form was defined as the form with huried chromophores (low absorption at 269 nm and high absorption a t 289 nm) and R as the form with exposed chromophores. A straightforward interpretation of the spectra suggests that T is stable in the absence of added ligand or in the presence of threonine, and R is stable in the pres( m e of potassium ions or of aspartate. The complexity of the spectra indicates that several aromatic residues are involved. It is not easy to determine whether these effects are equally distributed on each of the four suI)units or concern only some of them.

b. Effects of Ligands on the Protein Fluorescence. The change of the absorption spectrum of the protein upon addition of threonine in the presence of 0.15 M KCl is associated with a quenching and a slight shift of the protein fluoresrencc (28). A study of the excitation and emission spectra shows that the fluorescence excitation a t 295 nm is shifted to longer wavelengths by about 3 nm when threonine is added (28, 46). Addition of aspartate or potassium ions to the protein (in the absence of

528

PAOLO TRUFFA-BACHI

threonine) causes little change of the fluorescence obtained after excitation a t 295 nm: The quantum yield of the tryptophan residues in the R and T forms defined above does not differ significantly at this wavelength. Thus, the effects of the ligands on the fluorescence excited a t 295 nm cannot be accounted for by the two-state model suggested by the study of the ultraviolet absorption. The observed shift to longer wavelengths may be a short range effect of the binding of threonine or reflect the existence of another conformation of the protein. In contrast with these results the study of the fluorescence excited a t wavelengths below 280 nm, where the tyrosine residues may contribute,

eon asp art ate] (mM) (A)

[K'] (MI

(61

FIG.4. Effects of aspartate and of K' on the allosteric equilibrium. The values of R / ( R + T ) in the presence of various concentrations of ligands are derived from measurements of the fluorescence intensity of the protein. (A) Effects of aspartate. The following supplements are added to buffer P: (0) none, ( A ) 30 mM KCl. and ( X ) 40 p M L-threonine. (B) Effects of K'. The following supplements are added to the tris-Cl buffer (0) nonc. (A)2 mM L-aspartate, and ( X ) 40 p M L-threonine (46).

15.

529

MICROBIAL ASPARTOKINASES

is in full agreement with the two-state model: Addition of threonine quenches the fluorescence only if aspartate or potassium ions are present. Under proper conditions the measurement of the intensity of the protein fluorescence provided a determination of the relative proportions of the major conformations of the protein. The variation of the fluorescence excited a t 280 nm and observed a t 335 nm (near the isoemissive point of the effect specific for threonine) is compatible with a two-state model. If F R and F, are the fluorescence intensities of a given protein solution in the R and T conformation, the recorded intensity in the presence of intermediate concentrations of the ligands will be F:

where T and R are the relative concentrations of the two forms. The values of F T and F, can be measured under similar conditions of saturating concentrations of threonine and of aspartate (or of potassium ions), and such functions as R / ( R T j , T / ( R T ) or R / T can be derived from the value of F:

+

+

T / R = [ ( F R - F)/(F - FT)] (2) Figure 4 shows the variations of the R / ( R T ) function when increasing amounts of aspartate or of K+ ions are added to the protein in the presence or absence of other ligands. Aspartate and K+ give positive homotropic effects (sigmoidal curves) and positive heterotropic effects, but they give negative heterotropic effects with threonine. When threonine is added to protein solutions containing various amounts of the other ligands (Fig. 5 j , similar negative heterotropic effects are observed. Positive homotropic effects imply that the configuration change in one subunit of the protein leads to, at least partially, a similar change in other subunits. I n the extreme case, n subunits change conformation in a concerted way; the function T / R of the concentration A of a ligand binding T and R with dissociation constants KT and K R can then be written

+

\Vherc L+ is the value of T / R in the absence of ligand A and depends on the concentrations of the other ligands. When no information is available on the values of K T , KR,and n, a demonstration of the conccrted model based on formula (3) may be inconclusive. But in the case of threonine, a value of KT = 40 pLM can be derived from the study of the binding to the protein a t low concentration of K + ; wc also know that the binding of threonine to R is negligible (28). Equation (3) then simplifies to

530

PAOLO TRUFFA-BACHI

[~-Threonine](mM)

(A)

0

I

0

I

ai

a2

I

03

[~-Threonine](mM)

(B) FIG.5. Effects of threonine on the allosteric equilibrium. L-Threonine is added to a solution of protein in buffer P containing the following supplements: (A) (B) none, ( 0 )60 mM KC1, ( x ) 0.15M KCl, and (0) 0.45M KC1. (B) (B) none, ( A ) 5 mM L-aspartate, and (0) 20 mM L-aspartate (46).

T / R = L+(I

+ A/KT)n

where A is now the concentration of threonine and K T = 40 p M . A “modified Hill plot” of log ( T / R ) against log (I A/&.) should then give a straight line of slope n, the number of cooperating subunits, and extrapolate to log (L+) for A = 0. Figure 6 demonstrates that seven curves of Fig. 5 describing the effects of threonine on the R - T equilibrium yield parallel straight lines in the double logarithmic plot with a value of n close to three. Reciprocally, plots of ( T / R )1/3 against A (59) yield straight lines which extrapolate to A = -KT on the horizontal axis (Fig. 7). Convergence of the lines derived from the curves of Fig. 5 demonstrates that the dissociation constant KT of threonine is essentially

+

52. D. Blangy, H. Buc, and J. Monod, J . Mol. Biol. 31, 13 (1966).

15.

MICROBIAL AYPARTOKIKASES

+'

t

53 1 0

FIG.6. Double logarithmic plot of the effects of threonine. The values of T / R come from the same experiments as those of T ( R + T ) in Fig. 5 . The concentration of threonine added is A , and KT = 40 pM is an estimate of the dissociation constant. The slope of the dashed line corresponds to n = 3. The notations are the same as in Fig. 5 (46).

independent of the presence or absence of other ligends; moreover, the corresponding value of KT = 30 pM agrees with the value independently determined from binding measurements. The general properties of the R and T form are shown in Table IV. This description implies that the protein behaves as a pair of concerted trimers, and a discussion of this model has been presented ( 4 5 ) . It is clear that this must be rejected in view of the new structure pro-

I~-Threonine)(mM)

FIG. 7. Study of (T/R)'''. Effects of threonine. Plots of (T/R)'I" as a function of the concentration A of the ligand added, threonine in this case, yield a bundle of straight lines extrapolating to (L')''" on the vertical axis and to -KT on the horizontal axis. In this case n = 3 is determined from the slope of the double logarithmic plot (Fig. 6) and KT is found to be of the order of 30 p M . The notalions arc the same RS in Fig. 5 (466).

TABLE IV

GENERAL PROPERTIES OF R

AND

T form

Properties

T CONFORMATIONS~ R form

Remarks

~

Binding of ligands (dissociation constants) Threonine Aspartate I(+ ions NADPHc Spectrophotometric properties Relative fluorescenceintensity Tryptophan and tyrosine residues Enzymic activities Aspartokinase Homoserine dehydrogenase

Shifts the equilibrium toward: KT = 3 0 p M KT > 20mM Low affinity K T = 0.3 p M

K R = 1.8 mMb Kg = 3 mM High affinity K R = 0.3 p M

100

116

Buried

Exposed

None 10-15%

Active 10%

T form R form R form No detectable effect Excitation a t 290 nm, emission a t 335 nm Difference spectrum

T does not bind the substrate Different configuration of the active sites Y

Data from Janin and Cohen ($55). * Binding of threonine a t saturation of KCl aspartate. c Fluorescence titration. 0

r 0

e

I I

s -3

?

W ?-

c)

E!

15.

MICROBIAL ASPARTOKINAGES

533

posed for the protein (29). More data are needed to know whether aspartokinase I-homoserine dehydrogenase I is a concerted tetramer following the allosteric model or has a more complicated behavior. c. Relaxation Studies. Fast kinetic studies of this system were performed first by Barber and Bright (53) who demonstrated that the inhibition of the homoserine dehydrogenase activity appears with a significant delay after the addition of threonine. They interpreted this as an expression of a conformation change of the protein. The effects of the ligands on fluorescence and ultraviolet absorption spectra of aspartokinase I-homoserine dehydrogenase I reflect its overall conformation changes; T to R and R to T reactions can be followed in the stopped-flow apparatus. Janin and Iwatsubo (64) have shown that the expected increase of the protein fluorescence corresponding to a T to R conformation change is detected during a fraction of a second after mixing the protein with aspartate or KCl; a t the same time, the absorption at 269 nm increases and the absorption a t 289 nm decreases. When the reverse R to T reaction is performed, the opposite spectroscopic effects take place and are completed within a few seconds of mixing with threonine. Each reaction can be characterized by a single exponential process and therefore contains only one pseudo-monomolecular step. Stopped flow and equilibrium measurements of conformation change agree therefore both qualitatively and quantitatively. Since a single exponential process accounts for most of the phenomena in the range of time accessible to fast-mixing techniques, the conformation change has only one rate determinant step. A mechanism involving a process of dissociation-reassociation of the protein is ruled out by the fact that variations of the protein concentrations do not change the rate constant. As expected, Janin and Iwatsubo found that the protein molecules with different amounts of bound ligands change conformation a t different rates. The effects of K+ ions on the rate of the R to T reaction are shown in Table V. Since the ligands act on the equilibrium and on the rate constant of its relaxation, this isomerization step must be coupled with the binding steps. An extension of the study to shorter times was performed using temperature-jump techniques. When the temperature of the protein solution is varied from 20" to 28" a large relaxation time R , ( 2 7 0 msec) of the fluorescence or of the light absorption at 265 or 295 nm is observed. The 53. E. D.Barber and H. J. Bright, Proc. N u t . Acud. Sci. U. S. 60, 1363 (1968) J. Janin nnd M . Iwatsubo, E m . J . Biochem. 11, 530 (1969).

54.

534

PAOLO TRUFFA-EACH1

EFFECTOF

THE

TABLE V CONCENTRATION OF K+ IONSON OF THE R TO T REACTION'

THE

RATE

Rate constanb with KCl (mW

0.5 mM

5 mM

50 mM threonine

(sec-l)

(sec-l)

(sec-l)

50 100

8.5 3.0 1.6 1.1

11.5

9.2

23 22

6.6 10.0

14 18

200 400

Data from Janin and Iwatsuho (64).

decrease of the fluorescence intensity expresses an increase of the relative concentration of the T form (54).The same experiments also demonstrate the existence of a shorter relaxation time R z ( 2 1 msec) with a sign opposite to that of R , in the fluorescence and absorption measurements. From studies on the temperature relaxation a t different concentrations of protein and by studies on the influence of K+ ion on the relaxation time, it was concluded that RI and R , correspond to monomolecular steps. These results indicate also that the relaxation of the R 2 T equilibrium proceeds through two isomerization steps and suggest the existence of a t least one conformation of the protein, S, rapidly equilibrating with one of the major conformation and slowly with the other. AddiTABLE VI EQUILIBRIUM AND RATE CONSTANTS OF COENZYME BINDINW~ Dissociation constants

Rate constants ka

Enzyme

Coenzyme

R form T form R form T form

NADPH NADPH NADH NADH

(pM-1

sec-1)

100

290 9 65

kd

kd/ka

K

(sec-l)

( P W

( PW

0.3 0.27 70 70

0.27 0.27

30 80 650 4500

>50 -

Data from Janin (66). The values of the association and dissociation rate constants k , and k d are derived from second-order plot3 of the rates of thermal relaxations in the presence of 0.15 M KC1 with (Tform) or without ( Rform) 0.6 mMIrthreonine. Thedissociation constants K are measured independently by fluorescence titration of the binding sites with NADPH. The affinity for NADH is low and can only be estimated by this technique. 0

15.

535

MICROBIAL ASPARTOKINASES

tional studies of Janin (55) on the relaxation phenomenon have permitted the determination of the equilibrium and rate constant for the R T conformation change. From the satne set of studies the equilibrium and rate constant of coenzyme binding were determined (Table VI) .

-

d. Circular Dichroic and Optical Rotatory Spectra ( 3 6 ) . The near ultraviolet circular dichroic (CD) spectrum of the protein is shown in Fig. 8. The spectrum is a complex overlap of negative bands. The band arises from aromatic transitions, although a t shorter wavelengths there may be a contribution from disulfide bonds. The effects of threonine binding, shown in Fig. 8, primarily result in a decrease in the magnitude of the ellipticity of the bands. The change of ellipticity was used to titrate the protein with threonine; this titration is sigmoidal and is a measure of the saturation function. NADPH binding produces an extrinsic Cotton effect a t 248 nm that was used to determine the amount of NADPH bound to the protein (see Section II,B,8,b). Optical rotatory dispersion (ORD) was used as well as the CD spectra to calculate the a-helical content of the protein. Depending on the method used for the calculation, the a-helical content varies from 17 to 31%. This difference indicates that types of structure other than the a-helix are present in the protein and that a t least a part of structure is in the form of antiparallel pleated sheet. This hypothesis is indeed supported by the observation that deep ultraviolet circular dichroic

260

280

300

320

X , nm

FIG.8. Near ultraviolet circular dichroic spectra of aspartokinase I-homoserinc tlehydrogenasr 1:free enzyme: A, enzyme 2 mM L-threonine: B, enzyme 20 m M potassium baspartate : C ( 3 6 ) .

+

55. J. Janin, Cold Spritig Harbor Symp. Quant. Biol. 36, 193 (1971).

+

536

PAOLO TRUFFA-BACHI

trough is maximally intense a t about 218 nm, a feature characteristic of the antiparallel P-pleated sheet conformation. The effect of 6 M guanidinium chloride on the protein CD spectra and ORD was studied. Contrary to the results obtained by Tanford with certain proteins (56), aspartokinase I-homoserine dehydrogenase I is not entirely random coiled until the disulfide bonds are reduced. Heck and Truffa-Bachi suggested that this anomaly could be explained by the presence of an aromatic residue in the proximity of a disulfide bridge. Subsequent studies by Falcoz-Kelly et al. ($9) verified the existence of a peptide which contains a tryptophan residue between two cysteines involved in a disulfide bridge.

11. Distribution of the Two Activities on the Polypeptide Chain Mild proteolysis of aspartokinase I-homoserine dehydrogenase I results in the production of a homoserine dehydrogenase fragment whose subunit corresponds to the C-terminal part of the native polypeptide chain (57). It is difficult to decide whether proteolysis occurs first in a preferential region, giving rise to two fragments the first of which is further degraded, or if the N-terminal section of the polypeptide is randomly hydrolyzed. The proteolysis by a-chymotrypsin or trypsin leads to desensitization of homoserine dehydrogenase activity whereas the kinase activity is completely destroyed. The products of proteolysis analyzed on SDS gels showed a band of protein corresponding to a species of molecular weight 55,000. I n the presence of threonine the two enzymic activities are fully protected, and no difference with the native enzyme can be detected. The proteolytic fragment was purified on a Sephadex G-200 column and a species with a molecular weight of 110,000 can be isolated; it represents a dimer of the 55,000 molecular weight species. The C-terminal amino acid of this fragment was found to be same as in the native enzyme, whereas the N-terminal amino acid was found to be different. When homoserine dehydrogenase fragment was reduced and carboxymethylated with [ 14C]iodoacetic acid, only three radioactive peptides were detected (57) (Fig. 9 ) . A protein carrying only the sensitive aspartokinase activity had already been purified from a mutant. The kinetic parameters and the cooperative inhibition by threonine were very similar to those obtained 56. C. Tanford, Advan. Protein Chern. 23, 121 (1968). 57. M. Veron, F. Falcoz-Kelly, and G . N. Cohen, Eur. J . Biochem. 28,5-20 (1972).

15.

537

MICROBIAL ASPARTOKINASES

for the wild type protein (58). The mutant protein appears to retain, in the presence of threonine, the tetrameric structure of the wild type protein but has a considerably shortened polypeptide chain. This mutation cannot be a deletion since revertants with normal homoserine dehydrogenase activity are easily obtained. Therefore, the most likely explanation is that the mutation leads to the interruption of the reading of the gene and hence to a polypeptide chain shortened on its carboxyl terminal side.

+

Electrophoresis at pH 6.5

-

FIG.9. Fingerprint of the homoserine dehydrogenase fragment. Pure homoserine dehydrogenase fragment was obtained upon papain digestion of aspartokinase I-homoserine dehpdrogenase I. The figure shows the autoradiograph of the fingerprint of the soluble tryptic peptides obtained from the [l*Clcarboxymethylated fragment. The only radioactive spots detected are 'those corresponding to peptides T 60,T 30, and T Na (57). 58. J . Janin, P. Truffa-Bachi, and G. N. Cohen, Biochem. Biophys. Res. Commun. 26, 429 (1967).

538

PAOLO TRUFFA-BACHI

This conclusion is strengthened by chemical studies of the homoserine dehydrogenase fragment and of the mutant protein. The latter was shown to have a methionine N-terminal residue as in the wild type protein and, in contrast, preliminary results indicate that its C-terminal sequence is different from that of the wild type enzyme. Moreover, the mutant protein contains only three of the six soluble carboxymethylcysteinecontaining peptides normally present on the fingerprint of the wild type protein; these peptides (T85, T10, and TNb) are precisely those missing in the homoserine dehydrogenase fragment which contains the carboxyl terminal part of the polypeptide chain (Fig. 10). This confirms that the aspartokinase activity resides in the aminoterminal part of the subunit. Although the subunits of the homoserinc dehydrogenase fragment and of the mutant protein share a common sequence (the comparison of the molecular weights of the subunits indeed indicates an overlap of about 17,000 daltons), the two catalytic activities are likely to be carried by two distinct regions of the polypeptide chain. The aspartokinase activity is associated with the amino terminal and the homoserine dehydrogenase with the carboxyl terminal section of the polypeptide chain. The fact that such modified proteins with shortened subunits can be obtained, either by mutation or by mild proteolysis, gives some further indications regarding the configuration of native aspartokinase I-homoserine dehydrogenase I. Both fragments retain, almost unchanged, one or the other of the two catalytic activities and a t least some of the association areas between subunits. Therefore, as proposed for P-galactosidase and its complemented derivative (44), their structure must be close to that of the corresponding region in the native protein. The subunit of the aspartokinase I-homoserine dehydrogenase I appears in its native configuration to be composed of a t least two rather independent regions each being able to retain its own catalytic activity when the other is removed, either by mutation or mild proteolysis. Since the protein from the mutant Gif 108 retains a tetrameric structure, it is likely to contain the association areas found in the native enzyme. In the proteolytic fragments this region is presumably lost, but another region in which protein-to-protein interaction exists is retained as shown by the fact that the fragment is obtained as a dimer. Further structural studies may show whether the subunit interactions found in both fragments are similar to those found in the native enzyme. A bifunctional protein has been artificially produced by the fusion of two adjacent genes of the histidine operon (4W). The bifunctional protein obtained has been subjected to mild proteolysis (59),and it was 59.

T.Kohno and J. Yourno, J . Biol. Chem. 246, 2203 (1971).

15.

MICROBIAL ASPARTOKINASES

539

FIG.10. Fingerprint of the aspartokinase mutant protein from Gif 108 (67).Top: Autoradiograph of the [“C3carboxymethy1cysteine-containing peptides from the wild type (A) and mutant protein (B) separated in the first dimension electrophoresis. Bottom : Schematic pattern of the fingerprint obtained from the [‘4Clcarboxymethylated protein. The ninhydrin peptides are open areas while the dark areas correspond to the radioactive peptides revealed by autoradiography : T85, T10, and TNb.

observed that one of the activities is destroyed while a proteolytic fragment similar to one of the components of the fused protein is released. Gene fusion has been proposed as an important mechanism in the evolution of complex proteins (43) and aspartokinase I-homoserine dehy-

540

PAOLO TRUFFA-BACHI

drogenase I may have arisen by such a process. This hypothesis is strengthened by the finding that in all other bacteria studied, excluding the Enterobacteriaceae, the two activities, aspartokinase and homoserine dehydrogenase, are carried by two independent polypeptide chains.

C. ASPARTOKINASE 11-HOMOSERINE DEHYDROGENASE I1 Methionine-repressible aspartokinase and homoserine dehydrogenase exist in E . coli K12 at a very low level which precludes their study and, in fact, they were demonstrable only in a mutant devoid of the corresponding threonine-sensitive activities. From this organism, a mutant constitutive for aspartokinase I1 and homoserine dehydrogenase I1 was isolated. A preliminary study (22) definitely indicated the possibility that the two activities could be carried by a single protein as has been found for the two threonine-sensitive activities. Definite proof that the same protein carries both activities was subsequently given ( 6 0 ) . 1. Pirrificution and Criteria of Homogeneity The purification of the enzymc involves a two ammonium sulfate precipitation, a chromatography on DEAE-Sephadex, another ammonium sulfate precipitation, and a centrifugation. The aspartokinase and homoserine dehydrogenase activities remain associated throughout a 400-fold purification. The freshly prepared enzyme gives a single, sharp, and symmetrical peak in the ultracentrifuge. Electrophoresis on polyacrylamide gel gives two closely migrating bands that both show homoserine dehydrogenase activity. A charge differencc appears responsible for this double band. 2. Stability of the Enzy,ine

When stored at -15" in buffer containing 20% glycerol, the enzyme was stable, remaining 100% active and homogeneous for several months. However, when kept in solution at O", the enzyme tended to aggregate and to become heterogeneous with loss of free -SH groups. The aggregation process may thus involve the formation of intermolecular disulfide bridges. 60. F. Falcoe-Kelly, R. van Rapenbusch, and G. N. Cohen, Eur. J . Biochem. 8, 146 (1969).

15.

MICROBIAL ASPARTOKINASES

541

3. Extinction Coefficient The extinction coefficient a t the absorption maximum 282 nm is 0.87 absorbance unit/cm2/mg.

4. Molecular Weight of the Native Enzyme The value for sedimentation coefficient obtained upon linear extrapolation to infinite dilution was ~ 2 0 = , ~7.6 ( f 0 . 2 ) X 10-l3 cm. From the equilibrium sedimentation experiments a molecular weight of 169,000 2 9,000 can be calculated.

5. Subunit Structure Reduced guanidine-denaturated aspartokinase 11-homoserine dehydrogenase I1 protein travels as a single, symmetrical peak which appears monodisperse in schlieren optics throughout the centrifugation a t the five protein concentrations used. The molecular weight of the subunit, calculated from equilibrium runs, was found to be 43,000 f 5,000 daltons. In conclusion, aspartokinase 11-homoserine dehydrogenase I1 of E . coli K12 is composed of subunits of molecular weight 43,000 k 5,000. Since the niolcculnr weight of the nativc protcin is 169,000 & 9,000, it contains four subunits of equivalcnt molecular weight. The number of peptides obtained by tryptic treatment of thc carboxymethylated protein is compatible with four identical subunits.

6. Kinetic Parameters The apparent dissociation constant of the substrates of the homoserine dehydrogenase I1 are the following: K , = NADPH 1.5 X M, NADPH 3.2 X L-aspartatc p-semialdehyde 1.9 X lo4. For the M and aspartokinase I1 the values found are R, ATP = 1.9 X I<,,, L-aspartate = 2.1 X The molecular specific activity a t 27" of the enzymes has been calculated to be 4000 moles of aspartate semialdehyde reduced per minute per mole of dehydrogenase and 850 moles of aspartate phosphorylated per minute per mole of aspartokinase. The molecular specific activities are lower than those reported for the threoninc-sensitive multifunctional enzyme.

7. Amino Acid Composition The amino acid composition of the carboxymethylated protein is given in Table VII.

542

PAOLO TRUFFA-BACHI

TABLE VII AMINO ACID COMPOS~TION OF ASPIIITOKIN.WE IIHOMOSERINE DEWYDROQENASI~: 11.

a

Amino acid

Residues per 169,000 daltons

Lysine Histidine Arginiiie Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalaiiine Tryptophan

46 34 91 156 61 128 172 50 146 162 20 125 17 56 186 33 53 20

Data from Falcoz-Kelly ct al. (60).

D. ASPARTOKINASE 111 1. Purification and Criteria of Homogeneity The purification of thc enzyme involves aininonium sulfate precipitation, DEAE-Sephadex chromatography, Sephadex G-200 column, and hydroxyapatite chromatography (61). Purity was checked by ultracentrifugation and disc gel electrophoresis. 2. Extinction Coefficient

The extinction coefficient had been found to be 0.36 absorbance unit X cm'/mg. However, a new determination based on amino acid analysis gave a value of 0.46; the samc value was also found by refractometry (J. C. Patte, personal communication),

61. G. Lafuma, C. Gros, and J. C. Patte, Ew. J . Bioclte,ti. 15, 111 (1970).

15.

543

MICROBIAL ASPARTOKlKkCjES

TABLE VIII AMINOACID COMPOSITION OF ASPAILTOSINASE IIIa

Amino acid Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Phenylalanine Tyrosine Tryptophan a

Residues per 130,000 daltons 40 22 76 112 76 80 133 44 98 134 20 99 21 67 166 46 16

7

Data from Lafuma et al. (61).

3. Moleczilnr Weight Thc value for the sedimentation coefficient obtained upon linear extrapolation t o infinite dilution is s ~ , , = , ~ 6.9 ( f 0 . 1 5 ) X 10-13 sec. From equilibrium sedimentation runs the molecular weight was calculated to be 127,000 k 7,000 daltons. 4. Kinetic Parameters

The apparent dissociation constants of the substrates of the aspartokinase I11 arc I<,?,ATP = 4.8 X 10-3M; IC, aspartate = 4.7 X 10-3M. The molar specific activity a t 27" has been calculated to be 2350 moles of aspartatc phosphorylated per minute ( 6 2 ) .

5. Amino Acid Composition The amino acid composition of aspurtokinase I11 is giveii in Table

VIII. 62. P. Truffu-Bachi and G. N. Colien, Biochint. Bwphys. Actn 113, 531 (1966).

544

PAOLO TRUFFA-BACHI

6. Inhibition

The inhibition by lysine of the aspartokinase I11 shows a sigmoidal curve typical of allosteric enzymes (63). When other amino acids were tested as inhibitors it was found that leucine, isoleucine, and phenylalanine a t high concentration also affected the aspartokinase activity (64). Simultaneous addition of noninhibitory amounts of lysine and low amounts of leucine, isoleucine, or phenylalanine results in a higher inhibition than expected, i.e., there is a synergistic inhibition in the presence of lysine and of the other amino acids. Patte et al. (64) suggested the existence of a site for the lysine binding and one or more sites for the other nonpolar amino acids. This phenomenon was also described in Rhodopseudomonas capsulatus (see Section IV,A) . 111. Ofher Coliform Bacteria

Isofunctional aspartok-inases were also found in Salmonella typhimurium; Freundlich (20) has shown that a lysine-sensitive aspartokinase, a threonine-sensitive aspartokinase, and a threonine-sensitive homoserine dehydrogenase exist in these bacteria. He later showed in the same bacteria a homoserine dehydrogenase repressible by methionine (66). Cohen et al. (66),analyzing a large number of strains of coliform bacteria, have shown that isofunctional aspartokinases with allosteric properties of inhibition by lysine and threonine like those of E . coli arc universal in this genus, and that with the possibly exception of Edwardsiella and Providencia all the strains examined contain a fraction of nonsensitive enzyme which reflects the existence of a methionine-repressible enzyme. IV. Aspartokinaser Regulated by Concerted Feedback Inhibition

A. Rhodopseudomonas capsulatus The studies of Datta and Gest on the aspartokinssc of this photosynthetic bacterium resulted in the discovery of a phenomenon which 63. J. C. Patte and G. N. Cohen, C. R. Acad. Sci. 259, 1255 (1964). 64.

J. C. Patte, T. Loviny, and G. N. Cohen, Biochirn. Biophys. Actn 99, 523

(1965). 65. R. L. CafTerata and M. Freundlich, J. Bacterial. 97, 193 (1969). N. Cohen, R. Y . S t a n k , and G. Le Bras, J. Bacterial. 99, 791 (1969).

66. G.

15.

545

MICROBIAL ASPARTOKINASES

provided a basic element for one control scheme, viz., feedback inhibition by simultaneous action of two amino acids ; the authors designated this phenomenon “concerted feedback inhibition” (67). One single aspartokinase exists in Rhodopseudoinonas capsdatus; its activity is in: hibited to the extent of 15-20% at a concentration of 1 mM by L-threonine and L-lysine alone. The aspartokinase is, however, totally inhibited by addition of both amino acids (Fig. 11). By a series of experiments involving the protection of the enzymic avctivity against heat inactivation, the authors concluded that distinct regulatory sites exist for the two amino acids. This absolute requirement for two end products to accomplish efficient inhibition seems to be a less delicate control than the one using isofunctional enzymes since it does not allow the independent regulation of the first reaction in a branched pathway. It represents however an effective mechanism and a different control from the one we have seen in Enterobacteriaceac. ’

B. OTHERNONSULFUR PHOTOSYNTHETIC BACTERIA Rhodospirillum rubrum possesses one aspartokinase that was reported to be sensitive to threonine. No concerted feedback inhibition was found (68).I n contrast to these results, Cohen et al. (66) reported a concerted feedback inhibition by L-lysine plus L-threonine. The aspartokinase of Rhodospirillum tenue was purified 60-fold; this activity can be readily separated from the homoserine dehydrogenase activity (69). An apparent molecular weight of 100,OOO was found for the aspartokinase by gel filtration on a Sephadex G-200 column. The kinetic constants are I<,, aspartate = 9 X lO-‘M; I(, ATP = 3 X 10-3 M . The aspartokinase activity can be completely inhibited in a cooperative way by lysine or by threonine. The activity can also be inhibited by concerted feedback inhibition by low concentration of the two amino acids together. The main features of the aspartokinase from R. tenue are: (1) an additional, and so far unique, concerted feedback inhibition by L-threonine plus L-mcthioninc and (2) thc release of lysine inhibition by addition of methionine, isoleucine, glycine, or phenylalanine. Lysine, threonine, or methionine protect the enzymic activity against heat inactivation (69). 67. P.D a t t and H . Gest, Proc. N a f . Acad. Sn’. U.S. 52, 1004 (1966). 68. P.Datta and H. Gest, Nature (London) 203, 1259 (1964). 69. M. Robert-Gero, L. Le Borgne, and G . N . Cohen, J. Bacterial. 112, 251 (1972).

PAOLO TRUFFA-BACHI

546 100

*

c

-0 - 2 m M ~ - L y a - k(~-Lya)+2mM L-Thr-

80-

>

0

8o

10

20

t

40

50

/No threonine

+ 1 mM L-threonine

I: 2 0 01 0

30

I

2

I

I

I

6 8 L-Lysine concn. ( m M )

4

I

10

(6)

FIG.11 (A) Effects of L-lysine plus ~-threonineon R. cnpsulutits aspartokinase xctivity. Two solutions, each containing 0.75 m g protein per ml of standard reaction mixture, were incubated ~ i t or h without 2 niM lysine as indicated, n t 26'. At the times denoted by the points, 1 ml samples were removed and the quantities of nspnrtohydroxnmate estimated. At 18 minutes (arrow) both solutions were supplemented with ~-threonine(2 m M ) . (B) Progressive inhibition of aspartokinase activity with increasing concentration of L-lysine at a constant level (1 mM) of ~-threonine(67).

C . Bacilli 1. Bacillus polymyxa

a. Inhibition Studies. In these bacteria only one aspartokinase is present. Its activity is enhanced by ammonium or potassium ions. At 37" the requirement for these ions is nearly absolute; a t 25" it is less stringent. The enzyme is strongly inhibited in a concerted manner by addition of L-threonine and L-lysine a t concentrations below 1 mM and is also

15.

MICROBIAL ASPARTOKINASES

547

inhibited by higher concentrations of each amino acid alone. No effect of methionine was found (70, 7 1 ) . The inhibitors behave differently as a function of the temperature of the assay: While a t 25" the feedback inhibitors lead to a reduction of V,,,,,, at 37" the inhibition also reduces the affinity of the enzyme for L-aspartate. The K , for aspartate was also found to be dependent, a t 37", on enzyme concentration suggesting that a t 37" (but not at lower temperature) the active form of the aspartokinase dissociates to a lower molecular form which has a markedly lower affinity for aspartate. The effects of dioxane on the affinity of aspartate support this hypothesis. Nonpolar L-amino acids protect the aspartokinase from inactivation by heat and detergent (72). Since the amino acids protect the aspartokinase under conditions where L-aspartate is ineffective but are devoid of protective activity under conditions where aspartate protects, Paulus and Gray concluded that the stereospecific site(s) for nonpolar amino acids was distinct for the active center of the enzyme. The same nonpolar amino acids also reverse the inhibition of the aspartokinase caused by the feedback inhibitors L-lysine and L-threonine. These nonpolar amino acids differ in their specificities: Amino acids of group I (L-tryptophan, L-methionine, and L-isoleucine) counteract the inhibition by L-lysine only ; amino acids of group I1 (L-leucine, L-valine, L-isoleucine, and L-norvaline) counteract the inhibition by L-threonine only; finally, amino acids of group I11 (L-phenylalanine, L-alanine, glycine, and allyglycine) counteract the concerted inhibition by L-lysine plus L-threonine. Cooperative interaction does not occur between the amino acids of the different groups. From the prediction of the allosteric model (60) the authors concluded that all the activating ligands bind to the same site of the enzyme, this site being different from the threonine and lysine sites since cooperative interaction is observed between activators of group I and L-threonine and activators of group I1 and L-lysine (7s). A four-state model based on the kinetic properties was proposed for the interactions of activators and feedback inhibitors (73).

b . Enzyme Purification, Molecular Weight, and Amino Acid Composition (73).The aspartokinase of B. polymyxa was purified 1300-fold with 24% yield, Homogeneity was checked by gel electrophoresis and equilibrium centrifugation. The enzyme can be stored at -10" with a 4Q% loss of activity over a period of 6 months ; longer storage does not lead to further inactivation. A gel electrophoresis of an aged solution of enzyme shows the appear70. H.Paulus and E. Gray, J . Biol. Chem. 239, 4008 (1964). 71. H.Paulus and E. Gray, J . Biol. Chem. 242, 4980 (1967). 72. H.Paulus and E. Gray, J . Biol. Chem. 243, 1349 (1968). 73. C.Biswas, E. Gray, rind H. Paulus, J . Biol. Chem. 245, 4900 (1970).

548

PAOLO TRUFFA-BACHI

TABLE IX AMINOACIDCOMPOSITION OF THE ASPARTOKINASE OF Bacillus polymyxaa

Amino acid

Residues per 116,000 daltons

Lysine Histidme Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glyche Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalmine Cysteine Tryptophan

52 20 47 90 59 73 139 24 97 124 128 29 66 81 19 27 7 9

Data from Biswas et al. (73).

ance of a band of protein migrating more rapidly than the native enzyme. Addition of MgC1, to the electrophoresis buffer gives the pattern obtained with fresh enzyme. From equilibrium ultracentrifugation studies, the molecular weight of the enzyme was found to be 116,000 daltons. The amino acid composition is given in Table IX.

c. Subunit Structure. Gel electrophoresis in SDS reveals the presence of two types of subunits: one with a molecular weight of 47,000 daltons and the other with a molecular weight of 17,000.Gel filtration in SDS on Sephadex G-200 columns results in a complete separation of the two types of subunit. Biswas et al. (73) suggcsted that the native protein is composed of two subunits of each type. 2. Bacillus subtilis

Bacillus subtilis possesses two distinct aspartokinases. They have been separated by chromatography on a Sephadex G-100 column ( 7 4 ) . Table 74. A. Rosncr and H. Paulus, J . Biol. Chem. 246, 2965 (1971).

15.

549

MICROBIAL ASPARTOKIK AGES

TABLE X CQMPARISON OF ASPARTOKINASE 1 AND 11 OF B. subtilisa Property Molecular weight Monovalent cation requirement, pH range for 50% activity Apparent K , for Laspartatel Inhibitor Kinetics of inhibitionc Shape of inhibition curveb Effect, of pH on inhibition Effect of nonpolar amino acids Enzyme level during growth cycle Enzyme level in rich mediumd

Aspartokinase I

Aspartokinase I1

250,000

125,000

Not. specific

K+ or NH4+

6.0-9.5 3 mM

6.5-8.2

m-Diaminopimelate Noncompet it ive Hyperbolic Less a t low pH No Constant 70%

17 mM LThreonine and Llysine Competit,ive Sigmoid Less a t high pH Yes Decreases 10%

Data from Rosner and Paulus (74). Under standard assay condit.ions at, 1 mM ATP. With respect to L-aspartate. Relative to level in exponential cultures in minimal medium.

X shows the differences between the two enzymes. Aspartokinase 11, which is subject to a concerted feedback inhibition by L-lysine and L-threonine, resembles the aspartokinascs isolated from other organisms (see above). The activity of the aspartokinase I1 is also modulated by nonpolar amino acids as the aspartokinase 111 of E . coli (64) and the unique aspartokinasc of B. polyinyza ( 7 2 ) . The properties of aspartokinase I arc very different in being inhibited by an intermediate of the biosynthetic pathway, namely, diaminopimelic acid, a lysine precursor. The synthesis of the two enzymes varies with culture conditions: I n rapidly growing cultures, 80% of the total aspartokinase is represented by asllartokinase 11; in stationary culture or in rich media this enzyme is repressed 5- to 10-fold. These observations, as well as the nature of the end product regulations, suggest that the essential, physiological role of aspartokinase I1 is to supply precursors for the amino acid pool. A possible role for aspartokinase I would be to ensure the synthesis of diaminopimelate when the amount of aspartokinase I1 is repressed such as in rich media containing an excess of lysine and threonine ( 7 4 ) . In addition, aspartokinase I may contribute significantly to the synthesis of dipicolinic acid during sporulation, under conditions where the aspartokinase I1 is largely nonfunctional. The decline of the specific activity of the aspartokinase I1 is too rapid to result only from dilution of the enzyme during growth. Three hy-

550

PAOLO TRUFFA-BACHI

potheses have been envisaged by the authors: (1) instability, (2) specific degradation, and (3) conversion of aspartokinase I1 to I. The latter hypothesis would be consistent with the observation that the total aspartokinase activity remains constant throughout the bacterial growth (74). 3. Bacillus stearotherinophilus

The aspartokinase from this species has been partially purified and characterized (75). The temperature optimum is near 55", optimal p H is 8. The molecular weight determined by filtration on a Sephadex G-100 was found to be 110,000 daltons. This aspartokinase is inhibited in a concerted manner by L-lysine and L-threonine; each amino acid alone can totally inhibit the activity a t a concentration of 1 mM. The sensitivity of the enzyme to threonine or to lysine decreases as the assay temperature is increased from 23" to 55" ; however, this effect is more pronounced with respect to threonine sensitivity. In contrast to this result, the sensitivity of the enzyme to concerted feedback inhibition, at 55", is comparable to that found at much lower temperatures for aspartokinases of mesophilic bacteria. Lysine and threonine singly or together act as noncompetitive inhibitors with respect to ATP and as mixed competitive noncompetitive inhibitors with respect to aspartate. The absence of a competitive relationship between each inhibitor and aspartate was explained by Kuramitsu (75) as the result of either separate regulatory sites for lysine and threonine or by inhibitor sites which partially overlap the aspartate site (increasing the KuLfor aspartatel and also affect that portion of active site iiivolved in catalysis subsequent to substrate attachment (decreasing the V",,, of the reaction), Separate regulatory sites appear to be more likely since the enzyme can be desensitized to lysine and threonine under conditions where enzymic activity is increased. Homotropic interactions could not be shown for either substrate in the presence or in the absence of inhibitors; however, homotropic interactions with respect to threonine were observed as the assay of temperature was increased. No such effect was shown for lysine. Heat inactivation studies showed that the stabilization of the enzyme requires both inhibitors, suggesting a conformational change of the enzyme structure. Such a conformational change does not involve association-dissociation phenomena since no detectable alteration in sedimentation was observed following gradient sedimentation in the presencc of both inhibitors ('75). 75. H.

I<. Kurnmitsu, J. B i d . Chem. 245, 2991 (1970).

15.

MICROBIAL ASPARTOKINARES

55 1

4. Other Bacilli The aspartokinase of B. licheniformis is inhibited by lysine or by aspartate-p-semialdehyde (76), a fact reminiscent of the aspartokinase discovered in Rhodopseudonzonas spheroides (77). Addition of lysine plus threonine, as in other Bacilli, shows a concerted feedback inhibition. A combination of lysine and aspartate-p-semialdehyde also results in a concerted inhibition of the activity. The enzyme can be inactivated by storage in the cold and loses the capacity of being inhibited by lysine while its sensitivity to aspartate-p-semialdehyde remains unchanged ('76). In Bacillus cerew the only aspartokinase detected is totally sensitive to lysine (78). The sensitivity of the enzyme to lysine depends on the age of the culture, and the results indicate that a t the moment of the synthesis of dipicolinic acid the aspartokinase present is feedbackinsensitive (78).

D. PSEUDOMONADS In fluorescent pseudomonads (P. aeruginosa, P. putida, and P. Fuorescens), the single aspartokinase can be inhibited either by addition of large amounts of lysine or threonine or by lower concentration of the two amino acids together. Methionine has no effect (66). The aspartokinase of P. putida was studied in more detail by RobertGero et at. (79). I t was shown that the inhibition curves of lysine or threonine are cooperative. Both allosteric effectors acted as mixed competitive-noncompetitive inhibitors toward aspartate. Concerted inhibition was, however, noncompetitive. The affinity constants are I<, aspartate M and Knl ATP = 4.35 X M. = 4.8 X The molecular weight of the enzyme was estimated to be 126,000 by gel filtration 011 a Sephadex G-100 column. The aspartokinase is severely but not totally repressed when the bacteria is grown on culture media containing either lysine, methionine, or threonine. The enzyme synthesized under these conditions is different from that obtained from culture grown on ininimal medium: The sensitivity to lysine and threonine is lost. This result may appear as a physiological self-defense mechanism 76. D. P. Stahley and R. W. Bernlohr, Biochim. Biophys. Acta 146, 467 (1967). 77. P. Datta and L. Prakash, J. Biol. Chem. 241, 5827 (1966). 78. A. I. Aronson. E. Henderson, and A. Tincher, Biochem. Biophys. Res. Cornmiin. 26, 454 (1967). 79. M. Robert-Gero, M. Poiret. and G. R. Cohen, Biochim. Biophys. Actn 206, 17 (1970).

552

PAOLO TRUFFA-BACHI

for a cell which does not have isofunctional enzymes (79)to prevent the growth inhibition which would otherwise occur in the presence of an excess of a single amino acid that would act as a repressor and a feedback inhibitor. In the acidovorans group, lysine or threonine do not inhibit alone but exert a feedback inhibition when added together (66).

E. OTHERGENERA Concerted feedback inhibition of aspartokinase is also shown by Micrococcus glutamicus and Brevibacterium flavum (80, 81 ) . I n the latter, lysine or threonine alone do not inhibit; the concerted feedback inhibition exerted by these two amino acids can be released by L-isoleucine or L-valine (81). All the strains of Azotobacter investigated possess a single aspartokinase totally inhibited by high concentrations of lysine or threonine. Concerted feedback is shown a t low concentration by these two amino acids together. No inhibition or activation was observed with the two other end products of the pathway. Inhibitions by lysine and threoninc are highly cooperative. Both inhibitors are noncompetitive with respect to either substrate, ATP, and aspartate (89).

V. Rhodopseudomonos spheroides

The aspartokinase of these bacteria has been purified 240-fold by Datta and Prakash (77). The activity is totally insensitive to feedback inhibition by all the end products of the synthetic pathway, either alonc or in combination. However, the activity is strongly inhibited by aspartate-p-semialdehyde, a key intermediate for the synthesis of all the amino acids of the pathway. This inhibition is competitive toward both substrates. From kinetic experiments it was concluded that the enzyme possesses one binding site for aspartate and two sites for ATP. No inhibition has been found on the aspartokinase activity by lysine and threonine; however, both amino acids alone protect to a certain extent against heat inactivation (77). One explanation is that binding of threo80. K . Nakayana, H. Tanaka, H. Ogiwara, and S. Kinoshita, A g r . Biol. Chem. 30, 849 (1966). 81. R. Miyajina, S. Otsuka, and I. Shiio, J . Biochem. (Tokyo) 63, 139 (1968). 82. M. Robert-Gero, J. M. Sala-Trepat, and L. Le Borgne, J. Gen. Microbiol. 67, 189 (1971).

15.

553

MICROBIAL ASPARTOKINASES

nine or lysine does not influence the specific conformation required for catalytic functions.

0.f

the enzyme

.

VI Saccharornyces cerevisiae

The enzymic phosphorylation of L-aspartate was first discovered by Black and Wright in yeast extracts (If). The product of the reaction, p-aspartyl phosphate, was synthesized and its chemical properties were studied. The enzymic reaction was studied in both forward and reverse direction; in the reverse reaction the formation of ATP and ADP and p-aspartyl phosphate is stoichiometric. The equilibrium of the reaction was determined (see Section 1,B). The yeast aspartokinase was found to be fully sensitive t o threonine (19). ACKNOWLEDGMENTS The experimental work from our laboratory described in this review has been supportrd by the Centre National de la Recherche Scientifique, the DClCgation GCnCrale it la Recherche Scientifique et Technique, the Fondation pour la Recherche Medicale, and the Commissariat It 1’Energie Atomique.

This Page Intentionally Left Blank

Protein 1%inases DONAL ,4.WALSH

EDWIN G. KREBS

. .

.

. . . . . .

I. Introduction . . . . . . . 11. Substrate-Specific Protein Kinases . . . . . A. Phosphorylase Kinase . . . . . . . B. Pyruvic Dehydrogenase Kinase . . . . . 111. Cyclic Nucleotide-Regulated Protein Kinases . A. Cyclic AMP-Dependent Protein Kinases . . . B. Other Cyclic Nucleotide-Regulated Protein Kinases IV. Nonrlassified Protein Kinases . . . . . . A. Histonc Kinases . . . . . . . . B. Acidic Nuclear Protein Kinasrs . . . C. Phosvitin Kinases . . . . . . .

. .

. .

.

. .

. .

. .

. . . .

.

.

. .

. . . . . .

555 557 557 565 566 566 578 578 579 580 580

I. Introduction

The term ‘))rotein kinaee” was first used to describe an enzyme detected in liver, brain, and yeast which catalyzes the phosphorylation of casein or phosvitin, phosphoproteins that serve as nutrients in animals or in the developing embryo, respectively (1-3). This type of protein kinase was believed to be relatively nonspecific but nonetheless favored as substrates those proteins having a high content of serine, often containing runs of four or more serines in a row (4, 5 ) . Protein kinases 1. G. Burnrtt and E. P. Kennedy, JBC 211, 969 (1954). 2. T. A . Sundararajan, K. S. V. Sampath-Kumar, and P. S. Sarma. BBA 29, 449 (1958). 3. M. Rabinowitz and F. Lipmann, JBC 235, 1043 (1960). 4. S. Posternak, C . R. Acad. Sci. 184, 306 (1927). 5. J. Williams and F. Sanger, BBA 33, 294 (1959). 555

556

DONAL

A. WALSH

AND EDWIN G . KREBS

possessing more discrete functions have also been recognized and have generally been referred to on the basis of the particular substrate on which they act. In this latter category are phosphorylase kinase (6, 7 ) , glycogen synthetase I kinase (8),liistone kinase ( 9 ) , protamine kinase ( l o ) ,and pyruvic dehydrogenase kinase (11). Noteworthy of this latter group is their involvement in the regulation or control of various aspects of cellular function or metabolism. As indicated below, not all of these cnzymes are distinct entities. In 1968 ( l a ) ,it was recognized that certain protein kinases are stimulated by adenosine 3’,5’-monophosphate (cyclic AMP), a finding that was foreshadowed by knowledge of the effect of cyclic AMP on the activation and inactivation of phosphorylase kinase and glycogen synthetase, respectively (13-16). Because of the great interest in cyclic AMP as a rcgulatory agent, work on protein kinases has expanded enormously since that time, and emphasis is now being placed on the activatability or nonactivatability of a given enzyme by the cyclic nucleotide. No good system for classifying the protein kinases, which takes into account their specificities as well as their activation properties, has as yet emerged. To what extent the enzymes described prior to 1968 are identical to those now classified as cyclic AMP-dependent protein kinases is difficult to ascertain. Since 1968 protein kinases have been categorized on the basis of bcing either cyclic AMP-dependent or cyclic AMP-independent. This classification is ambiguous however as an expression of molecular and functional propertics. As discussed in detail below, some cyclic AMP-independent kinases occur by dissociation of cyclic AMPdependent enzymes. Although simple criteria are now available to distinguish this latter type of cyclic AMP-independent enzyme from those whose activities are definitely not regulated by cyclic AMP, these have in general not been applied. Because protein kinases have only recently been thrust into a position 6. E. G. Krebs and E. H. Fischer, BBA 20, 150 (1956). 7. T.W. Rall, E. W. Sutherlnnd, and W. D. Wosilait, JBC 218, 483 (1956). 8. D. L. Friedman and J. Larncr, Biochemistry 4, 2261 (1965). 9. T. A. Langan, in “Regulatory Mechanisms for Protein Synthesis in Mammalian Cells” (A. San Pietro, M. R. Lamborg. and F. T. Kenney, eds.), p. 101. Academic Press, New York, 1968. 10. B. Jrrgil and G. H. Dixon, JBC 245, 425 (1970). 11. T. C. Linn, F. H. Pettit, and L. J. Reed, Prim Nut. Acnd. Sci. U . S. 62, 234 (1969). 12. D. A. Walsh, J. P. Perkins, and E. G. Krebs, JBC 243, 3763 (1968). 13. E. G. Krebs, D. J. Graves, and E. H. Fischer, JBC 234, 2867 (1959). 14. J. Lamer, C. Villar-Palasi, and D. J. Richman, A B B 86, 56 (1960). 15. E. Belocopitow, ABB 93, 457 (1961). 16. D. L. Friedman and J. Lamer, BiochemistTy 2, 669 (1963).

16.

557

PROTEIN KINASES

of prominence, comparatively few review articles on this subject have appeared. The reader is referred, however, to an article by Rabinowitz in an earlier edition of this series (17) and to a recent review by one of the present authors (18). Reviews covering aspects of specific protein phosphorylation involved in glycogen metabolism are also available (19,2 0 ) . The present chapter will be concerned with (a) phosphorylase kinase and pyruvate dehydrogenase kinase currently classified as substrate-specific protein kinases ; (b) cyclic nucleotide-regulated protein kinases ; and (c) nonclassified protein kinases, i.e., enzymes about which little information is available concerning their specifioity or activatability by cyclic AMP.

II. Substrate-Specific Protein Kinases

A. PHOSPHORYLASE KINASE 1. Skeletal Muscle Phosphorylase Kinase

Because most of the work on skeletal muscle phosphorylase kinase has been carried out using rabbit muscle as the source, all specific information in this section will refer to that enzyme unless indicated otherwise. n. Reaction Catalyzed and Its Metabolic Significance. Phosphorylase kinase catalyzes the conversion of phosphorylase b to phosphorylase a in a reaction [Eq. ( l ) ] in which the terminal phosphoryl group of A T P is transferred to a specific serine residue in the phosphorylase subunit 2 Phosphorylase b (dimer)

+ 4 ATP

Mgn+

Phosphorylase a (tetramer)

+ 4 ADP

(1)

(21). The sequence of amino acids a t the phosphorylated site in rabbit skeletal muscle phosphorylase a as determined by a study of chymo(NH2)tryptic peptides is -Ser-Asp-Glu (NH,) -Glu-Lys-Arg-LysGlu Ile-Ser (P)-Val-Arg-Gly-Leu. Phosphorylase b, the dephospho form of phosphorylase, requires high concentrations of 5’-AMP for activity and is generally considered to be less active in vivo than phosphorylase a 17. M. Rabinowitr, “The Enzymes,” 2nd ed., Vol. 6, p. 119, 1962. 18. E. G. Krebs, Curr. Top. Cell. Regul. 5, 99 (1972). 19. E. H. Fischer, L. M. G. Heilmeyer, Jr.. and R. H Haschke, Curr. Top. Cell. Regul. 3, 211 (1971). 20. J . Lamer, Curr. Top. Cell. Regul. 3, 196 (1972). 21. E. G. Krebs and E. H. Fischer, Adwan. Enzymol. 24, 263 (1962).

558

DONAL A. WALSH AND EDWIN G . KREBS

(22). Physiological conditions under which conversion of phosphorylasc b to phosphorylase a occurs include muscle contraction (23) and epinephrine administration (24). The occurrence of a strain of mice in which the adults lack skeletal muscle phosphorylase kinase has provided confirmation and additional insight into the role of this enzyme in metabolism (25, 2 6 ) . I n these mice (I strain) there is no conversion of phosphorylase b to phosphorylase a in response to muscle contraction or epinephrine. Muscle glycogen does break down under these conditions, however, indicating that some process other than the phosphorylation of phosphorylase is involved (25,271. For the glycogenolytic response accompanying muscle contraction, it is probable that this can be accounted for on the basis of 5’-AMP formation which results in the stimulation of phosphorylase b. It is difficult to explain, however, the mechanism by which epinephrine causes muscle glycogenolysis in I strain mice although this may reflect an altered balance between synthesis and degradation.

b. Specificity. I n addition to catalyzing the phosphorylation of phosphorylase b by ATP, rabbit muscle phosphorylase kinase also catalyzes its own phosphorylation (28). This reaction, which is much slower than reaction ( l ) , appears to involve the phosphorylation of serine in more than one peptide sequence in phosphorylase kinase (29) and is accompanied by activation of the kinase as discusscd below. Phosphorylase kinase also catalyzes the phosphorylation of casein (28) but this reaction is also very slow, i.e., less than one thousandth the rate a t which phosphorylase b is phosphorylated. Very recently (SO) it has been determined that the 22,000-24,000 molecular weight component of troponin (TNI)(31) serves as a substrate for phosphorylase kinase. I n this instance the rate of the reaction is within an order of magnitude of that 22. C. Nolan, W. B. Novoa, E. G . Krebs, and E. H. Fischer, Biochemislry 3, 542 (1964). 23. C. F. Cori, in “Enzymes: Units of Biological Structure and Function” (0. H. Gaebler, ed.), p. 573. Academic Press, New York, 1956. 24. E. W. Sutherland, Symp. Phosphorus Metab. 1, 53 (1951). 25. J. B. Lyon, Jr. and J. Porter, JBC 238, 1 (1963). 26. J. B. Lyon, Jr., Biochem. Genet. 4, 169 (1970). 27. W. H. Danforth and J. R. Lyon, Jr., JBC 239, 4047 (1964). 28. R. J. DeLange, R. G. Kcmp, W. D. Riley, R. A. Cooper, and E. G. Krebs, JBC 243, 2200 (1968). 29. W. D. Riley. R. J. DeLange, G . E. Bratvold, and E. G. Krebs, JBC 243, 2209 (1968). 30. J. T.Stull, C. 0. Brostrom, and E. G. Krebs, JBC 247, 5272 (1972). 31. J. Potter and J. Gergely, Fed. PYOC.,Fed. Amer. SOC. Exp. Biol. 31, 501 (1972) (abstr.).

16.

559

PROTEIN KINASES

whicli occurs with phosphorylase b. The physiological significance of TNI phosphorylation has not been determined, but the existence of this reaction may cast doubt on the validity of classifying phosphorylase kinase as a specific protein kinase. Skeletal muscle phosphorylase kinase readily phosphorylates liver phosphorylase from the same or different species ( 3 2 ) . c. Purification and Molecular Properties. Phosphorylase kinase is a large rnoleculc and is located within the muscle fiber in close association with glycogen and its substrate, phosphorylase b ( 2 1 ) . When the pH of a muscle extract is lowered to 6.1, a large, almost flocculent precipitate develops which contains phosphorylase kinase, phosphorylase phosphatase, glycogen synthetase, phosphofructokinase, and part of the tissue content of phosphorylase and glycogen (13, 28, 33-36). This single step results in a 10-fold enrichment of the enzyme. Subsequent purification steps including differential centrifugation, ammonium sulfate fractionation, and gel filtration utilizing Sepharose 4B (37) result in the isolation of essentially homogeneous enzyme having a specific activity 200-fold that of the enzyme in the crude extract. Purified phosphorylase kinase sediments in the ultracentrifuge as a single boundary with s20,w = 26.1 S as extrapolated to infinite dilution (37). The weight average molecular weight determined by meniscus depletion equilibrium is 1.33 X loc. The absorption spectrum shows a maximum a t 279 nm and a minimum a t 251 nm; A:$ nm equals 11.8. On disc gel electrophoresis in the presence of sodium dodecyl sulfate the kinase exhibits three bands corresponding to molecular weights of 118,000, 108,000, and 41,000 which are referred to as subunits A, B, and respectively. From the relative densities of the bands as determined by scanning the gels, the formula A4B4C, has been assigned to the enzyme. Purified phosphorylase kinase forms a complex with glycogen ( 2 8 ) .-4s isolated, nonactivated phosphorylase kinase contains 0.4-0.8 mole of endogenous serine-bound phosphate per lo" g of protein ( 3 8 ) .

c,

32. M. M. Appleman, E. G . Krebs, and E. H. Fischer, Biochemistry 5, 2101 (1966). 33. A. Parmeggiani, J. H. Luft, D. S. Love, and E. G. Krebs, JBC 241, 4625 (1966). 34. E. G. Krcbs, D. S.Love, G. E. Bratvold, K. A. Trayser, W. L. Meycr, and E. H. Fischer. Biochemistry 3, 1022 (1964). 35. T. R. Soderling, J. P. Hickenbottom, E. M. Reimann, F. L. Hunkeler, D. A. Walsh, and E.G. Krebs, JBC 245, 6317 (1970). 36. S. S. Hurd, Ph.D. Thesis, University of Washington, 1967. 37. T. Hayakawa, J. P. Perkins, D. A. Walsh, and E. G. Krebs, Biochemistry (1973) (in presa). 38. S. E. Mnycr and E. G. Krebs, JBC 245, 3153 (1970).

560

DONAL A. WALSH AND EDWIN G . KREBS

d. Kinetic Properties. 1. Activation-The most interesting property of muscle phosphorylase kinase revealed from a study of its kinetic behavior is the existence of the enzyme in two interconvertible molecular forms (IS).One of these forms, nonactivated phosphorylase kinase, has very little activity a t p H 6.8 or lower but displays activity a t higher pH values (Fig. 1, curve A) (39). The other form, activated phosphorylase kinase, is active over a wide range of pH (Fig. 1, curve B) (39).These differences in activity relate primarily to the higher affinity of the activated form for the substrate phosphorylase b ( 9 4 ) .As isolated from rabbit muscle under conditions in which no particular precautions are taken, i.e., rabbits simply killed with an overdose of anesthetic, muscles removed without quick freezing, etc., essentially all of the enzyme is in the nonactivated form. Conversion of nonactivated phosphorylase kinase to activated phosphorylase kinase is achieved by preincubating the enzyme with ATPMg?'. During this procedure the enzyme becomes phosphorylated. Unlike the conversion of phosphorylase b to phosphorylase a , however, activation of the kinase is associated with phosphorylation of serine in more than one peptide sequence (29). Although it is possible that one sequence constitutes a specific activation site, no good evidence for this has yet been found. During activation by ATP-Mg2+ site (s) in subunit B are the first to be phosphorylated, but this is followed by the phosphorylation

PH

FIG.1. The activity-pH profile for nonactivated (curve A) and activated (curve R) rnbhit skeletal muscle phosphorylase kinase from Krebs et a / . (39). 39. E. G. Krcbs, R. J. DeLange, R. G. Kemp. and W. D. Riley, Phormacol. Rev. 18, 163 (1966).

16.

561

PROTEIN KINASES

of site(s) in subunit A (40). Subunit C is not susceptible to phosphorylation. Two different enzymes can serve as catalysts for the phosphorylation and activation of phosphorylase kinase (41). As noted above, one of these is phosphorylase kinase itself (autoactivation). The other enzyme is a cyclic AMP-dependent protein kinase (see below) present as a contaminant in purified phosphorylase kinase preparations. Stimulation of this latter kinase by cyclic AMP accounts for the effect of the cyclic nucleotide on the activation process as observed in early studies (IS,34). Activation of phosphorylase kinase by ATP, as catalyzed by the cyclic AMP-dependent protein kinase, is believed to constitute a key step in the regulation of glycogenolysis by epinephrine and other hormones (S9,49). This is illustrated for epinephrine in Fig. 2. Epinephrine stimulates adenyl cyclase leading to the production of cyclic AMP which in turn accelerates the rate of phosphorylation and activation of phosphorylase kinase. The latter enzyme converts phosphorylase b to phosphorylase a resulting in an increased rate of glycogen breakdown. Activation of phosphorylase Epinephrine I

i Adenyl cyclase

n

ATP

Cyclic 3', 5'-AMP 1

t

Protein kinase Nonactivated phosphorylase kinase

+ ATP

Activated phosphorylase kinase

n

Phosphorylase b

+

ATP

Phosphorylase a

n

Glycogen

+

Glucossl-P

Pi

FIG.2. The mechanism of action of epinephrine in the regulation of skeletal muscle plycogenolysis. 40. T. Hayakxwa, J. Perkins, and E. G. Krebs, Biochemistry (1973) (in press). 41. D. A. Walsh. J. P. Perkins, C. 0. Brostrom, E. S. Ho, and E. G. Krebs, JBC 246, 1968 (1971). 42. D. A . Wdsh, E. G . Krebs, E. M. Reimann, M. A. Brostrom, J. D. Corbin,

J. P. Hickenbottom, T. R. Sodding, and J. P. Perkins, Advan. Biochern. Psychophovmncol. 3, 265 (1970).

562

DONAL A. WALSH AND EDWIN G. KREBS

kinase by the autocatalytic mechanism is relatively slow and may not be of any physiological significance. In addition to the activation of phosphorylase kinase which results from phosphorylation, the nonactivated form of the enzyme can also be activated as a result of limited proteolysis. This process is probably of no importance physiologically, but it is of historical interest and may also serve as a useful tool in studying structurefunction relationships for this enzyme. In this type of activation (34, 43, 44) the nonactivated form is converted to a form exhibiting a pH optimum curve similar to that of the phosphorylated form of the enzyme, i.e., curve B of Fig. 1. Incubation of the kinase with proteolytic enzymes can be prolonged for a considerable period of time without a loss of activity ( 4 5 ) .I n this process subunits A and B undergo extensive degradation as evidenced by gel electrophoresis in the presence of sodium dodecyl sulfate, but subunit C remains unaltered suggesting that it may constitute the “catalytic subunit” of the enzyme. One of the proteolytic enzymes which is capable of activating phosphorylase kinase is a Ca2+-requiringprotease of skeletal muscle (44). This enzyme, originally referred to by the acronym, KAF, standing for kinase activating factor (43, 4 6 ) , causes activation of phosphorylase kinase during the isolation procedure unless a chelating agent is present to bind the metal, Activation of phosphorylase kinase by Cat+ ions derived from filter paper led to the original discovery of the conversion of phosphorylase b to phosphorylase a in vitro (47‘). 8. Stimulation by Cu2+-Tn addition to the effect of calcium brought about by preincubating phosphorylase kinase with KAF (see above), a second effect of this metal ion is seen in the phosphorylase b to phosphorylase u reaction itself. This effect, referred to as %timulation” to distinguish it from activation, is the result of a direct requirement of the enzyme for Ca2+ (4.3, 46, 48) and does not lead to a covalent alteration of the protein. Very low concentrations of Ca2+,i.e., at the micromolar level, suffice to saturate the enzyme so that the metal requirement is ordinarily not encountered as a result of the presence of contaminating calcium in the reagents used in activity tests. If chelating agents are used (49, 46, 4 8 ) , or if special care is used to purify all reagents ( 4 9 ) , 43. W. L. Meyer. E. H. Fischer, and E. G. Krebs, Biochemistry 3, 1033 (1964). 44. R. B. Huston and E. G. Krebs, Biochemistry 7, 2116 (1968). 45. D. J. Graves. T. Hayakawa, and E. G . Krebs, unpublished results (1972). 46. E. G. Krebs, R. B. Huston, and F. L. Hunkeler, Advan. Enzyme Requl. 6, 245 (1968). 47. E. H. Fischer and E. G. Krebs, JBC 216, 121 (1955). 48. E. Ozawa, K. Hosoi. and S. Ebashi, J. Biochem. (Tokyo) 81, 531 (1967). 49. C. 0. Brostrom. F. L. Hunkeler, and E. G. Krebs, JBC 246, 1961 (1971).

16.

PROTEIN KINASES

563

the need for Ca2+can be demonstrated readily. The concentration of Ca2+ required to stimulate the enzyme half-maximally is in the range of 1 x 10-7M to 3 X 10-6M, depending upon the pH and whether or not the enzyme is in its nonactivated or activated form (46, 48-50). Brostrom et al. (49) noted that the activated (phosphorylated) form of phosphorylase kinase displays an increased sensitivity to Ca2+. The kinase has a high affinity for Ca2+ions and binds about 25 moles per mole of enzyme under saturating conditions, i.e., M . Approximately half of the metal is bound more tightly than the remainder, the enzyme-Ca2+ complex for this portion having a dissociation constant of the order of M (46, 49). Heilmeyer et al. (50) obtained evidence that the kinase bound to glycogen has a somewhat greater requirement for Ca2+than is found for the purified enzyme. The requirement of phosphorylase kinase for Ca2+ is of particular significance with respect to the coupling of glycogenolysis to muscle contraction. Meyer et al. (43) first called attention to this control mechanism which has since been emphasized in numerous other reports (48, 50-54). When Ca2+is released from the sarcoplasmic reticulum of muscle in response to nerve stimulation, it triggers muscle contraction and a t the same time stimulates phosphorylase kinase (Fig. 3 ) . This leads to the conversion of phosphorylase b to phosphorylase a and subsequent glycogenolysis. An energy-requiring process (contraction) is thus linked t o an energy-producing metabolic sequence. Evidence for the validity of such a scheme was obtained by Brostrom et al. (49) who showed that an isolated sarcoplasmic reticulum fraction derived from muscle is capable of inhibiting phosphorylase kinase in vitro. 3. Interaction with glycogen-Phosphorylase kinase activity is stimulated by glycogen. I n the presence of Mgw, the interaction of the enzyme with this polysaccharide results in the formation of a flocculent precipitate (R8), a phenomenon more readily demonstratable with the activated (phosphorylated) than with the nonactivated form of the enzyme. Activation of the kinase by Mg2+-ATPis also enhanced by glycogen (42). The importance of glycogen as a regulatory agent in the conversion of phosphorylase b to phosphorylase a has been emphasized (50). 4. Other properties-The K , for the Mg2+-ATP complex in the phos50. L. M. G. Heilmeyer, Jr., F. Meyer, R. H. Haschke, and E. H. Fischer, JBC 245, 6649 (1970). 51. G. I. Drummond, J. P. Haraood, and C. A. Powell, JBC 244, 4235 (1969). 52. A. J. D. Friesen, G . Allen, and J. R. E. Valadares, Science 155, 1108 (1967). 53. S. E. Mayer, D. H. Namm, and J. P . Hickenbottom, Advan. Enzyme Regul. 8, 205 (1970). 54. C. Villar-Palasi and S. H. Wei, Proc. N o t . Acad. Sci. U. S . 67, 345 (1970).

564

DONAL A. WALSH AND EDWIN G. KRERS

Nerve stimulation

7

&lease of Caz+ from sarcoplasmic reticulum

Phosphorylase

A n Phosphorylase a

Phosphorylase b

Glycogen

Glucose- 1-P

+ pi

FIG. 3. The mechanism of action of Ca2+in the regulation of skeletal muscle glycogenolysis.

phorylase b to a reaction is 2.3 X lo-' M (34)and is essentially identical for nonactivated and activated phosphorylase kinase. ATP in excess of Mg2+ is sharply inhibitory. Heparin stimulates the activity of phosphorylase kinase at pH 6.8 (34).

2. Heart Muscle Phosphorylase Kinase Comparatively little work has been carried out on phosphorylase kinase from heart muscle as compared to the enzyme from skeletal muscle, but those studies which have appeared (55-57) emphasize that the major properties of the enzymes from the two sources are similar. Both kinases exist in nonactivated and activated forms differing in their ratios of activity a t pH 6.8 to activity a t pH 8.2, i.e., as in Fig. 1. The heart muscle enzyme, like its counterpart from skeletal muscle, is readily activated by MgZ+-ATPor by limited proteolysis. Heart muscle phosphorylase kinase also requires Ca2+ions for activity (47,58).It is of interest that in the fractionation of heart muscle homogenates the activated form of the kinase appears to adhere to the myofibrillar fraction (55); this could be significant in relation to the recently recognized activity of phosphorylase kinase toward troponin (30). 55. K. E. Hammermeister, A. A. Yunis, and E. G. Krebs, JBC 240, 986 (1965). 56. G. I. Drummond, L. Duncan, and A. J. D. Friesen, JBC 240, 2778 (1965). 57. G. I. Drummond and L. Duncan, JBC 241, 3097 (1966). 58. D. H. Namm, S. E. Mayer, and M. Maltbie, Mol. Pharmacol. 4, 522 (1968).

16.

PROTEIN KINASES

565

3. Liver and Other Mammalian Tissue Phosphorylase Kinuses The activation of glycogen phosphorylase in liver led to the original discovery of cyclic AMP more than 15 years ago [see reference (59) for a review]. Despite the probable importance of phosphorylase kinase in this effect, the enzyme has received comparatively little attention. Riley, in a preliminary note (60),reported that 75-fold purified dog liver phosphorylase kinase preparation was activated in a time-dependent process when incubated with Mg2+-ATP. The activation reaction which was accompanied by phosphorylation of the kinase was stimulated by cyclic AMP. The reaction caused a lowering of the K , for the substrate dephosphophosphorylase. Brain phosphorylase kinase has been studied by Drummond and Bellward (61) and was found to resemble the skeletal muscle enzyme. Smooth muscle phosphorylase kinase has been investigated by Mohme-Lundholme (62)and also by Albizati and Walsh ( 6 3 ) . 4. Phosphorylase Kinases frona Nonmammalian Sources

Insect flight muscle phosphorylase kinase from the blowfly, Phorinia regina, has been studied by Sacktor and his associates (64, 6 5 ) . This enzyme, like its mammalian muscle counterpart, requires Ca2+ions. It is also stimulated by inorganic phosphate. Activation as a result of phosphorylation has not been reported.

B. PYRUVIC DEHYDROGENASE KINASE Linn et al. (11, 6 8 ) have shown that the activity of pyruvic dehydrogenase is regulated by the phosphorylation and dephosphorylation of the dehydrogenase component (PDH) of this enzyme complex. The phosphorylated form of the enzyme is less active than the nonphosphorylated form. A specific protein kinase and a phosphoprotein phosphatase are involved in this system as shown in Eqs. (2) and (3). 59. G. A. Robison, E. W. Sutherland, and E. Butcher, “Cyclic AMP.” Academic Press, New York, 1971. 60. G. A. Riley, Pharmacologist 11, 253 (1969). 61. G. I. Drummond and G. Bellward, J. Neurochem. 17, 475 (1960). 62. E . Mohme-Lundholme, Acta Physiol. Scand. 59, 74 (1963). 63. L. D. Albizati and D. A. Walsh, Pharmacologist 13, 315 (1971). 64. R. G. Hansford and B. Sacktor, FEBS Lett. 7, 183 (1970). 65. B. Sacktor, N.-C. Wu, and W. D. Reed, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 30, 1176 (1971). 66. T . C. Linn, F. H. Pettit, F. Hucho, and L. J . Reed, Proc. Nat. Acad. Sci.

U. S. 64,

227 (1969).

566

DONAL A. WALSH AND EDWIN G. KREBS

PDH

+ ATP

PDH-P

PDH kinase

+ H20

PDH-P

+ ADP

PDH pbmphstase

PDH

(2)

+ Pi

(3) Pyruvic dehydrogenase kinasc is not affected by cyclic AMP. It is an intramitochondrial protein kinase and appears to be closely associated with the dehydrogenase itself. The relationship of this enzyme to the mitochondria1 phosvitin kinase (67) has not been determined, nor is it known what role pyruvic dehydrogenase kinase plays in the formation of the bulk of the serine-bound phosphate present in mitochondria (68). The relative amounts of phosphorylated and nonphosphorylated PDH can be varied in animals by administering fructose (69, 70), insulin (7072), or epinephrine (72).

111. Cyclic Nucleotide-Regulated Protein Kinares

A. CYCLIC AMP-DEPENDENT PROTEIN KINASES 1. Nomenclature In studying the mechanism whereby cyclic AMP stimulates glycogenolysis an cnzymc was identified in rabbit skeletal muscle which catalyzes the activation and concomitant phosphorylation of phosphorylase kinase ( l d , 4 l ). This enzyme, acting as a “phosphorylase kinasc kinase,” showed a complcte requirement for cyclic AMP for activity. It was also found to catalyze the phosphorylation of casein and protamine and was thus termed a “cyclic AMP-dependent protein kinase.” This term dcviatcs from standard nomcnclature in that it recognizes a specific activator but only a general class of substrates. As will be discussed in a later section, more than one cyclic AMP-dependent protein kinase may exist in a given tissue and eventually it may be important to classify these enzymes on the basis of substrate, specificity. For now, however, they will be considered collectively. M. Lorini, L. A. Pinna, V. Moret, and N. Siliprandi, BBA 110, 636 (1965). L. L. Bieber and P. D. Boyer, JBC 241, 5375 (1966). H. D. Soling and G. Bernhard, FEBS Lett. 13, 201 (1971). 0. Wieland, E. Siess, F. H. Schulse-Wethmar, H. G. von Funcke, and B. Winton, ABB 143, 593 (1971). 71. R. M. Denton, H. G. Coore, B. R. Martin, and P. J. Randle, Nature (London) New BioZ. 231, 115 (1971). 72. R. L. Jungas, Metab., Clin. Ezp. 20, 43 (1971). 67. 68. 69. 70.

16.

567

PROTEIN KINASES

2. Tissue and Subcellular Distribution

Cyclic AMP-dependent protein kinases have been identified in a wide variety of sources. Kuo and Greengard (73) , in a survey of bovine tissue, demonstrated the presence of the enzyme in each of thirteen different tissues examined. The enzyme was present in species from nine animal phyla. The subcellular distribution of protein kinase whose activity is dependent on cyclic AMP in various mammalian tissues is presented in Table I. In liver (74) and mammary gland (76) a t least 90% of the activity is detected in the cytosol and only small or negligible amounts are present in other fractions. In contrast to this in the anteriory pituitary (76) and whole brain (77) considerable activity is detected in other organelles, especially in thc mitochondria1 and microsomal fractions. The presence of a membrane-bound protein kinase was initially demonstrated by Jard and Bastide (78) who had shown that at least 30% of toad bladder cyclic AMP-dependent protein kinase was particulate. Cyclic AMP-dependent protein kinase is intimately associated with adrenal ribosomes (79) and with a wide variety of membraneous components of brain (77). Cyclic AMP-dependent protein kinases have not yet been described in plants, and a brief report (80) on the presence of the enzyme in Escherichia coli has not been extended. A protein kinase that is TABLE I SUUCELLULA~~ DISTHIUUTION OF CYCLICAMP-DEPENDENT PROTEIN KINASE Protein kinase activity

(% of total cyclic AMP stimulated activity) Tissue

Nucleus

Liver Mammary gland Anteriory pituitary Brain

6 0.4 7 8.6

Mitochondria 0

0.5 30 24.2

Microsomw 4

0.6 13 25.1

Cytosol

Ref.

90 98.5 50

76 76

77

42

78

73. J. F. Kuo and P. Greengard, Proc. Nut. Acad. Sci. U. S. 64, 1349 (1969). 74. L. J. Chen and D. A. Walsh, Biochemistry 10, 3614 (1971). 75. G. C. Majumder and R. W. Turkington, JBC 246, 2650 (1971). 76. S. Lemaire, G. Pelletier, and F. Labrie, JBC 246, 7303 (1971). 77. H.Maeno, E. M. Johnson, and P. J. Greengard, JBC 246, 134 (1971). 78. S. Jard and F. Bastide, BBRC 39, 559 (1970). 79. G. M. Walton, G. N. Gill, I. B. Abrass, and L. D. Garren, Proc. Nut. Acad. Sci. U . S. 68, 880 (1971). 80. J. F. Kuo and P. Greengard, JBC 244, 3417 (1969).

568

DONAL A. WALSH AND EDWIN G . KREBS

slightly stimulated by cyclic AMP has been reported in the Rauscher leukemia virus (81). 3. Purification

Cyclic AMP-dependent protein kinases have been purified from a variety of sources including skeletal muscle, 3000-fold (82) ; cardiac muscle, 1240-fold (83); adrenal cortex, 640-fold from cytosol (84) ; pineal gland, 17-fold (85); liver, 150-fold (86) ; anterior pituitary (87) ; mammary gland 1150-fold (75) ; brain, 277-fold (88); and trout testis, 35-fold (10). Of these preparations those from skeletal muscle, heart muscle and the adrenal are essentially homogeneous. The purification procedures utilized employed a wide variety of typical separation procedures and were marked by a low recovery of enzymic activity. 4. The Mechanism of Action of Cyclic A M P

a . Pertinent Kinetic Observations. I n the original description of a cyclic AMP-dependent protein kinase (12) the enzyme was found to exhibit a complete requirement for cyclic AMP. Later, as other methods of preparation of the skeletal muscle enzyme were used (89) and as enzymes from other tissues became available, a variable degree of cyclic nucleotide dependency was observed. A relatively pure preparation of the protein kinase from heart muscle (83) manifested as much as onesixth of its maximal activity in the absence of cyclic AMP. Aging of a protein kinase preparation was found to decrease the need for cyclic AMP (90). Reimann et al. (89) also showed that the order of addition of substrates markedly affected the cyclic nucleotide requirement; i.e., if a period of incubation of the protein kinase with histone were allowed, little or no cyclic AMP requirement was present. It was noted that the need for cyclic AMP is reduced a t low pH values (89). The concentration of cyclic AMP that is required for half-maximal 81. M. Strand and J. T. August, Nalure (London), New Biol. 233, 137 (1971). 82. C. 0. Brostrom, J. D. Corbin, and E. G. Krebs, Fed. Proc., Fed. A m e i . Soc. E z p . Biol. 30, 1089 (1971) (abstr.). 83. C. S. Rubin, J. Erlichman, and 0. M. Rosen, JBC 247, 36 (1972). 84. G . N . Gill and L. D. Garren, Proc. Nut. Acad. Sci. U . S. 68, 786 (1971). 85. J. A. Fontana and W. Lovenberg, Proc. Not. Acad. Sci. U . S. 68, 2787 (1971). 86. T. A. Langan, Science 162, 579 (1968). 87. F. Labrie, S. Lemaire, and C. Courte, JBC 248, 7293 (1971). 88. E. Miyamoto, J. F. Kuo, and P. Greengard, JBC 244, 6395 (1969). 89. E. M. Reimann, D. A. Walsh, and E. G. Krebs, JBC 246, 1986 (1971).

90. M. A. Brostrom, E. M. Reimann, D. A. Walsh, and E. G. Krebs, Adwan. Enzyme Regztl. 8, 191 (1970).

16. PROTEIN

569

KINASES

to 2 X 10-'M with protein stimulation of activity ranges from 1 X kinase from various tissues. As illustrated in Fig. 4 for rabbit skeletal muscle protein kinase the value is 1.5 X 10-sM, but the plot of cyclic A M P versus activity as indicated in Fig. 4 is atypical in that saturation of the enzyme occurs only a t very high cyclic A M P concentrations. This effect is also manifest by a nonlinear double reciprocal plot. The Hill plot is linear, however, and a coefficient of 1.26 suggests the possibility of a small degree of cooperativity. With nearly all protein kinases that have been investigated, i.e., the enzymes from skeletal muscle (89), cardiac muscle (90), anteriory pituitary (87) and reticulocyte (91), cyclic A M P has been found to increase the maximum velocity of the reaction, but is without effect on the K , for either ATP or the protein substrate. In contrast, it has been reported for the brain enzyme (88) that cyclic A M P causes a decrease in the K , for ATP,and with the enzyme from testis (92) that the cyclic nucleotide elevates the K , value for the nucleoside triphosphate. Except for these

[ C - A M P ] x 10'

FIG.4. The effect of cyclic A M P on the activity of rabbit skeletal muscle protein kinase. The inserts show (A) the Hill plot and (B) the Lineweaver-Burk plot of the data from Reimann et nl. (89). 91. M. Tao, ABB 143, 151 (1971). 92. A. H. Reddi, L. L. Ewing, and H. G. Williams-Ashman, BJ 122, 333 (1971).

570

DONAL A. WALSH AND EDWIN G . KREBS

last observations, however, all of the data support the concept that thc cyclic AMP-dependent protein kinase behaves like a “V system” as defined by Monod, thc simplest interpretation of which would suggest that cyclic AMP increases the number of available sites of catalytic activity. Cooperativity may be cxpectrd for thc activator, cyclic AMP, but not for the substrates. b. Activation Mechanism. I t has been determined that the cyclic AMPdependent protein kinase is made up of regulatory (R) and catalytic (C) subunits. Cyclic AMP binds to the regulatory subunit promoting dissociation of the holoenayme (RC) with the formation of the active catalytic subunit and a regulatory subunit-cyclic AMP complex [Eq. (4) 1 ,

+

+

RC cyclic A M P F! Re cyclic A M P C (inactive) (active)

(4)

Support for this mechanism originally came from several laboratories (90,93-96) and has since been extended to nearly all of the cyclic AMPdependent protein kinases investigated. Dissociation of RC in the presence of cyclic AMP has been demonstrated by the techniques of sucrose gradient ultracentrifugation (87, 89, 92), gel filtration (85, 97, 98), ion exchange chromatography (89), isoelectrofocusing electrophoresis ( 7 4 ) , and disc gel electrophoresis (82). The ready reversibility of the dissociation reaction has also been demonstrated. The addition of the regulatory subunit-cyclic AMP complex to isolated catalytic subunit blocks the activity of the latter in the abscnce of exogeneous cyclic AMP but is without effect in the presence of saturating concentrations of cyclic AMP (97, 99). Similarly, free catalytic subunit promotes the dissociation of [3HJ cyclic AMP from [“H] cyclic AMP-regulatory subunit complex (97). Either of two mechanisms could explain the action of cyclic AMP in dissociating the protein kinase. As indicated in Eqs. ( 5 ) and (6) an equilibrium may exist between the associated and dissociated form of the protein kinase. Addition of cyclic AMP would, by formation of a cyclic AMP-regulatory subunit complex, displace this equilibrium. A. Kumon, H. Yamamura, and Y. Nishizuka BBRC 41, 1290 (1970). 94. E. M. Reimann, C. 0. Brostrom, J. D. Corbin, C. A. King, and E. G. Krehs,

93.

BBRC 42, 187 (1971). 95. M. Tao, M. L. Salas, and F. Lipmann, Proc. Nut. Acad. Sci. U. S. 67, 408 (1970). 96. G. N. Gill and L. D. Garren, Proc Nut. Acad. Sci. U . S. 63, 512 (1969). 97. C. 0. Brostrom, J. D. Corbin, C. A. King, and E. G. Krebs, Proc. Nut. A&. Sci. U . S. 68, 2444 (1971). 98. J. Erlichman, A. H. Hirsch, and 0. M. Rosen, Proc. N u t . Acatl. Sci. 11. S. 68, 731 (1971). 99, G. N. Gill and L. D. Garren, Proc. Nut. Acnd. Sci. U . S. 63, 512 (1969).

16.

571

PROTEIN KINASES

(5)

RCeC+R

11 + cyclic AMP EL. cyclic AMP (6) Alternatively, a ternary complex may be formed which then leads to dissociation [Eqs. (7) and @)I.

+

RC cyclic AMP RC * cyclic AMP RC . cyclic AMP 5 R . cyclic AMP C

+

(7) (8)

No evidence is currently available to discern with certainty between the two possibilities. The question as to what cellular components might influence the dissociation of RC promoted by cyclic AMP is of importance to a full understanding of the regulation of protein kinase activity in vivo. Two effectors have been recognized which influence this reaction. Haddox et al. (100) have noted that preincubation of protein kinase with ATP results in a decreased binding of cyclic AMP to the protein kinase and concomitantly causes an increased requirement for cyclic AMP in the activation of the enzyme. Ashby and Walsh (101) have shown that a protein inhibitor of cyclic AMP-dependent protein kinases interacts directly with the catalytic subunit. At suboptimal concentrations of cyclic AMP this results in an increase in the concentration of R-cyclic AMP complex. Dissociation of the protein kinase can also occur in the absence of cyclic AMP. The alterations in cyclic AMP dependency that occur following preincubation of the enzyme with histone (89) have been shown to result from a histone-promoted dissociation of the protein kinase ( l o g ) . Tao (10.3) has shown likewise that protamine causes dissociation of the enzyme. The question is open as to whether undissociated holoeneyme, as assayed in the absence of cyclic AMP, possesses catalytic activity. Purified preparations of holoenzyme, apparently devoid of free catalytic subunit as based on physicochemical criteria, often do possess discernible protein kinase activity in the absence of cyclic AMP (85). To what extent this activity is the result of dissociation of the holoenzyme occurring under the conditions of low protein kinase concentration in the assay has not been evaluated. The changes in cyclic AMP dependency that occur on aging (90)or at low pH (89) probably also reflect dissociation of the protein kinase. 100. M. K. Haddox, N . E. Newton, D. K. Hartle, and N. D. Goldberg, BBRC 47, 653 (1972). 101. C. D. Ashby and D. A. Walsh, JBC (1973) (in press). 102. E. Miyamoto, G. L. Petzold, J. S. Harris, and P. Greengard, BBRC 4, 305 (1971). 103. M. Tao, BBRC 46, 56 (1972).

572

DOi’iAL A. WALSH AND EDWIN G . KREBS

The concentration of cyclic AMP required for half-maximal activation of protein kinase probably represents something other than a simple dissociation constant of the enzyme-nucleotide complex as indicated by Eqs. ( 5 ) - ( 8 ) . Nevertheless, it is of interest that the latter was found to M (104). This is in close agreement with concentration giving be 2 X half-maximal activation of the same enzyme (Fig. 3 ) . The apparent binding affinity of cyclic nucleotide is increased 5-fold in the presence of a heat-stable protein inhibitor of the protein kinase (104). 5. Properties

Studies of cyclic AMP-dependent protein kinases from a wide variety of tissues have in general indicated a marked similarity between the various enzymes. In addition to a general survey of tissues that has been performed by Greengard and his colleagues (7S, 105, 106) individual studies have included those on the adipose tissue (107), adrenal (84, 96) anteriory pituitary (87), bladder (78), brain (88), cardiac muscle (85, &I mammary ), gland (75, 108), liver (74, 86), pineal gland (85), reticulocytes (91), skeletal muscle (12, 82, 89), and mammalian (92) and trout testis (10).

a. Nucleoside Triphosphate Requirement. The li, value for ATP for each of the enzymes studied lies within the range of 7 pM to 20 pLM as tested at saturating concentrations of Mg2+.CTP, GTP, UTP, and ITP can replace ATP as the phosphoryl donor only poorly with a difference in K , value of at least 15-fold (109). At low concentrations (50 pM) GTP, UTP, CTP, and dTTP had no effect on the activity of the brain protein kinase as assayed with ATP as the phosphoryl donor (88). In contrast the enzyme from the anteriory pituitary was inhibited approximately 45% by GTP, CTP, and UTP when each of the latter were at a concentration of 0.1 mM and in a 4-fold excess of ATP (87). With either the skeletal muscle enzyme (109) or the cardiac muscle enzyme (98) the K , for ATP was identical for either the holoenzyme assayed in the presence of cyclic AMP or the isolated catalytic subunit. Although poor substrates the K , values for GTP and ITP for the skeletal muscle holo104. D. A. Walsh, C. D. Ashby, C. Gonzales, D. Calkins, E. H. Fischer, and E. G . Krebs, JBC 248, 1977 (1971). 105. J. F. Kuo, B. K. Krueger, J . R. Sanes, and P. Greengard, BBA 212, 79 (1970). 106. J. K. Kuo and P. Greengard, BBA 212, 434 (1970). 107. J. D. Corbin and E. G. Krebs, BBEC 38, 328 (1969). 108. C. T. Waddy and A. G. MacKinlay, BBA 250, 491 (1971). 109. C. 0. Brostrom, J. D. Corbin, and E. G. Krebs, unpublished observations (1972).

16.

PROTEIS KINASES

573

enzyme or catalytic subunit were found to be identical, lending further support to the proposcd activation mechanism [ Eq. (4) 1. b. Cyclic Sucleotide Specificity. Analogs of cyclic AMP in which either the 3’-0 or 5‘-0 position are substituted with a methylene group, the 2’-deoxy derivative, the xylofuranosyl analog, adenosine 3’,5’-cyclic phosphorothioate, 5’-AMP, 3’-AMP, 5’-IMP, and 5’-UMP are all inactive as stimulators of protein kinase (90, 105, 110). In contrast, modification of the purine ring such as in the 8-thio-, 8-0x0-, 8-benzylthio-, 8-amino-, and 3’,5’-tubercidin monophosphate results in compounds of equal or greater affinity for the protein kinase with K , values between 1- to 3-fold lower (110, 111 ) . Cyclic 3’,5’-monophosphate derivatives of naturally occurring nucleosidcs are poor activators of cyclic AMP-dependent protein kinases with K, values of approximately 40-fold, lOO-fold, and 200-fold higher than cyclic AMP for the inosine, guanosine, and cytosine nucleotides, respectively. c. Divalent Metal Ion Specificity. All protein kinases require a divalent metal for activity. The K, value for Mg2+is in the range of 1-3 mM. Magnesium ion is required for the expression of catalytic activity but not for the binding of cyclic AMP to the protein kinase components (112). Substitution of Mg2+by Co2+results in approximately a 3-fold increase in catalytic activity for all enzymes except those from testis in which the latter ion is inactive and from the pituitary in which the activity with Co2+was 60% of that expressed in the presence of Mg*+.For the mammary gland and brain enzymes, Mn2+ was 2-fold and 5-fold more effective, respectively, than Mg?+, but with other protein kinases it was considerably less effective. A notable exception to this is the enzyme described from E . coli (80) which exhibits a complete requirement for A h 2 + that cannot be replaced by Mg2+.A wide range of other divalent ions has been tested with the mammalian enzymes, and these ions are in general either totally ineffective or of minimal activity. Concentrations of Ca2+equiniolar to Mg2+are markedly inhibitory for both the hepatic (116) and anteriory pituitary (87) enzymes.

d. Protein Substrate Specificity. A number of proteins have been identified which act in vitro as phosphoryl acceptors for the reaction catalyzed by cyclic AMP-dependent protein kinases (Table 11) (10,1.2, 36, 110. G. I. Drummond and C. A. Powell, Mol. Pharmacol. 6, 24 (1970). 111. R. J . Bayer, K. R. Swiatek, R. K. Robins, and L. N. Simon, BBRC 45, 526 (1971). 112. A. M. Chambaut, F. Lefray, and J. Hanoune, FEBS Lett. 15, 328 (1971).

574

DONAL A. W A L S H AND E D W I N G . KREBS

TABLE I1 SUBSTRATES FOR CYCLIC AMP-DEPICNDENT PHOTICIN KINASES Substrate Phosphorylase kinase Glycogen synthetase Triglyceride lipase Histone Protamine Ribosomal proleills Tropoiiiii Neurotubule protein RNA polymerase Polyniicleot.idephosphorylase Casein Phosvitin Membrane proteins

Source of substrate Skeletal muscle Skeletal muscle Adipose tissue Thymus Trout testis Eschrrichia coli Liver Adreiial Skeletal muscle Brain Escherichia wli Eschrrichia coli Milk

Ref.

41 35, 113 114,116 86 10 116 117,118 79 110

180 181 188 18

Em

183

Liver Erythrocyte

126

124

41, 79,86, 113-126). To what extent, if any, all of these proteins serve as substrates physiologically remains uncertain. To date, strong evidence exists in support of the cyclic AMP-dependent phosphorylation of phosphorylase kinase and glycogen synthetase in both skeletal muscle and heart (14,58, 70,1267,and triglyccride lipase in adipose tissue (127).In 113. K. K. Schlender, S. H. Wei, and C. Villar-Palasi, BBA 191, 272 (1969). 114. J. D. Corbin, E. M. Reimann, D. A. Walsh, and E. G. Krebs, JBC 245, 4849 (1970). 115. J. K. Huttunen, D. Steinberg, and S. E. Mayer, BBRC 41, 1350 (1970). 116. J. Traugh and R. R. Traut, Fed. Proc., Fed. Amer. Soc. Exp. BioZ. 30, 1204 (1971) (abstr.). 117. C. Eil and I. G. Wool, BBRC 43, 1001 (1971). 118. J. E. Loeb and C. Blat, FEBS Lett. 10, 105 (1970). 119. C. Bailey and C. Villar-Palasi, Fed. Proc., Fed. Amer. Soc. Exp. Biol. 30, 1147 (1971) (abstr.). 120. D. B. P. Goodman, H. Rasmussen, F. DiBella, and C. E. Guthrow, Jr., Proc. N a t . Acad. Sci. U . S. 67, 652 (1970). 121. 0. J. Martelo, S. L. C. Woo, E. M. Reiniann, and E. W. Davie, Biochemistry 0, 4807 (1970). 122. M. N. Thang and F. Meyer, FEBS Lett. 13, 345 (1971). 123. E. Miyamoto, J. F. Kuo, and P. Greengard, Science 165, 63 (1969). 124. L. Shlatz and G. V. Marinetti, BBRC 45, 51 (1971). 125. M. J. Duffy and V. Schwarz, BJ 126, 12 (1972). 126. J. B. Posner, R. Stern, and E. G. Krebs, JBC 240, 982 (1965). 127. R. W. Butcher, R. J. Ho, H. C. Mcng, and E. W. Sutherland, JBC 240, 4515 (1965).

16.

575

PROTEIN KINASES

addition, the phosphorylation of hepatic f, histone ( I % ) , ribosomes (12929) , and membrane proteins (130)has been shown to occur in response to a cyclic AMP-mediated signal. Although a wide variety of histones have been shown to be phosphorylated in vivo, only with the f, species has this been demonstrated to occur in response to a cyclic AMP signal. Studies of substrate specificity have been made primarily with those proteins that. can be obtained readily from commercial sources, i.e., protamine, phosvitin, casein, and the histones. Of these, the first two do not occur in mammalian tissues and casein is confined primarily to the mammary gland. The relative order of activity obtained with the different substrates is glycogen synthetase phosphorylase kinase > histone > casein > phosvitin. Comparison made with various histone fractions indicate that the rate with different histone fractions is f2,, > f, > f, or f,. Activity with protamine, as assayed at high ionic strength with the trout testis enzyme, is 13-fold greater than with unfractionated histone, but these two are essentially equivalent a t low ionic strength (10).With other protein kinases tested, rates with protamine were comparable to those obtained with either histone fPb or fl. I n general, these comparisons have shed little information on the physiological significance of phosphorylation except to imply that cyclic AMP-dependent protein kinases exhibit a broad specificity. This conclusion may be invalid since it is to be expected that a comparison of inappropriate substrates would shed little information of differences between enzymes. It may be more justified to conclude that with many enzymes the physiological substrate has not been recognized. The K , values determined for histone and casein are in the range of 0.2-2.0 mg/ml. In all instances evaluated, the phosphoryl acceptor in protein kinase reactions is a seryl residue. Histone fl is phosphorylated in response to a cyclic AMP signal on the serine residue following alanine in the sequence Lys-Ala-Ser-Gly- (Pro),-Val-Ser-Glu-Leu-Ile-Thr-Lys ( I S 1) . Protamine (132), as phosphorylated in vivo during the development of the trout testis, can be isolated with phosphate associated with all 4 seryl residues in a sequence

>

Pro-(Arg)lSer-Ser-Ser-Arg-Pro-Val-(Arg)a-Pro-(Arg)TVal-Ser-(Arg)~Gly-Gly-(Arg)r -(Ile)-(Ah)-

Comparison of tryptic digests of 32P-phosphoprotamine produced either 128. T. A. Langan, JBC 244, 5763 (1969). 129. C. Blat and J. E. Loeb, FEBS Lett. 18, 124 (1971). 130. R. N. Zahken, A. A. Hockberg, F. W. Stratman, and H. A. Lardy, Proc. Nat. Acad. Sci. U . S. 69, 800 (1972). 131. T. Langan, A7m. N . Y . Acad. Sci. 185, 160 (1971). 132. M. M. Sanders and G . H. Dixon, JBC 247, 851 (1972).

576

DONAL A. WALSH AND EDWIN Q. KREBS

in vivo in whole spermatid cells or in vitro by incubation with cyclic AMP-dependent protein kinase indicates that each of the seryl residues can be phosphorylated in response to cyclic AMP.

e. Ionic Strength. As assayed with casein as substrate, high ionic strength (0.1 M NaC1) inhibits the skeletal muscle enzyme resulting in a 5-fold increase in the K, of the protein substrate (89). With the enzyme from adipose tissue 0.1 M NaCl is partially inhibitory (-30%) as assayed with casein as substrate but slightly stimulatory (czlOo/o) when histone is used (133). This reault is observed when either holoenzyme or isolated catalytic subunit is used as the enzyme source. The enzyme from trout testis is stimulated 3-fold by 0.3 M of either potassium chloride, ammonium chloride, sodium chloride, or sodium acetate, when assayed with protamine as substrate (10).This high ionic strength strongly inhihits the activity when histone is used. It would appear from the results obtained that the effects of ionic strength are primarily on the protein substrate rather than the enzyme per se. f. pH Optima. Maximum activity of skeletal muscle protein kinase is expressed at pH 5.8 with casein as substrate and pH 6.9 with histone as substrate. T o what extent these differences reflect proton dissociations of the protein substrates rather than titrations of specific residues of the protein kinase has not been evaluated. A wide variety of pH optima in the range of pH 6.0-9.0 has been described for the enzymes from various sources. Variations in reaction conditions and substrates used do not permit a meaningful comparison to be made. The concentration of cyclic AMP required for half-maximal activity and the maximum stimulation of activity are both markedly sensitive to pH, but this effect is not reflected in the amount of cyclic AMP required to saturate the binding sites (119). g . Efiects of Temperature. I n a study of the cyclic AMP-dependent protein kinase from reticulocytes (91) it has been shown that a difference exists in the stability to heat inactivation of the cyclic nucleotide binding and the catalytic activities. A combination of ATP and Mgw stabilizes both activities to heat inactivation; however, in the presence of Mg2+ alone the binding activity is more susceptible to denaturation than the catalytic function. I n contrast, cyclic AMP addition stabilises the binding function but destabilizes the catalytic activity. Cardiac protein kinase was found to be more sensitive to inactivation by heat in the presence than the absence of cyclic AMP (90). 133. J. D. Corbin, C. 0. Brostrom, R. L. Alexander, and E. G. Krebs, JBC 247, 3736 (1972).

16.

577

PROTEIN RINASES

h. Effects of Metabolites. Cyclic AMP-dependent protein kinases are not known to be subject to physiologically significant feedback regulation. At very low concentrations of ATP (‘2 X 10-5M), however, rat liver protein kinase can be inhibited by high concentrations (5 X lo4 M ) of adenine, adenosine, ADP, and AMP (134). Inhibition is observed with both holoenzyme and catalytic subunit but can be eliminated by a 25-fold increase in the ATP concentration. This would appear to indicate that interaction of these compounds with the enzyme occurs a t the catalytic binding site. An inhibition by high levels of cyclic AMP (1 mM) that has been reported for several protein kinases presumably occurs by a similar mechanism.

i. Effects of a Heat-Stable Protein. All mammalian tissues examined contain a heat-stable protein that inhibits the activity of cyclic AMPdependent protein kinases (104). The effect is noncompetitive with respect t o the substrates, ATP and proteins, and also to cyclic AMP. The inhibitor protein acts by a direct combination with the catalytic subunit (101).

j . Molecular Size. A summary of the sedimentation coefficients and molecular weights of various cyclic AMP-dependent protein kinases and their subunits as determined in several laboratories is presented in Table I11 (74,87,91,92,96,135, 136).The major form of the holoenzyme appears to be a 6.3 to 7.0s species with a molecular weight in the range of 120,000-160,000. A catalytic subunit having a sedimentation coefficient of about 4.0 S and molecular weight between 35,000 and 60,000 is common TABLE I11 THESEDIMENTATION COEFFICIENTS AND ESTIMATED MOLECULAR WEIGHTS M.4MMALIAN CYCLIC AMP-DEPENDENT PROTEIN KINASES AND THW SURUNITS DE RI VE D THEREFROM Holoenzyme Enzyme source Skeletal muscle Cardiac muscle Adrenal Liver Anteriory pituitary Mammalian testis Reticulocyte

6.8 6.3 7.04 6.8

=7

-

Catalytic subunit

OF

Regulatory subunit

MW

s20,w

MW

SX,,~

MW

Ref.

123,000 158,000 157,000 h.160,OOO

4.1 3.3

49,000 36,550 60,500 ?45,000 60,000

4.9 4.1

82,000 83,800 92,000 -

1.36 136 96

-

87 9.2

-

91

-

140,000

4 4.1 =4

-

-

-

-

134. H. Iwai, M. Inamasu, and S. Takeyama, BBRC 46, 824 (1972). 135. C. 0. Brostrom, C. A. King, and E. G. Krebs, unpublished results (1972). 136. J. Erlichmm, C. S. Ruhn, and 0. Rosen, personal communication (1972).

74

578

DONAL A. WALSH AND EDWIN G . KREBS

to all protein kinases that have been studied. The regulatory subunit is of approximately 80,000 molecular weight. Smaller forms of the regulatory subunit and holoenzyme have been recognized for the skeletal muscle enzyme (89),but these have been attributed to proteolysis (136).The stoichiometry of subunits in the protein kinase has not yet been conclusively established.

Ic. Charge of Cyclic AMP-Dependent Protein Kinases and Their Subunits. Holoenzyme and isolated regulatory subunits are acidic proteins, the isoelectric points of which for the liver enzyme are 5.2 and 4.5, respectively ( 7 4 ) . The catalytic subunit is basic with isoelectric points ranging from 6.3 to 8.6 ( 7 4 ) .It would appear that the charge characteristics of the holoenzyme reflect predominantly those of the regulatory subunit. At least three forms of hepatic catalytic subunit have been identified on the basis of differences in charge ( 7 4 ) .No significant differences between these forms has yet been identified with respect to substrate specificity. B. OTHERCYCLICNUCLEOTIDE-REGCLATED PROTEIN KINASES Kuehn (137)has described a protein kinase in the slime mold, Physarum polycephalum, whose activity is inhibited by cyclic AMP. Concentrations of the cyclic nucleotide greater than M inhibit the enzyme by more than 90%. Both this enzyme and a cyclic AMP-activated enzyme from the same slime mold are specific for casein as substrate and are essentially inactive with protamine, histone, or phosvitin. The existence of cyclic GMP-dependent protein kinase has been shown by Kuo and Greengard (138). The enzyme is predominant in Arthropoda and has been identified in many tissues. The apparent K, for cyclic GMP is between 2.5 X 10-sM and 4.8 X 10-'M. Cyclic AMP is approximately two orders of magnitude less effective. The divalent metal ion requirement of Coz+> MnZ+> Mgz+is similar to some cyclic AMP-dependent enzymes. The function of the cyclic GMP-activated enzyme is unknown.

IV. Nonelassifled Protein Kinases

Protein kinase activity which is detectable in a subcellular fraction and which does not require cyclic AMP could conceivably represent (1) 137. G. D. Kuehn, JBC 246, 6366 (1971).

138. J. F. Kuo and P. Greengard, JBC 245, 2493 (1970).

16.

PROTEIN KINASES

579

the free catalytic subunit of a cyclic AMP-dependent protein kinase; (2) a protein kinase that is not regulated by cyclic AMP; or (3) activi ty of undissociated cyclic AMP-dependent protein kinase, i.e., activity of the holoenzyme. Of these three possibilities, the last is not known to exist. It is relatively simple, however, to distinguish between the first two possibilities. Free catalytic subunit is inhibited by isolated regulatory subunit. In addition, currently available evidence would indicate that the heat-stable protein described by Walsh et al. (104) specifically inhibits all cyclic AMP-dependent protein kinases and the catalytic subunits derived therefrom. Thus, the inhibition by both regulatory subunit and inhibitor can serve as a mechanism for distinguishing free catalytic subunit. In general, these tests have not been applied and there exists considerable ambiguity in the field as t o whether various protein kinases whose activity is not stimulated by cyclic AMP represent free catalytic subunit of cyclic AMP-dependent protein kinase or alternat,ively a different class of enzyme. This section will be restricted to those enzymes for which there is some ambiguity concerning classification and about which new information is available since the previous review (17).

A. HISTONEKINASES Because of their ready availability, especially in comparison to other potential physiological substrates, histones have been used extensively in vitro in the study of cyclic AMP-dependent protein kinases. The phosphorylation of histones occurs in vivo in response to a wide variety of physiological stimuli including glucagon administration (1.98), hepatic regeneration (139, I@), phytohemagglutin-stimulated transformation of lymphocytes (141), pancreatic degeneration and regeneration (I&), and in response to x-irradiation (143). Of these the best characterized to date is that of hepatic growth in response to partial hepactectomy in which it is shown that the species of histone phosphorylated is a function of the time course of regeneration. In synchronized cell culture species-specific histone phosphorylation is a function of the cell cycle and mitotic activity. Whereas the hepatic cytoplasmic cyclic AMPdependent protein kinase does catalyze the phosphorylation of the four major classes of histones in vitro, only histone f i has been shown to 139. R. Gutierrer and L. S. Hnilica, Science 157, 1324 (1967). 140. M. G. Ord and L. A. Stocken, BJ 112, 81 (1969). 141. M. E. Cross and M. G. Ord, BJ 124, 241 (1971). 142. P. J. Fitrgerald. W. H. Mnrsh, M. G. Ord, and L. A. Stocken, BJ 117, 711 (1970). 143. L. R. Gurley and R. A. Walters, Biochemistry 10, 1588 (1971).

580

DONAL A. WALSH AND EDWIN G . KREBS

be phosphorylated in vivo in response to a cyclic AMP signal (128). The site of f, phosphorylated in vitro by cyclic AMP-dependent protein kinase is identical to that phosphorylated in vivo in response to cyclic AMP (see Section III,A,5,d). However, to what extent the other histone phosphorylations occurring under the various conditions cited above are catalyzed by cyclic AMP-dependent or cyclic AMP-independent enzymes remains to be established. Langan has identified a second enzyme, designated HK,, that is specific for a different seryl residue if f, histone in the sequence (Thr, Ser, Gly,, A1a)-Gly-Ser-PO1-Phe-Lys (131). Phosphorylation of this site also occurs in vivo. The activity of HK, is not stimulated by cyclic AMP nor is it inhibited by either cyclic AMP-dependent protein kinase regulatory subunit or the protein kinase inhibitor (144).

B. ACIDICNUCLEAR PROTEIN KINASES Protein kinase activity has been demonstrated as a component of the acidic nuclear proteins associated with rat liver chromatin (146, 146). Four distinct kinases have been identified (147). The enzymes catalyze the phosphorylation of casein, phosvitin, and acidic nuclear protein but not histone or protamine. The activity is not stimulated by cyclic AMP.

C. PHOSVITIN KINASES The phosphorylation of histone and phosvitin is catalyzed by separate enzymes from rat liver cytosol. Langan (86),in achieving a 150fold purification of hepatic histone kinase, altered the relative activities toward the two substrates by greater than 90-fold. The histone kinase was later shown to' be stimulated by cyclic AMP and also catalyzed the phosphorylation of protamine (86). Baggio et al. (148) have purified rat liver cytoplasmic phosvitin kinase extensively and found no detectable activity toward protamine. Multiple forms of hepat.ic phosvitin kinase have been recognized on the basis of differences in both size and change (1499).As mentioned earlier, Lorini e t aZ. (67) detected a phosvitin kinase in mitochondria. The function of these enzymes and 144. T.Langan, C.D. Ashby, and D. A. Walsh, unpublished observation (1972). 145. M. Kamiyama and B. Dastugue, BBRC 44, 29 (1971). 146. M. Takeda, H.Yamamura, and Y . Ohga, BBRC 42, 103 (1971). 147. R. W.Ruddon and S. L. Anderson, BBRC 46, 1499 (1972). 148. B. Baggio, L. A. Pinna, V. Moret, and N . Siliprandi, BBA 207, 516 (1970). 149. B. Baggio and V . Moret, BBA 250, 346 (1971).

16. PROTEIN

58 1

KINASES

their potential regulation by cyclic AMP has not been evaluated. An active phosvitin kinase is present in brain that is stimulated 3- to 4-fold by Na+ and K+ (150). It is considered unlikely, however, that this enzyme is associated with protein phosphorylation that occurs during active transport of monovalent cations (151). The cellular distribution of this enzyme is markedly similar to the cyclic AMP-dependent protein kinase described recently by Maeno et d.(77). ACKNOWLEDGMENTS The authors wish to acknowledge the support of the National Institutes of Health (Grants No. AM 13613 and No. AM 128421, the Muscular Dystrophy Associations of America, Inc., and the American Heart Association. D.A.W. is an Established Investigator of the American Heart Association.

150. R. Rodnight and B. E. Lnvin, BJ 93, 84 (1964). 151. 1,. Dccsi and R . Rodnight. J . Neurochem. 12, 791 (1965).

This Page Intentionally Left Blank

Author Index Numbers in parentheses are reference numbers and indicate that an author’s work is referred to, although his name is not cited in the text.

A Aanning, H. L., 436 Aaron, H. S., 213 Abdul-Baki, A., 60 Abeles, R. H., 145, 148 Abelson, P. H., 510 Abita, J. P., 173 Abou-zamzam, A. M., 281 Abraham, H. D., 68, 69(138) Abrahams, S. L., 248 Abramovitz, A. S., 178 Abrams, R., 24, 25(172), 37(238), 38, 39(238), 40(238) Abrass, I. B., 567, 574(79) Ackermann, W. W., 131 Adams, C. A., 35(227), 36 Adams, G. A., 108 Adams, I. T., 301 Adams, M. J., 454 Adelberg, E. A., 511(14), 512 Adelman, R. C., 286 Aebi, H., 386 Afting, E.-G., 261 Agarwal, K. C., 309, 310(27), 311(75), 312(56), 313(27), 314(25), 315(25, 27), 316(25) Agarwal, R. P., 308, 309(8), 310(8, 24, 27, 33), 311(25, 331, 312(56), 313(27), 314(8, 24, 25), 315(8, 24, 25, 271, 316 (8, 24, 25), 321(24), 322(8, 24), 323 (8), 324(8), 325(8, 24), 326(8, 241, 329(8, 24), 330(8), 331(241), 332 (8, 24, 33) Ahlfors, C. E., 240, 241(13), 242(13), 259 (131, 260(13), 262(13), 267(13), 267 (131, 272 583

Ailhaud, G. P., 156, 158, 159, 165, 173 Ainsworth, S., 380 Airth, R., 20 Akazawa, T., 57,58,74 Alberts, A. W., 156, 161, 166(8), 167(8), 168(36), 169, 170, 171, 173, 176(8), 179, 183, 18503, 50), 187, 188(1), 189, 190(81), 193(51), 194(51, 811, 195 (41, 81), 196(41), 197(41), 198(41) Alberty, R. A., 2, 3(4, 51, 4(4), 37(4), 415, 422 Albizati, L. D., 565 Albrecht, G. J., 53, 55(16), 62(16), 63 (161, 65(16), 60(16), 67(16), 68(16) Alden, R. A., 454 Alexander, M., 22(167), 24, 25(167) Alexander, R. L., 576 Allen, G., 563 Allen, G. W., 204, 232(13), 233(13) Allende, C. C., 6(32), 7, 9, lO(57, 64,651, 11, 12(57), 13(64, 65) Allende, J. E., 6(32), 7, 9, lO(57, 64,65), 11, 12(57), 13(64, 65) Allderdice, P. W., 340 Alpers, J. B., 259, 269(102), 342 Ames, B. N., 119 Anders, M., 21 Anderson, B. M., 185 Anderson, H. M., 340 Anderson, J. W., 262(113), 264 Anderson, L., 232, 233(105) Anderson, M. L., 56, 61(53) Anderson, P. W., 29 Anderson, R. L., 241, 245, 250(43) Anderson, S. L., 580 Anderson, W. A., 43, 44(266)

584

AUTHOR INDEX

Anderson, W. B., 42, 43(256), 44(258) Andersson, M., 379 Ando, K., 291, 292 Andrews, P., 315, 318(58) Anido, V., 396 Anraku, Y., 44, 46(274), 47(274) Appleman, M. M., 559 Argyrakis, M. P., 312(52), 313, 325(52), 329(52)

Arnett, E. M., 206 Arnold, H., 356, 359(15), 379(15) Amon, D. I., 83 Aronson, A. I., 551 Arrington, S., 85, 372 Ashby, C. D., 571, 572, 577(101, 1041, 579 (104), 580

Ashworth, J. M., 61 Askonas, B. A., 400 Asnis, R. E., 492 Atherton, R. S., 392, 445, 469, 470(69) Atkinson, D. E., 105, 106, 114(58, 63), 240, 247, 263(53, 541, 267(54), 285,

286, 332, 357, 366(33), 374(33), 376 (33), 504

Atkinson, M. R., 2, 3(7) Atzpodien, W., 240, 241(22), 242(22), 248

'

Ball, E. G., 488, 489(11), 490(11), 493 (111, 498(11) Ball, E. P., 278 Ballard, F. J., 55 Bamburg, J. R., 351(21), 356, 374, 376 (21)

Bandurski, R. S., 35(230), 36, 337, 340 (101, 347(10)

Bangham, M. A., 340 Bank, W. J., 240, 241(9), 242(9), 249(9), 258(9), 262(9)

Bar, H. P., 213 Barbalace, D. S., 374 Barber, E. D., 533 Barber, G. A,, 19 Barker, H. A., 123, 144, 145(73), 146(73), 148, 151(73), 152(73)

Barman, T. E., 232, 233(91a) Barnes, E. M., Jr., 165 Baron, D. N., 281 Barrio, J. R., 245, 300 Bartels, H., 337 Baskin, R. J., 385 Bass, S. T., 52, 53, 54, 55(16), 62(16), 63 (161, 65(16), 66(16), 67(16), 68(16)

Bassham, J. A., 74, 92(4), 337 (22), 261 Bastide, F., 567, 572(78) August, J. T., 21, 22(162), 23, 24(148, Basu, D. K., 54 162), 25(162), 26, 568 Battig, F. A., 42 Axelos, M., 52 Baumann, P., 249, 263(66), 264(66), 267 Axelrod, B., 280, 337, 340(10), 347(10) (66) Axelrod, J., 53 Baumgardt, B. R., 488, 490(21) Ayengar, P., 309, 310(29, 34), 312, 324 Bautz, E. K. F., 21 Bayer, R. J., 573 (29) Ayling, J., 169, 170 Bayley, P. M., 394 Azzone, G. F., 277 Bazylewicz, G. A., 259, 269(102) Becker, A., 47 Becker, R. R., 371 B Bedfort, N., 9(106), 10, 13(106) Baddiley, J., 167 Beere, A., 287 Baggio, B., 580 Beinert, H., 6, 12(19) Bailey, A., 281 Bell, J. L., 281 Bailey, C., 574 Bell, R. M., 171, 195(41), 196(41), 197 Bailey, E., 357, 373, 374(44), 376(44), 379 (41), 198(41) Bailey, K., 431 Bello, L. J., 312(53), 313, 325(53) Bakerman, H. A,, 160 Bellward, G., 565 Balbinder, E., 522 Baldwin, E., 460, 461(16), 462(16), 464 Belocopitow, E., 556 Bendall, J. R., 288, 338 (16), 468(16), 471(16), 482(16) Baldwin, R. L., 54,66(24) Bender, M. L., 185,220

AUTHOR INDEX

Benesch, R., 431 Benesch, R.E.,431 Benkmann, H.-G., 283 Benkovic, P. A., 205, 221, 229, 241, 244 Benkovic, S. J., 201(4, 51, 202, 203(9), 204(9), 205, 206(9), 210(9), 220, 221, 223(74), 225(4, 741, 226(74, 78), 227 (74), 228, 229, 232(78), 233(78, 99), 241, 244, 381 Bennett, T. P., 9(109), 10, 13(109) Benson, R. W.,183 Bent, H., 215 Bentley, M.,37(238), 38, 39(238), 40 (238) Benveniste, R.,29, 30 Benziman, M.,357, 366(32), 374(32), 375 (32) Berg, K.,283 Berg, P.,6, 7(15), 8(15), 9, lO(58, 59), 12(15, 16, 171, 13(58, 59, 94, 951, 21, 22(156, 1641, 24, 25(164), 308, 309, 310(3, 311, 312(3, 31), 315(31), 320 (31),321(3, 311, 324, 325(3, 311, 329 (31) Berg, T. L., 14 Bergmeyer, H. U., 4(9), 5, 492, 493(46), 494(46), 495(46), 496(46), 499(46), 501(46), 502(46) Bergren, W. R., 69 Berman, M.,503 Bernard, S.,356 Bernhard, G.,566 Bernheim, F.,123 Bernheim, M.L.C., 123 Bernlohr, R. W.,356, 370, 374(113), 375 (113), 376(113), 379(29), 551 Bernstein, I. A.,281 Bernstein, R. L.,59, 62(98), 66(98), 68 (98) Berry, R. S., 215 Bertrand, J. A., 219 Bessman, M. J., 312(53, 53d, 313 Beutler, E.,339 Bew, M.,258 Bianchi, P. A,, 311(44), 313, 325(44) Bieber, L.L.,566 Bilik, E.,142 Bird, I. F.,56 Birge, C. H.,191, 193(88)

585 Birktoft, J. J., 185,454 Birnbaum, E.R.,473 Bischoff, E.,67 Bishofberger, H.,356, 358(13), 359 Biswas, C.,547, 548 Black, J. A., 356,361 Black, M.K.,309, 312(19), 319(19), 325 (19) Black, S., 6, 12(13, 14), 13(13), 504, 510, 511, 512, 513, 533 Blagoveshchenskyi, V. A., 491, 501(3) Blair, J. McD., 286 Blake, C. C. F., 340, 341(30), 344(30), 345(30), 351(30) Blangy, D., 240(25, 261, 241, 242(25, 261, 245, 246(48), 253(25), 256(25), 263 (48), 265 (48), 267(48), 530 Blat, C., 574, 575 Blethen, S. L., 460, 461(13, 18, 25, 26), 462(13, 181, 464, 466(13, 18, 25, 481, 467(13, 181, 468(18), 469(13, 18), 470 (13), 471(25), 472(25, 481, 474(25), 476(25, 48), 478(18), 479(13, 18, 481, 480(18, 481, 482(25), 483(25), 484 (251, 486(25) Bloch, K., 156, 157(7), 158, 159(17), 160 (171,161, 190(7), 191 Bloemers, H.J. P., 9, 12(78) Blow, D.M., 185, 454 Bloxham, D.P.,245, 255 Bluestein, H.,9, 10(64), 13(64) Blume, A. J., 522 Blume, K. G., 356, 359(15), 379(15) Blumenthal, H. J., 505 Bock, A.,9(103), 10, 12(76), 13(103), 503 Bock, R. M., 6, 12(18), 160 Bode, H., 240, 241(22), 242(22), 248(22) Boezi, J. A., 24, 25(169), 54 Bognara, A. S., 115 Boiteux, A., 276 Bolton, E.T.,510 Bomsel, J.-L., 281,285 Bondar, R. J. L., 355, 361, 371 Bondinell, W., 365 Bonner, D. M.,522, 539(43) Bonnett, R.,144 Bono, V. H., Jr., 289(70), 291, 293(70) Bornmann, L., 358, 359(52a) Borsook, H., 123

586 Boulanger, Y., 9(106), 10, 13(106) Bourgeois, S., 492 Bowen, T. J., 281 Bowen, W. J., 298,299(82) Bowman, I. B. R., 357, 361, 374(40) Boyd, D. B., 218 Boyer, P. D., 9, 183, 250, 353, 354(1), 355, 358(1), 360(1), 364, 366, 368'3, 369(1), 370, 371, 372(1), 375(1), 376 (1, 114), 379(1), 380(1), 385, 566 Bozzi, M. L., 55 Brachet, J., 326,330(64) Bradley, M. E., 56 Bradshaw, R. A., 169, 170(39) Brady, R. J., 374 Brady, R. O., 144, 145(73), 146(73), 148 (711, 151(73), 152(73) Brady, W. T., 17 Brand, K., 373 Brand, L., 241(24), 242(24), 263(24) Bratvold, G. E., 558, 559, 580(29, 341, 561(34), 562(34), 564(34) Brdicrka, D., 282 Brennan, P. J., 491, 492(44) Breslow, R., 212 Bright, H. J., 533 Brindley, D. N., 158, 159(17), 160(17) Brink, N. G., 144 Britten, R. J., 510 Brock, D. J., 283 Brock, D. J. H., 240, 241(16), 242(16), 244(16), 262(16), 283 Brodie, A. F., 492 Brodie, J. D., 178 Bromel, H., 335 Bromilow, R. H., 209, 220, 223, 224(76, 771, 225(77), 226(77) Brosemer, R. W., 460, 461(24), 462(24) Brostrom, C. O., 558, 561, 562, 563, 584 (30, 411, 568, 570(82), 572(82), 574 (41), 576, 577, 578(135) Brostrom, M. A., 561, 563(42), 568, 569 (%I), 570(90), 571(90), 573(90), 576 (90) Brown, C. S., 446 Brown, D. H., 57 Brown, G. M., 174, 175(47) Brown, K., 281 Brown, K. K., 496

AUTHOR INDEX

Brown, M. S., 43 Brown, P. R., 309, 310(33), 311(25, 331, 312, 314(25), 315(25), 316(25), 332 (33) Brownlow, E. K., 385 Bruice, T. C., 201(4), 202, 225(4), 422 Brummond, D. O., 57 Brunner, G., 282 Bruno, R., 55 Brunswick, D. J., 274 Bryant, R. G., 368 Bublitz, C., 488, 489(10, 12), 490(10, 121, 493(10, 12), 494(12), 495(10, 121, 496 (10, 121, 497(10, 121, 499(10, 12), 500(12), 501(10, 121, 502(10) Buc, H., 245, 246(48), 263(48), 265(48), 267(48), 530 Buchanan, B. B., 83 Buchanan, C., 59, 60(93) Biicher, T.,.301, 335, 336(7), 337(3), 338, 340(3, 71, 341(3, 71, 342(7), 346(7), 347(2, 7), 348(7), 349(7), 371, 384, 512 Buchhawat, B. K., 54 Bucovaz, E. T., 9(97), 10, 13(97) Buehner, M., 454 Buist, G. J., 231 Bulcke, J. A., 394 Bunton, C. A., 202, 205, 207, 231, 232, 233(92, 951, 234(95) Burch, H. B., 56 Burdon, R. H., 24, 25(174) Burger, A., 386 Burma, D. P., 58, 311(48), 313 Burnett, G., 555 Burnham, B. F., 332 Burton, C. J., 9(101), 10, 13(101) Burton, D. N., 160 Burton, K., 337 Busch, D., 356, 359(15), 379(15) Butcher, E., 565 Butcher, R. W., 278,574 Butterworth, P. H. W., 160, 178, 186(56) Bygrave, F. L., 357, 369, 370(43)

C Caderata, R. L., 544 Calderon, R. O., 59, 60(96)

AUTHOR INDEX

Calendar, R., 9(94, 951, 10, 13(94, 95) Calkins, D., 572, 577(104), 579(104) Callaghan, 0. H., 280, 298(14, 15), 299, 3ofL Calvin, M., 337 Cameron, E. C., 77, 86(24), 89(24), 90 (24) Campagnari, F., 9, 10(54), 12(54) Campbell, J. W., 344, 348(42), 359 Canellakis, E. S., 22(184, 185, 186, 1881, 24, 25(170), 26(180), 310(38a, 38c), 312 Canellakis, Z. N., 22(185), 24, 25(170) Cannon, J. R., 144 Cantero, A., 55 Cantoni, G. L., 9(105), 10(64), 13(64, 105), 123, 124(11), 125(14), 127(14), 128(14, 28), 129(11, 14, 22), 130(14, 22, 23), 133(10, 141, 134(14), 135(11, 14), 136(41), 138(14), 139 Cantrenne, H., 6(29), 7, 12(29) Cantz, B., 384 Caputto, R., 59, 60(96), 245 Carbonell, J., 357, 374(49), 376(49), 377 (49) Cardini, C. E., 57, 58,245 Carlson, C. W., 460, 461(24), 462(24) Carlson, D. M., 53 Carmeli, C., 371 Carminatti, H., 356, 357, 372, 373, 374 (50), 375, 376(123, 1431, 377(123), 378(122, 123, 1431, 379(50) Carr, S., 377 Carroll, W. R., 17 Carter, J., 117 Cartier, P., 354 Cashel, M., 96 Cassells, A. C., 74 Castanera, E. G., 144, 145(73), 146(73), 148(73), 149(73), 151(73), 152(73) Cattaneo, J., 110 Cerletti, P., 281 Chaimovich, H., 9, 10(65), 13(65), 205, 232, 233(95), 234(95) Chambaut, A. M., 573, 576(112) Chamberlin, M., 21, 22(156) Chambliss, G. H., 374 Chan, T. L., 310(36), 312 Chang, G. G., 143

Changeux, J. P., 79, 87(29), 89(29), 267, 373, 526, 547(50) Chapeville, F., 9, 10(61, 66), 13(61) Chapman, A., 271, 273 Chapman, A. G., 106, 114(63) Chatman, V. B., 365 Chegwidden, R., 427 Chelala, C. A., 49 Cheldelin, V. H., 491, 492(35) Chen, L. J., 567, 570(74), 572(74), 577 (741, 578(74) Chen, S., 62, 63(124) Chen, S.-H., 340 Cheng, Y. C., 309, 310(22, 24), 311(25), 313(22), 314(22, 24, 25), 315(22, 24, 251, 316(22, 24, 251, 317(22), 319, 321(22, 241, 322(22, 241, 325(24), 326 (24), 329(24), 331(24), 332(24), 333 (22) ChCruy, A., 309, 310(15), 315(15), 316 (15), 321(15), 322(15), 325(15), 326 (151, 329(15), 330(15) Chesterton, C. J., 178 Chien, J. R., 44, 46(274), 47(274) Chiga, M., 280, 324, 325 Chin, B., 281 Chiriboga, J., 58 Chiu, C.S., 281, 282(28), 289(28), 291 (29, 296(28), 297(28) Chojnacki, T., 58, 66(86),69(86) Chou, T. C., 128, 130(30), 131(30), 132, 133, 134(30), 135(30), 136(30), 137, 138(30), 139(30) Chousterman, S., 9, lO(66) Ciardi, J. E., 41, 43 Clames, R. B., Jr., 491, 492(40) Clark, B., 488, 489(16), 493(16) Clark, J. B., 263(122), 264, 267(122) Clark, J. M., 9(93), 10, 13(93) Clark, J. R., 432 Clark, R. A., 365 Clark, R. B., 519 Cleland, W. W., 70, 138, 249, 250(63), 309, 315(16), 319(16), 320(16), 322 (16), 325(16), 326(16), 327(16), 328, 368, 396, 411, 414(72, 102), 415, 416, 418(104), 465, 486, 501 Clement, G. E., 185 Coch, E. H., 142 Cochran, D. G., 281, 298(26), 299(26)

588 Cohen, G. N., 520(7, 9, lo), 511, 512, 513(17), 514(17), 515(19), 516, 517 (%), 518(25, 29), 519(29, 33, 34), 520 (25, 29), 521(25, 29), 522(29), 523 (28, 291, 524(28), 525(25), 526(25, 49), 527(28, 45), 528(45), 530(45), 531(45), 532, 533(29), 536(29), 537, 538(57), 539(22, 57), 540, 542(16), 543, 544, 545(69), 549(64), 551(66), 552(66), 553(19), Cohn, J. A.,185 Cohn, M.,71, 295, 303, 304, 360, 362, 364, 368, 369, 370(109), 381(109), 405 (831, 406, 407, 408, 411, 412(94), 426 (941, 428, 430(116), 437(94), 438(94, 116), 439(137), 440(89, 1371, 441 (89, 94, 143), 442(94), 444(144), 449 (83, 84, 94), 450(89), 453(94) Cole, H. A.,115 Coleman, J. E.,213 Colli, W.,285 Collin, R.L.,211,212(34) Colomb, M. G.,309, 310(15), 315(15), 316(15), 319, 321(15), 322(15), 325 (15, 591, 326(15), 329(15), 330 Colowick, S. P., 27, 252, 280, 293, 301 Colvill, A. J. E., 24, 25(173) Colwell, R.R.,108 Condliff, P.,26 Conn, R.B.,396 Contopoulou, R.,82 Conway, M.M.,258 Coon, M. J., 358, 359(52), 360(55), 364, 365(83), 370(55), 376(55), 382(55, 83) Cooper, D. V.,339 Cooper, R. A., 558, 559(28), 563(28) Cooperman, B. S., 228, 229, 274 Coore, H.G.,566 Coputo, A., 357 Corbin, J. D., 561, 563(42), 568, 570(82), 572(82), 574, 576 Corbridge, D. E. C., 203,213 Cordes, W. V.,185 Cori, C. F.,274,277,558 Cornish, A., 357, 374(35) Cottam, G. L.,358, 359(52), 364, 365, 369, 370(110), 382(83) Coultate, T. P.,247, 248,262(51, 581, 266 (51)

AUTHOR INDEX

Coulter, A. W., 128, 129(33), 130(33), 135(33), 136(33), 137(33) Courte, C., 563, 569(87), 570(87), 572 (871, 573(87), 577037) Courtright, J., 492, 494, 495(61), 501 (61) Covitz, F., 217 Cowie, D. B.,510 Cox, D. J., 519 Cox, J. R.,201 (3), 202,219 Cox, R.,135, 138(44) Cozzarelli, N.A., 492 Crabtree, G. W.,312(56), 313 Cramer, F.,9(86), 10, 13(86) Crane, R.K.,278,280,295 Crawford, I. P.,108 Creighton, T.E.,522 Criddle, R. S., 158, 159(13), 172(13) Crisa, W. E.,280, 282, 287, 289(19, u)), 291(19, 20), 292(20), 296(19, 201, 297 (19, 201, 298(19), 299(19) Cronvall, E.,9(85), 10, 13(85) Crook, E. M.,431, 432, 445(122), 446 (120) Cross, M.E.,579 Cullen, J., 190 Cunningham, G.N.,518,519 Cunningham, L. W.,394, 432, 438(58), 446, 449(58) Curti, B., 85 Curtino, J. A., 59,60(96) Cutinelli, C., 82,570 Cutolo, E.,52 Czok, R.,337, 384

D DaCosta, W. A.,399 Dahlberg, D., 61 Damotte, M.,110 Dance, N.,432 Danenberg, P.V.,9(104), 10, 13(104) Danforth, W.F.,301 Danforth, W.H., 265,558 Daniel, V., 22(165, 1661, 24, 25(165, 166, 177) Danishefsky, I., 59 Dankert, M.,74 Darnall, D. W.,473 Dastugue, B.,580

AUTHOR INDEX

Datta, A., 232,233(103) Datta, P.,545, 546(67), 551, 552 d’Auzac, J., 366 Davidoff, F., 377 Davie, E.W.,574 Davies, J. E.,28, 29(194), 30(194) Davis, C., 371 Davis, J. J., 2, 3(3), 6, 12(3) Dawson, D. M., 232, 233(108), 384(16), 385, 386, 387, 389(16), 390(16), 391 (16), 393(16), 395(32), 397(16), 400 (16), 402(16), 403(16), 408(16), 409 (16),431(16), 445(32) Deal, S.J., 491,492(40) Deal, W.C., Jr., 358, 360, 370(63) Deamer, D. W.,385 De Bruin, K.E., 213 de Caputto, D. P.,31(218), 33 de Caputto, R., 31(218), 33 Decker, K.,67 DeCrombrugghe, B., 492 Decsi, L., 581 de Fekete, M. A. R., 56, 57, 58 Degani, C., 232, 233(96), 234(96) Degtar, R. G.,459, 461(11), 462(11) De La Haba, G., 126 Delaney, R.,167 DeLange, R. J., 558, 559(28), 560(28), 561(39), 563(28) Delluva, A. M., 510(6), 511 Delo, J., 161 DeLuca, C., 31(206, 2071, 32 Deluca, M., 7, 11(51), 12(130), 19, 20 (51) De Moss, J. A., 522, 539(43) Dempsey, W.B., 117 Dennis, D. T., 247, 248, 262(51, 581, 266 (51) Dennis, E. A., 216,217,219 Denton, M.D., 44 Denton, R. M., 262(112), 264, 277, 278, 566

DeRitis, G., 281 De Robichon-Szulmajster, H., 513, 514 (19),515(19), 553(19) der Terrosian, E., 431, 432, 460, 461(15, 17), 462(15, 17, 361, 466(17), 467 (17), 469, 470(65, 661, 471(36), 472

589 W ) , 475(36), 476(36), 477(36), 478 (17,761, 479, 480(76) Desvages, G.,469, 470(66) Devalle, J., 379 DeVito, P. C., 35(221), 36, 37(234), 38 Diago, K.,18 DiBella, F.,574 Dickerman, H., 123 Dickinson, D. B., 93,94(47) Diefenbach, H., 71, 437, 440, 441(143) Diekmann, M., 22(164), 24, 25(164) Diesterhaft, M., 357 Diezel, W., 254,265,267(126) Di Jeso, F., 459, 461, 462(36, 38), 471 (36), 472(36), 475(36, 38), 476(36), 477(36) Diller, R. F. B., 9(96), 10, 13(96) Dils, R.,160 Dirheimer, G.,365 DiSabato, G.,208, 232(24), 233(24) Distiche, A., 384(10), 385 Dixon, G. H.,556, 568(10), 572(10), 573 (lo), 574(10), 575(10), 576(10) Dixon, M.,329 Doelle, H.W.,249, 250(71), 263(71), 264 (711, 267(71) Doherty, M. D., 384(14), 385, 390 Domagk, G.,360 Domschke, W.,360 Donnel, G.N.,69 Donnicke, M., 268 Donohue, J., 213 Dorizzi, M.,9(90), 10, 13(90) Dorsey, J. A., 178,189 Doudoroff, M., 82,517 Doughty, C. C., 504, 505(78), 506(78), 507(78), 508(78) Drago, R.S.,213,215(44) Dreyfuss, J., 35(221, 2261, 36, 37(234), 38 Drummond, G. I., 563, 564, 565, 573 Duba, C.,177, 178(55), 181(55), 182(55) Dubin, J. C.,356 Dubnoff, J. W.,123 DuBe, E ., 344,348(42), 359 Duffy, M. J., 574 Dunaway, G. A., Jr., 374, 376(139) Duncan, L., 564 Dunikoski, L. K., Jr., 220, 228

590

AUTHOB INDEX

Ennis, H. L., 115 Dunhcher, M. P., 9(!%), 10, 13(98) Duntze, W.,240(31), 241, 260(31), 278 Ennor, A. H., 410, 431, 460, 461(19), 462 (19, 32, 37), 463(37), 464(32), 471 (31) (19, 32, 34), 472(19, 32, 341, 474(19), Durell, J., 125, 128(22), 129(22), 130 475(19, 32, 341, 476(19, 32, 34), 477 (22) (19, 32) Dvornikhova, P. D.,459, 461(11), 462 Eppenberger, H. M.,232, 233(108), 384 (11) (W, 385, 386, 387, 388, 389(16), 390 Dyson, J. E., 240(34), 243 (16), 391(16), 393(16), 395(32), 397 (161,400(16), 402(16), 403(16), 408 (161, 409061, 431(16), 445(32) E Eppenberger, M. E., 386,387 Erenrich, E. S.,236 Ebashi, S., 288, 562, 563(48) Erikson, R.L.,21, 23(149) Ebel, J.-P., 9(106), 10, 13(106), 365 Eriksaon, A. W.,283 Ebner, E., 43, 44 Ebner, K.E., 53, 59(20), 62(20), 63(124), Erlichman, J., 568, 570, 571(83), 572 (83,98), 577 66(20) Erst-Fonberg, M.L.,161 Eckstein, F., 213, 236 Eskola, M.-R., 283 Edelhoch, H., 232,233(98) Espada, J., 31(209, 211), 32, 74 Edelman, G.M., 47 Estabrook, R. W.,493 Edelmann, P. L.,75, 110, 118(75) Edlund, B., 232, 233(106), 308, 309(6, Evans, H.J., 362,366,371 lo), 311(10), 312(6), 313(6), 3 W 6 , Evans, M. C.W., 83 lo), 3156, 101, 316(10), 319(6, 10) Evans, P., 340, 341(30), 344(30), 345 (301,351(30) Edmonds, M., 24, 25(172) Eveleigh, J. W., 384, 389(43), 390(43), Eggerer, H.,189 391, 392(45), 393(43), 469, 470(67) Eggleston, L. V., 298, 299, 302, 374, 379 Ewing, L. L.,569, 570(92), 572(92), 577 Eggleton, G. P., 458 Eggleton, P., 458 (92) Eyer, P., 241, 244, 245 Ehrensvard, G.,82, 510 Eidels, L., 75, 78(17), 83(17), 110, 118 Eyzaquire, J. P.,9(93), 10, 13(93) (75) Eigner, E. A.,9,13(79) F Eil, C.,574 Falcoz-Kelly, F., 517, 518(29), 519(29), Elhilali, M.M.,386 520(29), 521, 522(29), 523(29), 533 Ellfolk, N.,48 (29), 536, 537(57), 538(57), 539(57), Ellingson, J. S.,61 541 Elliot, W.H.,12(28) Elodi, P., 460, 461(12), 462(12), 466(12), Fall, L.,106,114(63), 286 Fanshier, E.W.,52, 70(4) 467(12) Elovson, J., 161, 166, 171(33), 173(33), Farber, S.J., 207 Fareed, G. C.,44, 46(272), 47(272) 174(46), 175(27) Farina, M. V., 311(44), 313, 325(44) Elsasser, S.,43 Farnham, W.B.,213 Emery, R. S.,54,66(24) Farrell, F.J., 230 Engelhardt, V. A., 269 Fasiolo, F., 9(106), 10, 13(106) Engers, H.D., 338,339(17) Engstrom, L., 232, 308, 309(6), 312(6), Fattoum, A., 433, 434(128), 453(128), 454(128), 455(128), 481, 482(80, 82, 313(6), 314(6), 315(6), 319(6, 18), 83), 483(821,485(82) 320(18), 326(18)

591

AUTHOR INDEX

Fedor, J., 229 Feiner, L., 45(276), 46, 47(276) Feingold, D. S., 59 Fellman, J., 283 Fendler, E. J., 202, 205, 231 Fendler, J. H., 205 Fenrych, W., 145 Ferdinandus, J., 263(122), 264, 267(122) Fernald, N., 312(51), 313, 319(51), 324 (51)

Fife, T. H., 202, 206(8), 221 Fildes, R. A., 282 Filler, R., 287 Filmer, D. L., 232, 233(93), 267, 526 Finch, L. R., 115 Fine, I. H., 384 Fink, S. C., 460, 461(24), 462(24) Finkelstein, J. D., 124, 126(16), 129(16), 134, 142, 143(40)

Fischer, E. H., 338, 556, 557, 558, 559 (13, 21), 560(13, 341, 561(13, 34), 562(32), 563(43), 564(34, 47), 572, 577(104), 579(104) Fish, D. C., 505 Fisher, E. A., 379 Fiske, C. H., 458 Fitzgerald, D. K., 62, 63(124) Fitzgerald, P. J., 579

Fitzsimmons, J. A. E., 390 Flanders, L. E., 351(21), 356, 376(21) Flashner, M., 359, 360(55), 370(55), 376 (55), 382(55)

Flatt, J. P., 278 Flavin, M., 141, 142(54,55) Fling, M., 509, 510 Flodgaard, H., 55 Floyd, B. F., 399 Flynn, I. W., 357, 361, 374(40) Flynn, R. M., 6, 12(13, 141, 13(13) Focant, B., 384(13), 387, 390(13), 391 (13), 392, 393(13), 395(13, 341, 400 (13), 408(47), 409(47), 419(47) Folk, W. R., 9(99), 10, 13(99) Folkers, K., 144 Follmann, H., 232, 233(109) Fondy, T. P., 493 Fontana, J. A., 568, 570(85), 572(85)

Ford, G. C., 454 Forest, P. B., 270, 271(147), 273(147) Fosmire, G. J., 255 Franke, J., 54, 62(22) , 63 (22), 65 (221, 66(22), 70(22)

Franklin, R. M., 21, 23(149, 150) Franz, G., 57, 59 Freedberg, W. B., 492 Freedland, R., 379 Freese, E., 357 Frenkel, R., 240, 241(10), 242(10), 262 (10)

Freundlich, M., 514, 544 Frey, P. A., 234 Freyer, R., 265, 267(126) Friedberg, F., 395, 399 Frieden, C., 253, 259, 269(100) Friedman, D. L., 556 Friesen, A. J. D., 563, 564 Fritz, P. J., 374, 379(133) Froede, H. C., 270 Frflholm, L. O., 14 Fromm, H. J., 252 Fujikawa, K., 14 Fujimoto, T., 60 Fukasawa, T., 59,60(89) Fukunaga, K., 53, 62(21), 63(21), 65 (21), 66(21), 67(21), 69(21), 70(21)

Furlong, C. E., 75, 77(18, 19), 81(18, 191, 118(19)

Furth, J. J., 21, 22(167), 24, 25(167)

G Gabriel, T. F., 47 Gaffney, T. J., 461, 462(32), 464(32), 471 (32, 34), 472(32, 34), 475(32, 341, 476 (32, 341, 477(32)

Galsworthy, S. B., 142 Gammack, D. B., 385, 466, 467(59) Gancedo, C., 43, 492, 497(47) Gancedo, J. M., 261, 492, 497(47) Gander, J. E., 65, 66(130), 67(130) Gangali, N. C., 312(50), 313 Ganther, H., 85 Garces, E., 309, 315(16), 319(16), 320 (16), 322(16), 325(16), 326(16), 327 (161, 328

AUTHOR INDEX

Garfinkel, D., 269, 378 Garland, P.B., 265, 277(127), 490 Garrard, L. A.,262(120), 264, 266(120) Garren, L. D.,567, 568,570, 572(84, 96), 574(79), 577(96) Garson, 0. M., 340 Gatica, M.,9, lO(57, 65), 12(57), 13(65) Gaudy, E.T.,496 Gauer, J., 254 Gay, D.C.,217 Gefter, M.,47 Gehring, V.,183 Geiduschek, E.P.,21 Geismann, H., 212 Gell, J., 310(33), 311(33), 312, 332(33) Gellert, M.,45(275), 46, 47(275) Gentner, N., 75, 94(ll, 131, 97(11, 13, 521, 99(11, 13, 521, 100(52), 101(57), 102(11, 571, 103(57), llO(11, 521, 113 (52, 56, 571, 114(11, 56, 57) George, P.,2, 3(6) Geraci, G., 270 Gerday, C.,384, 389(44), 390(44), 391, 393(5), 400(5), 448(5) Gergely, J., 558 Gerschenson,, L.,379 Gest, H., 545, 546(67) Gevers, W.,6(43, 44, 451, 7, 9, 10(68), 11(43, 44, 45, 68, 75), 12(46), 14(75), 15(75) Ghosh, H. P., 33, 77, 86(21, 22, 23, 87 (21, 22, 231, 88(22, 231, 89(23), 93 (23) Gibbons, A. P., 57 Giblett, E.R., 283, 340 Gibson, D. M.,309, 310(29, 34), 312, 324 (29) Gilchrist, M., 184, 209, 210(27), 211(27), 232(29), 233(29) Gill, G. N.,567, 568, 570, 572(84, 961, 574(79), 577(96) Gillespie, P.,215 Gillett, T.A.,63,69 Gilvary, C.,396,405(66) Ginsburg, A., 40, 41, 43(242), 44(258, 266) Ginsburg, B., 45(276), 46, 47(276) Ginsburg, V.,33, 36(215), 52, 53, 62(19), 66(19), 70(4, 191,74 Giovanelli, J., 6, 12(27)

Gitlin, J., 284 Gitzelmann, R.,68 Giunta, C.,55 Glaser, L.,54, 117 Glass, J., 85, 372 Glasaiou, K.T., 58 Glaze, R. P., 309, 310(28, 37), 312, 313 (281, 314(28), 321(28), 325(28), 326 (28), 329(28), 330(28) Goedde, H. N., 283 Goff, C. G., 47, 48(284), 49(284) Goffeau, A., 309, 322(17), 325(17), 326 (17), 329(17), 330(64), 331(17), 333 ( 17)

Goldberg, M. E.,522, 527(44), 538(44) Goldberg, N.D.,571 Goldberg, N.O.,68 Goldemberg, S. H., 59 Goldfeder, A.,281 Goldfine, H., 158, 159(15) Goldman, P.,156, 188(1) Gonzales, C., 572, 577(104), 579(104) Goodman, D.B. P., 574 Gopinathan, K. P., 492 Gorna, M.,145 Gosselin-Rey, C., 340, 384(1), 385, 388, 389(44), 390(44), 391, 393(5), 400 (5), 448(5) Gottesmann, M.E.,22(188), 24, 25(170), 26 Gottschalk, E.,9(86), 10, 13(86) Govons, S.,94, 97(52), 99(52), 100(52), 110(52), 113(52), 118(73, 75) Grant-Greene, M. C., 395 Graves, D. J., 556, 559(13), 560(13), 561 (13), 562 Gray, E., 547, 548(73), 549(72) G ~ D. E., ~ 6, 12(19) ~ ~ , Greenawalt, J. W.,310(36, 43), 312, 313, 333(43) Greenbaum, A. L., 504 Greenberg, D.M.,504 Greenberg, E.,33, 75, 77, 86(24), 89(24), 90(24), 94(11, 13), 97(11, 13, 521, 99 (11, 13, 521, 100(52), 102(11), 110 (11, 52), 113(52), 114(11), 115(78), 117(78), 118(75) Greenberg, G. R., 312(54), 313 Greenberg, R.,488, 489(3)

AUTHOR INDEX

593

Greene, R. C., 127, 128, 129(27), 130

(271, 131(27), 132(27), 133, 134(27), 135(31), 136(27), 137(27), 138, 139 (27), 141, 142(52, 56) Greengard, P. J., 27, 567, 568, 569(88), 571, 572(73, 88, 1051, 573(80), 574, 578, 581(77) Greenleaf, A. L., 403, 404(81), 405(81), 406(81) Greenspan, M. D.,189, 191, 193 Gries, F. A., 488, 489(13), 490(13, 22), 493(13, 22) Griffin, C. C.,241(24), 242(24), 250, 251 (73,263(24) Griffiths, D. E., 460, 461(19), 462(19), 471( 19), 472(191, 474(191, 475(19), 476(19), 477'39) Grisolia, S., 232, 233(98) Groot, G.S. P., 333 Gros, C.,516, 517(25), 518(25), 519(34), 520(25), 521(25), 525(25), 526(25), 542, 543 Grosjean, C. H., 9,lO(60) Gross, C.,9(89), 10, 13(89) Grunberg-Manago, M.,4(8), 5, 21, 22 (139), 23(139) Grunnet, N.,494, 497(60), 498(60), 499 (60),500(60), 503(60), 504(60) Grreschik, A.,340 Grzeschik, K. H., 340 Gumaa, K. A.,504 Gumport, R.I., 46,47(277) Gunsalus, I. C.,492 Gunther, H. L., 504, 505(78), 506(78), 507(78), 508(78) Gunther, S., 177, 178(55), 181(55), 182 (55) Gurin, S.,6 Gurley, L. R.,579 Gussin, A. E.S., 56 Gustafson, G., 54, 62(27), 67(27) Gustafson, G. L.,65, 66(130), 67(130) Guthrow, C.E.,Jr., 574 Gutierrez, R., 579 H Haag, U.,377 Haake, P.C.,204, 217, 232(13), 233(13) Haas, T.,379

Haavik, A. G., 160 Hackert, M.L.,454 Haddox, M.K.,571 Haeckel; R.,373, 375(127) Haessler, H. A., 488, 489(9), 493(9), 494

(91, 496(9) Hageman, E.,63 Hageman, J. G., 286 Hagenmaier, H., 127, 129(26) Hagmaier, V.,261 Hahn, P.,488,489(3) Hales, C.N.,277 Hall, M.A., 56, 57(57) Hall, Z. Q.,44, 46(274), 47(274) Hall, Z.W.,47 Halmann, M.,232, 233(96), 234(96) Hamer, N.K.,216,217 Hamilton, W.C.,213 Hammermeister, K.E.,564 Hammee, G. G., 362, 396, 413(68), 437

!@I, 451(68) Hamoire, G.,384(11), 385,388 Hancock, R. L., 129, 135(35), 142(35),

143(35) Hancock, W.S.,191, 193(88) Handler, P., 9(107), 10, 13(107), 31

(202,203,2041,32 Hanoune, J., 573, 576(112) Hansen, R. G., 52, 53, 64, 55(16), 02(16,

17), 63(16, 17), 65(16, 17), 66(16, 171, 67(16, 17), 68(16, 17), 69(17), 71 Hansen, R. W.,504, 507(77), 508(77) Hansford, R. G., 565 Hanson, J., 464 Hanson, R. L., 249, 250, 251(70), 252 (69, 70) Hanson, T.E.,241,245 Happold, F. C.,281 Harano, Y.,356, 359(16), 360(16), 374 (16), 376(16) Hardie, J., 77, 86(24), 89(24), 90(24) Hardy, S.J. S., 22(168), 24, 25(168) Harmey, M.A., 74 Harper, E.M.,52 Harris, E.J., 117 Harris, H., 282, 283 Harris, I., 177, 183(52) Harris, J. I.,183 Harris, J. S.,571

594

AUTHOR INDEX

Harrison, W. H., 485 Hartle, D. K., 571 Hartley, B. S., 9(100, 1011, 10, 13(100, 1011, 454

Hartmann, G., 275 Harvey, C. L., 47 Harwood, J. P., 563 Haschke, R. H., 557, 563 Hashimoto, T., 31(219), 35, 340, 341(31) Hass, L. F., 250, 370, 376(114) Hasselberger, F. X., 275 Hassid, W. Z., 52, 57, 59, 70(4) Hastings-Park, J., 177, 183(52) Hatch, M. D., 58 Hathaway, J. A., 247, 263(53), 267 Hauge, J. F., 491, 492(35) Hawker, J. S., 94 Hayaishi, O., 27, 47, 49(285), 259, 269 (101)

Hayakawa, T., 559, 561, 562 Hayashi, H., 9, 13(81) Hayashi, J. A., 504, 505(78), 506(78), 507(77, 78), 508(77, 78)

Hayashi, S., 491, 492(30), 493, 494(59), 496(59), 497(59), 498(59), 499(59), 500 (591, 501(59) Heber, U. W., 91, 92(43, 44, 45) Hecht, L. I., 24, 310(38b), 312 Heck, H., 185 Heck, H. d’A., 519, 521(36), 524(36), 535(36) Hedrick, J. L., 118 Heider, H., 9(86), 10, 13(86) Heilmeyer, L. M. G., Jr., 557, 563 Heinrich, C. P., 42 Heinrikson, R. L., 41 Heldt, H. W., 385 Hele, P., 6, 1 2 W Hempfling, W. P., 117 Hemphill, R. M., 269 Hems, R., 298, 299, 301, 302, 308, 310 (41, 311 Henderson, E., 551 Hendrickson, R. L., 9(100), 10, 13(100) Hennig, S. B., 41, 43, 44(258, 266) Heppel, L. A., 23,161 Herberg, L., 488, 489(13), 490(13), 493 (13) Herbert, E., 24, 26(180), 310(38, 38b), 312

Heritier-Watkins, O., 59 Hers, H. G., 504 Hess, B., 276, 356, 358(13), 359(20, 52a), 360, 361, 373, 375 (127), 378 (1421, 379 Heumann, W., 355 Hevey, R. C., 206 Heyde, E., 421, 424(111), 427(111) Hickenbottom, J. P., 559, 561, 563(42), 573(35), 574(35) Hift, H., 6, 12(18) Hill, A. J., 79, 87, 89 Hill, R. L., 158, 159(16), 166(16), 167, 168(16), 169(16), 170(16) Hilmoe, R. J., 23 Himms-Hagen, J., 488, 489(14) Hinae, H., 43 Hirs, C. H. W., 179, 186(67) Hirsch, A. H., 570, 572(98) Hirsch, M. L., 510(7, 8, 9, 101, 511 Hirsch, R., 9(85), 10, 13(85) Hirschbein, L., 49 Hirschfield, I., 9, 12(78) Hirsh, D. I., 9, 10(63), 13(63) Hizukuri, S., 57 Hnilica, L. S., 579 Ho, E. S., 561, 566(41), 574(41) Ho, R. J., 278, 474 Ho, W., 262(113), 264 Hoagland, M. B., 31(208), 32 Hochachka, P., 357, 374(45, 471, 375 (451, 376(45) Hockberg, A. A., 575 Hodgkin, D. C., 123, 144(7) Hodgson, G., 359 Hofer, H. W., 241, 244, 245(39), 259, 268, 270 Hofer, M., 117 HolTbauer, R. W., 356, 359(15), 379(15) Hoffman, P., 215 Hoffmann, E., 254, 265, 267(126) Hoffmann-Ostenhof, O.,280 Hofstatter, T., 341, 344(35) Hogdson, G., 344, 348(42) Hohorst, H. J., 337 Holbrook, K. A,, 232, 233(101) Hollenberg, P., 358, 359(52), 360, 370 (551, 376(65), 382(55) Holloway, C. T., 141, 142(52, 56)

595

AUTHOR INDEX

Holmsen, H., 376 Hololian, P. D., 493 Holtzcr, R. L., 324 Holz, G., 4(9), 5, 492, 493(46), 494(46), 495(46), 496(46), 499(46), 501(46), 502(46) Holzer, H., 41, 42, 43(246), 240(31), 241, 260(31), 261, 278(31) Hornmes, F. A., 281 Honjo, T., 47, 49(285) Hooton, B. T., 384(15), 388(15), 389 (15), 390(15), 393(9), 400(9), 402 (15), 443(15), 445(15), 446(15) Hopper-Kessel, I., 189 Homing, D. P., 222, 225(75), 226(75) Horowitz, N. H., 509, 510 Horsley, W. J., 21, 22(136) Hoskins, D. D., 248, 262(62), 266(62) Hosoi, K., 562, 563(48) Hossler, F. E., 310(35), 312, 318(35), 319 (351, 333(35) Houck, B. N., 241(24), 242(24), 263 (24) Howell, R. R., 68, 69(138) Hsia, Y. E., 145 Hsieh, H.-S., 340 Hsu. D. S., 354 Huber, W. A., 56 Hubscher, G., 488, 489(16), 493(16) Hucho, F., 565 Huennekens, F. M., 144, 145(75), 147 (75, 831, 148(75), 150(75), 151(75), 152(75) Hughes, D. E., 115 Hulrne, E. C., 249, 252(65), 269 Humeres, E., 202 Humphreys, T. E., 262(120), 264, 266 (120) Hunkeler, F. L., 559, 562, 563(46, 49), 573(35), 574(35) Hunsley, J. R., 356, 358(14), 369, 373, 375(129), 378(129) Hunter, G. J. E., 491 Hurd, S. S., 559 Hurst, J. K., 396, 413(68), 437(68), 451 (68) Hurwitz, J., 21, 22(138, 162, 1671, 23(153,

154), 24(148, 1621, 25(162, 1671, 26, 45(276), 46, 47(276), 49 Huston, R. B., 562, 563(46) Huttunen, J. K., 574

I Ibscn, K., 356, 379 Ichihara, A., 504 Ide, M., 259, 269(101), 356, 374(28), 375 (28) Ikura, Y.,257 Imahori, K., 240(32), 243, 263(32), 264 (32), 266(32), 267(32) Irnarnura, K., 356 Irnsande, J., 31 (2031, 32 Inamasu, M., 577 Ingraharn, J., 110, 118(73) Ionino, S., 14 Irreverre, F., 124, 126(16), 129(16) Isselbacher, K. J., 55, 68, 488, 489(9), 493(9), 494(9), 496(9) Itoh, H., 14 Ives, D. H., 311(45), 313 Iwai, H., 577 Iwatsubo, M., 533, 534

J Jachirnoaicz, M., 60 Jacob, E. J., 178, 186(56) Jacob, F., 60 Jacob, J.-L., 366 Jacob, M., 309, 310(29), 324(29) Jacobs, G., 438 Jacobs, H., 385 Jacobs, H. K., 308, 315(12), 320(12), (17), 385, 394, 395(61), 399(63a), (63a), 401(17, 63a), 402(17), (63a), 418, 419(106), 420(106), (106), 447, 448(150), 466 Jacobs, N. J., 491, 492(39) Jacobsen, D. D., 250, 370, 376(114) Jacobson, K. W., 356, 361 Jacquemin-Sablon, A., 44, 46(272), (272) Jaffe, E. R., 340 James, E., 403, 406, 407(86), 408, (101), 415, 416(101), 417(86),

384 400 403 427

47

414 418

596

AUTHOR INDEX

(1011, 422(101), 436(101), 437(101), 464, 465(49) James, H. L., 9(97), 10, 13(97) Jameson, G.W.,209 Jamieson, G. A., 125, 126, 130(23) Jampel, E.,203 Janin, J., 516, 517(25), 518(25, 291, 519 (29), 520(25, 291, 521(25, 291, 522 (29), 523(28, 291, 524(28), 525(25), 526(25), 527(28, 45), 528(45), 530 (45), 531(45), 532, 533(29), 534, 535, 536(29), 537 Janjigian, J. A., 29 Jarabak, J., 508 Jard, S., 567, 572(78) Jencks, W. P.,4(11), 5, 6, 9, 11, 12(20, 21), 156, 184, 185, 208, 209(25), 210 (27), 211(27), 232(24, 291, 233(24, 29), 237 Jergil, B., 556, 568(10), 572(10), 573(10), 574(10), 575(10), 576(10) Jimenez de Asua, L.,356, 357, 372, 373 (123), 374(50), 375, 376(123, 1431, 377(123), 378(122, 123, 1431, 379(50) Jirgenssons, B., 344 Johnson, A. W.,144 Johnson, E.,357, 374(35) Johnson, E. M., 567, 581(77) Johnson, G.S.,358, 360, 370(63) Johnson, J. C.,54 Johnson, R.E.,35(227), 36 Joklik, W.K., 308, 310(3, 31), 312(3, 311, 315(31), 320(31), 321(3, 311, 324, 325(3, 30, 329(31) Jokura, K., 59, 60(89) Jolles, G.R.,232, 233(105) Jones, E.A,, 277 Jones, M. E.,6, 12(13, 141, 13(13) Jones-Mortimer, M. C.,35(224), 36, 42 (225) Joseph, D. R., Q(91,921, 10, 13(91, 92) Joshi, V. C.,178, 179, 180(63), 183(63), 186(62, 64) Jungas, R. L.,666

K Kabsch, W., 295 Kachmar, J. F.,366,372

Kaffarnick, H.,284 Kagi, J. H., 393 Kahlbacher, B.,96 Kaiser, E.T.,219 Kaji, A,, 252 Kalckar, H. M., 33, 52, 53, 54, 59, 62 (31), 280,488 Kaldor, G.,284 Kalnitsky, G.,85 Kalyankei, G. D.,6(36, 37, 38, 39), 7, 12(36, 39) Kamiynma, M., 580 Kammen, H.O.,22(184, 187), 26 Kamogawa, A., 54, 62(28), 66(28) Kamp, H., 356, 374(26), 375(26), 376(26) Kamp, R. G.,558, 559(28), 563(28) Kamper, D.,371 Kamper, J., 144 Kampf, S. C.,488, 489(7), 493(7) Kanazawa, K., 74,92(4) Kanazawa, T.,74, 92 Kandler, O.,74 Kaplan, N.O.,31(206), 32, 232, 233(108), 384(16), 385, 386, 387, 389(16), 390 (l6), 391(16), 393(16), 395(32), 397 (IS), 400(16), 402(16), 403(16), 408 (161, 409(16), 431(16), 445(32), 460, 461(13, 18), 462(13, 181, 466(13, 181, 467(13, 181,468(18), 469(13, 18), 470 (131,478(18), 479(13, IS), 480(18) Karibian, D., 491, 492(32) Karnovsky, M. L.,491, 492(32) Kassab, R.,431, 432, 433, 434(128), 443 (129), 453(128), 454(127), 455(128, 129), 460, 461(14, 15, 17, 29, 301, 462 (14, 15, 17, 29, 30, 31, 38), 464(31), 465, 466(17, 31), 467(17, 31, 58), 468 (14, 30, 311, 469, 470(65, 661, 471 (31), 472(31, 71), 475(38, 711, 476 (711, 477(71), 478(14, 17, 29, 761, 479(18), 480(71, 781, 481, 482(14, 52, 71, 78, 80, 82, 831, 48362, 78, 821, 484(52), 485(82) Kates, M., 491, 501(34) Kato, I.,47,49(285) Katunuma, N.,48 Katz, I., 212 Katze, J. R., 9, 13(81, 84)

AUTHOR INDEX

Kauder, E. M., 492, 493(46), 494(46, 481, 495(46), 496(46), 499(46), 501(46), 502(46) Kauss, H., 74 Kawade, Y.,9, 12(52), 13(52) Kawahara, K.,119 Kay, G.,358 Kaye, A. M., 26 Kayne, F. J., 85, 355, 361, 362(71), 363 (71),366(8), 367, 368(8),370(8), 371 (701, 372(8), 373(70), 378(124) Kayne, M. S., 358 Kee, A.,250, 251(72) Kelly, G.J., 262(116, 117, 118, 1191,264, 266(116, 117, 118) Kernp, R.C.,560,561(39) Kernp, R. G.,240, 241(15), 242(15), 254, 258, 259, 262(15), 266(15), 269(82), 270, 271, 273(147, 148, 149, 150) KendrickJones, J., 287 Kennedy, E. P., 115, 488, 489(12), 490 (12), 493(12), 494(12), 495(12), 496 ( 12), 497( 12), 499( 12), 500( 12), 501 (121,555 Kenyon, G.L.,365,403, 404(81), 405(81), 406(81) Keppler, D. 0. R.,67 Kerr, D.S.,141, 142(54, 55) Kerson, L. A., 378 Kerwar, S.S.,145, 148(79) Kerwin, T.D.,298,299032) Ketelaar, J., 212 Keto, A. I.,384(14), 385 Keutel, H.J., 384(17), 385, 394, 395(61), 399(63a), 400(63a), 401(17, 63a), 402 (17), 403(63a) Kesdy, F. J., 185 Khan, S. A., 207, 209(33), 210, 222, 223, 224(76, 771, 225(75, 76, 771, 226(75, 76, 77) Khoo, J. C.,283 Khorana, H.G.,218 Kikuchi, G.,82 Kirnberg, D.V.,377 Kirnhi, Y.,21, 22(140), 23(140) King, C. A., 570, 577, 578(135) King, T.E., 491,492(35) Kingdon, H.S.,41, 42(248) Kinoshita, J. H., 249, 262(67), 264(67)

597 Kinoshita, S., 552 Kipnis, D.M.,277 Kirby, A. J., 201,202, 205(7), 206(7), 208 (23), 209(25, 26, 32, 331, 210, 211 (7), 212, 220, 222, 223, 224(76, 77), 225(32, 75, 771, 226(32, 75, 77) Kirjarinta, M., 283 Kirk, M.R.,74, 92(4) Kirkland, R. J. A.,311(49), 313, 315(49), 320(49), 321(49), 329(49), 330(49) Kistler, W.S.,492, 503 Kjellstrorn, W.A.,230 Klaus, D.,284 Klee, W.A., 139 Kleineke, J., 365 Kleinkauf, H.,6(40, 43, 44, 451, 7, 9, 10 (681, ll(40, 43, 44, 45, 68, 75), 12 (461, 14(75), 15(75) Klernperer, H. G.,22(186, 187), 24, 25 (1711,26 Klenow, H.,280,310(32), 312 Klethi, J., 281, 296(79), 297 Klingenberg, M.,385 Klingenburg, H.,341, 344(35) Klisacek, H.,215 Klots, C.,460, 462(23), 462(23), 466(23), 467(23, 611, 468(23), 474(23), 476 (23), 482(23), 484(23) Klotsch, H., 4(9), 5 Kluger, R.,217 Klungsoyr, L.,286 Knight, B. L.,488, 489(8), 490(8) Knivett, V. A., 190 Knop, J., 53, 62(17), 63(17), 65(17), 66 (17), 67(17), 68(17), 69(17) Knowles, J. R., 9,13(81) Knox, W.E.,490 Koch, J. P.,491, 492(30) Koedam, J. C.,396 Koeppe, 0.J., 9 Kohno, T., 522,538(42) Koike, M.,6(50), 7, 17 Kokesh, F. C.,234 Koler, R.D., 356 Kondo, M.,145 Kong, C. M.,309, 311(26), 314(25), 315 (25), 316(25) Konigsberg, W.,9, 13(W Koniuszy, F.R.,144

598

AUTHOR INDEX

Kono, N., 240, 241(17, 191, 242(17, 191, Krishnaswamy, P. R.,9, lO(55) 243(17), 252(19), 255(19), 257(19), Krueger, B. K., 572, 573(105) Krug, R.,n(1671, 24, 25(167) 258(19), 270(19) Kruger, J., 276 Koopmans, K., 212 Krystek, E., 241, 244, 245(39) Kopperschlager, G.,265, 267(126) Kreanowski, J., 266, 275(129) Korey, S: R.,4(8), 5 Kuby, S. A., 232, 233(106, 1071, 280, 291, Korman, E.F.,227 292(72), 293, 296(12), 297(12), 301(4, Korn, E. D.,488, 489(8), 491(18) 751, 308, 312(5), 314(5), 315(12), 316 Kornberg, A., 5, 9, 11, 21(71), 31(12, (ll), 320(12), 321(5), 324(5), 384 2051, 32, 281, 312(53a), 313 (17), 385, 389(41, 42), 390(42), 391 Kornberg, H. L., 357, 374011, 370(31), (411, 392(41, 421, 393(42), 394(42), 505, 506 395(3, 48, 52, 611, 396(64), 399(42, Kornfield, R.,57 63a, 73), 400(3, 63a, 73), 401(3, 17, Korzybski, T.,54 63a, 64), 402(17), 403(52, 63a, 731, Kosakowaski, M. H. J. E.,9(103), 10, 405(64), 407(73), 412(73), 413, 416 13(103) (73, 97), 418(64), 419(106), 420(73, Koschinsky, T., 488, 489(13), 490(13, n), 106), 421(64, 73, 97), 422(73, 97), 427 493(13, 22) (106), 428, 429, 430(113), 431(64), Koshland, D.E.,Jr., 98,103(54), 232,233 432(97), 443(64), 445(64), 446(64), (93), 267, 526 447, 448(150), 449(64), 463, 465(45), Kosicki, G. W.,366, 381(97) 466(45) Kosow, D.P., 18 Kuczenski, R.,359, 362, 363, 375(80) Koster, J. F., 354, 356, 374(4, 281, 375 Kuehn, G.D.,578 Kulbertus, H., 384(10), 385 (261,376(26), 379(4) Kumagai, H.,288 Kotel’nikova, A. V.,280 Krakow, J. S., 21, 22(136, 137, 184, 186), Kumar, S., 189 Kumon, A., 570 26 Kumudavalli, I., 384, 386, 389(8), 390 Krall, A. R.,364 (81, 39309, 400(8), 445(8), 447 Kratky, O.,359 Kunisawa, R.,82 Krause, E.G.,56 Kuo, J. F.,567, 568, 569(88), 572(73, 881, Kraut, J. A.,454 573(80, 105), 574,578 Krebs, E. G.,240, 241(6, 71, 242(6, 71, Kupiecki, F.,364, 365(83), 382(83) 243, 253(7), 254(7), 255(7), 260(7), Kurahashi, K., 14, 54, 59, 60(89), 62(28), 269(82), 270(7), 338, 556, 557, 558, 66(28), 311(46), 313 559(13, 21, 28), 560(13, 29, 341, 561 Kuramitsu, H. K., 550 (13, 34, 39), 562(34), 563(28, 42, 43, 49), 564(30, 34, 471, 566(12, 411, 568 Kurashina, Y., 27 (12), 569(89, 901,570(82, 89, 90), 571 Kurland, C.G.,22(168), 24, 25(168) (89, 901, 572(12, 82, 89, 901, 573(12, Kurooka, S., 240(27), 241, 242(27), 243, 253(27), 256(27), 263(27), 267(27) 35, 901, 574(12, 35, 41), 576(89, 90). 577(104), 578(90, 1351, 579(104) Kutzbach, C., 356, 359(20), 379 Krebs, H. A., 301, 308, 310(4), 311(4), Kyle, W.E.,143 337, 379 Kress, L. F.,289(70), 291, 293(70), 294 Kreutner, W.,68 Krietsch, W. K. G.,336(7), 337, 338, 340 (7), 341(7), 342(7), 343, 344(35), 346 (7), 347(7), 348(7), 349(7)

599

AUTHOR INDEX

Lacombe, G., 460, 461(22), 462(22, 39), 463(39), 466(22, 391, 467(22, 391, 468 (22), 474(22, 391, 475(22), 478(22), 479(22), 485(84) Ladd, J. N., 148 Lafuma, G., 542, 543 Lagerkvist, U., 9, 10(67), 13(67), 37(239), 38, 39(239), 40(239) Lahiri, S., 491, 492(42) Laland, S. G., 14 Lambion, R., 492 Lammel, C., 75, 77(12), 93, 94(12), 96 (121, 97(12), 99(12), 100(12), 110, 118(75) Landon, M. F., 393, 394(57), 434, 466, 467(57), 482 Lane, M. D., 6(47), 7, 13(47), 18(47) Lang, G., 4(9), 5 Langan, T. A., 556, 568, 574(86), 575, 579(128), 580(128) LaPlaca, S. J., 213 LaPointe, J., 9, 13(81) Lardy, H. A., 54, 232, 233(107), 240, 241, 242(5, 8), 244, 245, 246(46), 247(7), 248(7), 249, 251(70), 252(70), 253(5, 8 ) , 254(8), 255, 256(83), 259(8), 262 (57), 265(57), 269(8), 270(83), 271 (1441, 273(144), 308, 312(5), 314(5), 321(5), 324(5), 384, 392, 395(3, 48), 396, 399(73), 400(3, 73), 401(3), 403 (731, 407(73), 412(73), 416(73), 420 (731, 421(73), 422(73), 428, 431, 575 Larner, J., 52, 54, 55, 56(25), 60, 62(25), 66(25), 378, 556, 557, 574(14) Larrabee, A. R., 156, 160, 174, 175(48) Larsson-Rainikiewics, M., 336(6), 337, 339, 340(23), 341(4), 342(5, 23), 343, 344(23), 346(4, 51, 347(4, 5, 6, 46), 348(44), 349(4, 5, 6) Laster, L., 124, 126(16), 129(16) Laturase, J. G., 319, 325(39) Lauterborn, W., 373, 375(127) Lavin, B. E., 581 Lawlor, J. M., 220, 222, 225(75), 226(75) Laws, J. F., 384(12), 385, 389(43), 390 (43), 391, 392, 393(43), 400(12), 445, 469, 470(69) Layser, R. B., 240, 241(9), 242(9), 249 (91, 257, 258(9), 268(9)

Lasdunski, M., 173 Lea, M. A., 327 Leach, F.R., 6(50), 7 Le Borgne, L., 545(69), 552 Le Bras, G., 513, 514(19), 515(19), 516, 518, 519(33), 525, 526(49), 539(22), 544, 545(66), 551(661, 552 (661, 553 (19) Lech, J., 488, 489(15) Lee, E. Y . C., 94 Lee, L. M. Y., 240, 241(18), 242(181 Lee, S.C., 143 Lee Peng, C. H., 6, 12(22) Lefray, F., 573, 576(112) Lehman, I. R., 44, 46(274), 47(274, 277), 312(53a), 313 Lehninger, A. L., 309, 310(42), 313, 322 (171, 325(17), 326(17), 329(17), 330 (64), 331(17), 333(17) Lehmann, F. G., 284

Lehmann, W., 283 Lehninger, A. L., 502 Leiderman, B., 372, 373 ( 123), 376( 123), 377(123), 378(123)

Leigh, J. S., Jr., 295, 303(77a), 304, 364, 368(84), 380, 439

Leigh, R. A., 360 LeliBvre, Y., 340 Leloir, L. F., 33, 57, 58, 73, 74, 245 Lemaire, G., 9(89, 901, 10, 13(89, 90) Lemaire, S., 567, 568, 569(87), 570(87), 572(87), 573(87), 577(87)

Lenhert, P. G., 123, 144(7) Lennarz, W. J., 156, 157(7), 190(7) Lents, P. J., Jr., 454 Leonard, K. R., 255 Leonard, N. J., 245, 300 Lepp, C. A., 493 Leroux, J. P., 354 Levi, A. S., 35(222), 36 Leveille, G. A., 56 Levin, A. P., 491,492(32) Levine, S., 63,69 Levitski, A., 98, 103(54) Levy, H. L., 145 Li, T. K., 393,438 Liao, C.-L., 357, 366(33), 374(33), 376 (33)

Liberti, M., 175

600 Lichtler, E., 310(32), 312 Lieberman, M., 59, 60(93) Lifshitz, Y.,371 Light, R. J., 156, 157(7), 190(7) Lilljeqvist, A. C., 145 Lilly, M.D.,358 Lin, E. C. C.,491, 492(30, 32), 493, 494 (591, 496(59), 497(59), 498(59), 499 (59), 500(59), 501(59), 502(31, 38, 67), 503 Lindahl, T., 47 Lindell, T. J., 240, 241(21), 242(21), 247 (211, 265(21), 267 Lindsey, J., 144 Lineback, D. R.,52 Ling, G.M., 377 Ling, K.-H., 240, 241(5), 242(5), 245, 246(46), 253(5) Linn, T. C., 558,565 Lipmann, F., 2, 3(1), 6(40, 41, 42, 43, 45), 7, 9, 10(68), 11, 12(13, 14, 21), 13(13), 14(75), 15(75), 21, 27, 35 (220,229), 36, 555, 570 Liston, J., 108 Littauer, U. Z.,21, 22(140, 165, 166), 23 (140), 24, 25(165, 166, 177) Little, J. W.,45(275), 46, 47(275) Littmann, C., 355 Litwack, G., 280, 289(19, 201, 291(19, 20), 292(20), 296(19, 20), 297(19, 20), 298(19), 299(19) Live, T.R.,44, 46(272), 47(272) Llewellyn, D. R.,232, 233(92) Llorente, P.,357, 374(49), 376(49), 377 (49) Lloyd, G. J., 228 LO, C.-H., 286 Lochmuller, H.,2, 3(3), 6, 12(3), 18 Lockwood, D. H.,124 Loeb, J. E.,574,575 Loerch, J. D., 54 Loftfield, R. B.,6(33), 7, 9, 10, 13(79), 39(33) Lohmann, K., 458, 459, 461(9, lo), 462(9, 10) Lohr, G. W.,356, 359(15), 379(15) Lombardini, J. B., 125, 128, 129(33), 130 (33),135(20, 33), 136(20,33),137, 142 (20) Long, F. A,, 202

AUTHOR INDEX

Longlands, M.,61 Lorenson, M. Y.,240, 241(14), 242(14), 249, 269(14), 272(14) Lorini, M., 566, 580 Lou, M. F.,249, 263(67), 264(67) Love, D. S., 240, 241(7), 242(7), 253(7), 254(7), 255(7), 260(7), 270(7), 559, 560(34), 561(34), 562(34), 564(34) Love, E.J., 203 Lovenberg, W.,568, 570(85), 572(85) Loviny, T.,518, 519(33), 543, 549(64) Lowenstein, J. M., 248, 266(59), 297, 301, 302, 303 Lowry, 0. H.,66, 117, 240, 241(4), 242 (4), 244(4), 248(4), 249(4), 258, 262 (4, 109, ill), 264, 265(4, log), 266 (128), 267(109), 269(4), 275, 278(4, 94) Lowy, J., 464 Lubin, M., 115 Luchsinger, W. W.,9 Luft, J. H.,240, 241(7), 242(7), 253(7), 254(7), 255(7), 260(7), 270(7), 559 Luh, W., 338 Lui, N. S. T., 394, 438(58), 449(58) Lundquist, F.,494, 497(60), 498(60), 499 (60), 500(60), 503(60), 504(60) Lusk, J. G.,115 Lyn, G.,47 Lynen, F.,6(47), 7, 13(47), 18(47), 158, 159(18), 160(18), 169, 170(40), 177 (20), 178(55), 181(53, 54, 551, 182 (55), 182(55), 186(53, 541, 189(50, 53), 198,275 Lyon, J. B., Jr., 558 Lysenko, O., 108

M Maas, W. K., 6(46), 7 McAuslan, B.,145, 148(79) McBride, 0.W.,488, 489(18), 491(18) McCormack, J. H., 56 MeDaniel, E.G.,160 MacDonald, H.L., 55 MacDonald, P. W., 77, 86(25), 92(25) McDonald, T.,281 McElroy, W. D., 7, 11, 12(128, 129, 1301, 19, 20(128) Macfarlane, N.,380

AUTHOR INDEX

Mack, B., 14 MacKay, M., 144 MacKinley, A. G., 572 McLaughlin, A. C., 405(83), 406, 407, 409, 440(89), 441(89), 449(83, 841, 450(89) McLean, P., 504 McLick, J., 227 McPherson, A., Jr., 359, 454 Maddaiah, V. T., 56 Medley, T. I., 263(121), 264 Madsen, N. B., 56, 57, 338, 339(17) Maeba, P., 286, 357, 359(30), 366(30), 374(30) Maeno, H., 567, 581 Magns, S., 145 Magasanik, B., 39, 40(241), 491, 492(32) Magni, G., 43 Mahler, H. R., 6, 12(18), 24, 25(176) Mahowald, T. A., 291, 292(72), 293, 301 (751, 389(41), 390, 391(41), 392(41), 393, 395, 396(64), 401(64), 405(64), 413, 416(97), 418(64), 421(64, 97), 422(97), 431(64), 432(97), 436, 438 (51), 443(64), 445(64), 446(64), 449 (641,469, 470(68) Maier, K. P., 258 Mailhammer, R., 47, 48(283) Maitra, U., 21, 23(153, 1541, 24 Majerus, P. W., 156, 166(8), 167(8), 188 (36), 171, 172(42), 173, 176(8), 179, 183(50), 185(50), 187, 190, 191, 193 (51, 88), 194(51) Majumder, G. C., 567, 572(75) Makman, M. H., 9(105), 10, 13(105) Maland, L., 308, 315(12), 320(12), 395, 399(63a), 400(63a), 401 (63a), 403 (ma) Malay, F., 54 Malay, G. F., 54 Malcolm, L. A., 340 Malcovati, M., 357, 374(31), 379(31) Maltbie, M., 564, 574(58) Manchester, K. L., 277 Mandel, M., 108 Mandel, N., 213 Mandel, P., 281 Mangum, J. H., 43 Mann, J. D., 131, 132(36), 133(36), 140, 149(36)

601 Mansour, T. E., 240(29, 30), 241(4, 11, 12, 13, 19, 242(11, 12, 13, 141, 249, 253, 259(12, 13), 260(13, 29), 262 (13, 29,' 301, 265(13, lOa), 267(13), 269(14, 29, 98), 270, 272(14, 29, 79); 273, 278(104) Mantel, M., 42 Mantsavinos, R., 310(38a, 38c), 312 Marchall, R., 61 Marco, R., 357, 374(49), 376(49), 377 (49) Marcus, F., 240, 241(5), 242(5), 253(5), 460, 461(20), 462(2O), 465(20), 468 (201, 484(20), 486(20) Marechal, L. R., 59 Marguarding, D., 215 Markland, A. W., 312(56), 313 Markland, F. S., 280, 281, 289(17), 291 (17), 292(17), 294(17), 296(17), 297 (171, 298(18), 299(18), 301(18), 302, 303(18) Markovita, A., 59, 60(93) Marsh, W. H., 579 Marshall, R. D., 9, 13(83) Martell, A. E., 235 Martelo, 0. J., 574 Martin, B. R., 566 Martin, R. G., 119 Martin, R. J., Jr., 488, 490(21) Masaki, J., 490, 491(23) Masamume, Y., 44, 46(272), 47(272) Massey, V., 85 Masuda, H., 492 Matamala, M., 9, 10(57), 12(57) Mathias, M. M., 270, 271, 273(148) Matschinsky, F. M., 266, 275(129) Matsumura, S., 158, 159, 160(17), 166 (14), 169, 170 Matsuo, I., 281 Matthews, B. W., 185 Maul, S. B., 518, 519 Mavis, R. D., 247, 248(56) Max, P. F., Jr., 56 Maxwell, E. S., 53 Mayer, J., 488, 490(20), 493(20) Mayer, S. E., 559, 563, 564, 574(58) Mecke, D., 41,43(246) Mehler, A. H., 9(108), 10, 11, 12(77), 13(82, 108) Meier, H., 59, 60(95)

AUTHOR INDEX

Meister, A., 6(34, 35, 36, 37, 38, 391, 7, 9, 10(55), 12(24, 25, 26, 36, 39) Melander, W., 85, 361 Melchoir, J. B., 361, 368, 371 ( 105), 377 (116) Melhorn, D. K., 108 Meng, H.C.,278,574 Menge, H.,284 Menaies, R.,354 Mercer, W. D.,344, 348(42), 359 Meriwether, B. P., 177, 183(52) Metaenberg, R.L.,142 Meyer, F.,563, 574 Meyer, W. L.,559, 560(34), 561(34), 562 (341,563, 564(34) Meyerhof, O., 458 Meyers, C.,148 Michelson, A. M., 21, 22(133) Mildvan, A. S.,70, 71(146), 85, 231, 241, 244, 303, 360, 364, 367, 388032, 841, 369(106), 370(110), 372, 377(106), 378, 381, 382, 441, 444(144), 485 Miles, J. B., 384, 389(43), 390(43, 45), 391, 392(45), 393(43), 469, 470(67, 69) Miller, D. L., 204, 223041, 229 Miller, D.R.,374 Miller, E. M., 229 Miller, F., 282 Miller, L. A,, 281 Miller, L. K., 311(47), 312(47), 313, 332(47) Miller, 0.J., 340 Miller, 0. N.,384(19), 385 Miller, S.I.,213 Mills, G.T.,52 Mills, R. R.,281, 298(26), 299(26) Mills, S.E.,520,539(43) Milman, L.S.,357, 374(41) Milner-White, E. J., 396, 409(65), 410 (65,%), 412(65), 416(65), 4210351, 423(65), 424(65), 425, 427(65), 435 (140a), 437(65), 438(65), 439, 448 (65), 449(65), 453(65) Milstien, S., 202, 206(8), 221 Minamikawa, T.,58, 74 Mindich, L.,491,493(36) Mislow, K.,213, 218

Mitchell, H.L., 215 Mitra, S. K.,9,12(77) Miyajina, R.,552 Miyamoto, E.,568, 569(88), 571, 572(88), 574 Miyoshi, Y.,325 Moeski, H.,6 Moffatt, J. G.,218 Mohme-Lundholme, E.,565 Mohri, H., 490, 491(23) Moldave, K.,6, 9, 10(56), 12(24, 25, 26) Mollering, H.,492, 493(46), 494(46), 495 (461, 495(46), 499(46), 501 (46), 502 (46) Monod, J., 60, 246(48), 263(48), 265(48), 267(48), 373, 526, 530, 547(50) Moore, D.,357, 374(39), 376(39) Moore, E.,446 Moore, S.,179 Morales, M. F., 395, 397(63, 85), 407 (63), 408(85), 413(63), 416(63), 420 (981, 421(85, 98), 422(63, 98), 423 (98), 424(63, 981, 427(98), 524 Moreland, B. H.,384, 387, 389(8), 390 (8), 393(8), 395(34), 400(8), 443, 445 (S), 450(147), 461, 462(41), 463(41), 466(41), 467(41) Moret, V., 566,580(67) Morimura, H.,356, 359(16), 360(16), 373, 374(16), 376(16) Morningstar, J., Jr., 371 Morrison, J. F.,396, 403, 406, 407, 408, 409, 411, 412(93), 413(95), 414(10), 415, 416(101), 417(86, 1051, 418(101), 421, 422(95, 101, 1051, 424(111), 427 (1111, 428(105), 429(105), 430, 436 (lOl), 437( 101), 439(69), 460, 461 (19, 20, 211, 462(19, 20, 21, 371, 463 (37), 464(21), 465(20, 21, 491, 468, 468(20, 211, 471(19, 551, 472(19, 21, 55), 473(21, 72), 474(19, 721, 475(19, 72), 476(19, 72), 477(19, 54), 480(55), 482(21, 72), 483(55), 484(20, 551, 485 (21, 55), 486(20, 211, 485, 486(90) Mortimer, D. C., 58, 311(48), 313 Morton, R.K.,2,3(7) Mosley, W. H., 31(218), 33

603

AUTHOR INDEX

Mourad, N., 308, 309(13), 310(7), 313 (7), (13, (7), 328,

314(7), 315(7, 13), 316(7), 317 14), 319(7), 321(7), 322(7), 323 324(7), 325(7, 14), 326(7), 327, 330(14) Mourant, A. E., 283 Moyed, H. S., 39, 40(241) Mudd, S. H., 124, 125(14), 126(15, 16, 21), 127(14, 24), 128(14, 24), 129 (14, 16, 241, 130(14, 23, 24, 28, 291, 131(28), 132(24, 28, 361, 133(14, 361, 134(14, 24), 135(14, 211, 136(29, 411, 137(24), 138(141), 139(24, 28), 140, 142(21), 143(40), 145, 146(24), 149 (36) Muench, K. H., 9(91, 921, 10, 13(91, 92) Muesing, R. A., 189 Muetterties, E. L., 214, 215(49) Muirhead, H., 344, 348(42), 359 Muller, X., 359 Munch-Peterson, A., 33, 52, 53, 54, 62 (31), 65, 66(131), 70(15), 148 Munk, P., 519 Munro, G. F., 374 Muntz, J. A., 247, 248(55) Murnkarni, Y., 228 Murata, T., 57, 58, 74 Murphy, A. J., 524 Murphy, S., 145, 147(83) Mushak, P., 213 Mushfa, T., 357, 374(45), 375(45), 376 (45) Muto, A., 82 Myant, N. B., 488, 489(8), 490(8)

N Najjar, V. A., 232,233(97) Najman, A., 354 Nakae, T., 54, 63, 65, 66(126), 67(126) Nakarnura, H., 309, 310(30), 313(30), 315(30), 321(30), 322(30), 325, 326 (301, 329(30) Nakata, Y., 21, 23(154) Nakayana, K., 552 Narnm, D. H., 563, 564, 574(58) Nass, G., 9, 13(80) Nasu, T., 258 Nath, K., 49

Natori, Y., 143 Naurnann, K., 213 Nawa, H., 17 Naylor, R. A., 213 Negelein, E., 335 Neidhardt, F. C., 503 Nelson, 0. E., 56,58(51), 94 Nemethy, G., 267, 526 Neri, G., 357 Netter, K. F., 275 Neufeld, E. F., 52, 59, 70(4) Newell, P. C., 55, 61(38) Newsholme, E. A., 240(36), 244, 248, 262 (36, 611, 265(36), 266(36, 61), 267 (361, 275, 277(127, 164), 488, 489 (6, 17), 490(17, 191, 491(17), 493(6, 171, 494(6), 496(19), 498(6, 19), 500 (6), 501(6), 502(6), 504(6) Newton, M. G., 219 Newton, N. E., 571 Ng, W. G., 69 Nigam, V. N., 55 Nihei, T., 395, 397(63, 851, 407(63), 408 (851, 413(63), 416(63), 420(98), 421 (85, 981, 422(63, 981, 423(98), 424 (63, 98),427(98), 446 Nijessen, J. G., 354, 374(4), 379(4) Nikaido, H., 54, 63, 66(126), 67(126) Nikiforuk, G., 301 Niles, E. G., 45 Ning, J., 252 Nishikawa, M., 57, 257, 258 Nishizuka, Y., 47, 49(205), 570 Nisrnan, B., 510(10), 511 Nixon, J. E., 178 Noda, L., 232, 233(107), 280, 289(70), 291, 292, 293(70), 294, 295(73), 296(12), 297(12), 298, 299(81), 301(4), 384, 392, 395(3, 481, 396, 397(63, 851, 399 (731, 400(3, 731, 401(3), 4 0 3 0 3 , 407 (63, 731, 408(85), 412(73), 413(63) 416(63, 731, 420(73, 981, 421(73, 85, 98), 422(63, 73, 98), 423(98), 424 (63, 98), 427(98), 428, 446 Nolan, C., 558 Noll, F., 355 Noltmann, E. A., 240(34), 243, 291, 292 (72), 293, 301(75), 389(41), 390, 391 (41), 392(41), 395, 396(64), 401(644),

604

AUTHOR INDEX

405(64), (64, 97), 431(64), 446(64), (45)

413, 416(97), 418(64), 421 422(97), 428, 429, 430(113), 432(97), 443(64), 445(64), 449(64), 463, 465(45), 466

Norman, A. W., 309, 312(19), 319(19),

Omston, L. N., 505, 507(79), 508(79) Ornston, M. K., 505, 507(79), 508(79) Ortir, P. J., 21, 22(162), 23, 24(162), 25 (162), 26

Osborn, M., 119 Oshima, T., 240(32), 243, 263(32), 264 (32), 266(32), 267(32)

325(19)

Norris, A. T., 9, 10(58), 13(58) Novelli, G. D., 6(30, 31, 461, 7, 11(30), 31(208), 32.

Novoa, W. B., 558 Nowak, T., 364, 367, 369, 381, 382

0 Oakenfull, D. G., 227 Ochoa, S., 4(8), 5, 21, 22(137), 23, 232, 233(100), 355

Oda, A., 324 Oesper, P., 341,347(34) Oesterhelt, D., 177, 181(54), 186(54) Ogilvie, J. W., 518, 519 Ogiwara, H., 552 Ogston, A. G., 396,439(69) Ohga, Y., 580 Ohrmann, E., 357, 374(37) Okabe, K., 384(17), 385, 394, 395(61), 399(63a), 400(63a), 401(17, 63a), 402 (17), 403(63a), 447, 448(150) Okamoto, H., 27 Okamoto, T., 9, 12(52), 13(52) Okanishi, M., 29 Okuno, G., 57,257 Old, L. O., 286 Oldham, K. G., 232,233(92) Oliver, I. T., 55, 65, 66(129), 67(129), 301, 396 Olivera, B. M., 44, 46(274), 47(2’74) Olson, 0. E., 293, 389(42), 390(42), 391, 392(42), 393(42), 394 (421, 395(52), 399(42), 403(52) Olsson, P., 232, 233(106), 308, 309(6), 312(6), 313(6), 314(6), 315(6), 319 (6) Opie, L. H., 266 Opits, J. M., 340 Ord, M. G., 579 Ordin, L., 56, 57(57) Oriol, C., 434, 466, 467(57), 482 Oriol-Audit, 393, 394(57)

Oshinsky, C. K., 45(275), 46, 47(275) Osterbaan, R. A., 185 Osterman, J., 374, 379(133) O’Sullivan, W. J., 298, 299(81), 346, 396, 409, 412, 413(95, 961, 422(95), 429 (96), 430(96, 116), 437, 438(116), 439 (69, 137), 440(137), 464, 466(48), 472(48), 476(48), 479(48), 480(48), 483(48), 484 Otsuka, K., 492 Otsuka, S., 552 Ottaway, J. H., 385 Ouellet, L., 232, 233(101) Ozaki, H., 118, 119(89), 357, 366(36), 374 (361, 375 (36) Orawa, E., 562, 563(48) Ozbun, J. L., 94

P Packmann, U., 9(85), 10, 13(85) Paetkau, V., 240, 241(8), 242(8), 247(8), 248(8), 253(8), 254(8), 255, 2560331, 259(8), 262(8), 270(83), 271(144), 273 (144) Paglia, D. E., 340, 374 Paladini, A. C., 245 Palm, P., 48 Palrnieri, R. H., 308, 315(12), 320(12), 389(42), 390(42), 391, 392(42), 393 (42), 394(42), 399(42) Pan, F., 128, 135(32), 138(32), 143 Panar, M., 219 Pande, S. V., 491, 495(41), 496(41), 501 (41) Pannbacker, R. G., 61 Pant, R., 461, 471(33), 475(33) Papas, T. S., 9(108), 10, 13(108) Pardee, A. B., 35(226), 36, 240 Parks, C. C., 309, 311(25), 314(25), 315 (25), 316(25) Parks, L. W., 141, 143(50) Parks, R. E., Jr., 247, 262(57), 265(57), 308, 309(8), 310(7, 8, 22, 24, 271, 311

AUTHOR INDEX

(25, 331, 312(56), 313(7, 8, 22, 271, 314(7, 8, 22, 24, 25), 315(7, 8, 22, 24, 25, 27), 316(8, 22, 25), 317(22), 319 (7), 321(7, 22), 322(7, 8, 22, 24), 323 (7, 8 ) , 324(7, 81, 325(7, 8, 241, 326 (7, 8, 241, 327(7), 328, 329(8, 24), 330(8), 331(8, 22), 332(8, 24, 331, 333 (22) Parmeggiani, A., 240, 241(6, 71, 242(6, 7), 243, 253(7), 254(7), 255(7), 260, 262(110), 264, 270(7), 277, 559 Partridge, M., 75,94(10) Parvin, R., 491, 495(41), 496(41), 501 (41) Passeron, S., 357, 370, 374, 376, 379(38, 135) Passoneau, J. V.,240, 241(4), 242(4), 244 (4), 248(4), 249(4), 258, 262(4, 109, lll), 264, 265(4, lW),266(128), 267 (1091, 269(4), 275, 278(4) Pastan, I., 492 Pasternak, C. A.,35(224), 36, 37(235), 38 Patte, J, C.,514, 515, 516, 517, 518, 519 (331, 539(22), 542, 543, 544, 549(64) Patterson, B. D.,281 Paule, M. R.,75, 83(20), 84, 85, 86 Paulus, H.,259, 269(102), 491, 494(28), 497(28), 547, 548(73), 549(72), 550 (74) Pavlinova, 0. A.,59 Pawelkiewicz, J., 145 Payne, K.J., 24,25(169) Pazur, J. H., 52 Peaud-Lenoil, C.,52 Peck, E.J., 232,233(94) Pedersen, P. L.. 309, 310(36, 39, 401, 312, 315, 316(57), 319(67), 322(17), 325 (17), 326(17), 329(17), 330(57, 641, 331(17, 57). 33307. 39, 40) Pegg, A. E., 124 Peller, L.,422 Pelletier, G.,567 Penefsky, H.S., 232, 233(102, 103) Pennington, R. J., 311(46), 313 Penny, I. F.,342 Pentchev, P. G.,341,344(35) Perkins, J. P., 556, 559, 561, 563(42), 566 (12, 411, 568(12), 572(12), 573(12), 574(12, 41) Perlman, R.L.,492

605 Perlaweig, W. A.,123 Perriard, J. C.,388 Perrin, D. D., 346, 412, 413(96), 429(96), 430(96, 115), 484 Perry, S. V., 287,431 Peterkofsky, A., 135, 144, 145(74), 146 (74, 841, 148(74), 149(74, 86) Petricciani, J. C., 53, 62(21), 63(21), 65 (211, 66(21), 67(21), 69(21), 70(21) Pette, D.,241, 244, 245(39), 259, 268, 282, 338 Pettit, F.H.,556, 565(11) Petzold, G.L.,571 Pfleiderer, G.,301,371 Pfohl, S.,215 Phillips, G.T.,178 Phillips, T. M.,366, 381(97) Phillips, R.C.,2, 3(6) Piccinini, F., 177, 178(55), 181(55), 182 (55) Pickworth, J., 144 Pigg, C.J., 141, 143(50) Pihl, A.,271, 273(151) Pinna, L. A.,566,580(67) Pitra, C.,56 Plackett, P., 492 Plate, C. A., 178, 186(62, 64) Plaut, G.W.E.,280,325 Plowman, K.M.,364 Pogson, C. I., 240(35), 244, 262(35), 266 (35), 271, 357, 363, 374(82), 375(48), 377 Poiret, M., 551, 552 Polli, E.,311(44), 313, 325(44) Pon, N. G.,355,361,371 Poole, W.E.,339 Poon, W.M.,258 Porter, H.K.,56 Porter, J. W.,160, 178, 186(56), 189 Posner, J. B.,574 Posternak, S.,555 Potter, J., 558 Potter, V. R.,131, 302, 310(38, Bb), 312 Powell, C.A.,563,573 Powell, G.,161, 175(27), 191, 193(88) Poyer, J. L.,31(218), 33 Pradel, L. A., 431, 432, 433, 434(128), 443(129), 453(128), 454(127, 1281, 455(128, 129), 460, 461(14, 15, 17, 29, 30), 462(14, 15, 17, 29, 30, 31, 35,

606

AUTHOR INDEX

Racker, E., 232, 233(103), 245, 246(47), 247(47), 262(47), 265, 266(47), 493 Raijman, L., 232, 233(98) Rall, T.W.,26, 556 Ramaiah, A.,247, 262(114), 263(53), 264, 268, 276 Ramakrishaan, C. V., 6, 12(18) Ramakrishnan, T.,492 Ramaley, R., 308, 309(9), 312(9, 51), 313, 315(9), 316(9), 319(9, 51), 320(9), 322(9), 324(51), 325(9), 326(9), 327 (91,329(9), 331(9) Ramirea, F., 213, 214, 215 Ramsay, B.,201(3), 202 Randerath, K.,301 Randle, P. J., 240(35), 244, 262(35, 1121, 264, 265, 266(35), 275, 277(127), 278, 490, 566 Ranney, H.M.,257 Rao, D.R.,341, 347(34) Rao, K.M.K.,59 Raper, J. H.,466,467(59) Rapin, A., 59 Rapley, S.,283 Rask, L.,232, 233(106), 308, 309(6), 312 (61, 313(6), 314(6), 315(6), 319(6) Rasmussen, H., 574 Ratliff, R. L.,232, 233(106), 308, 312(5), 314(5), 316(11), 321(5), 324 Ratner, S., 2, 3(2), 37(236, 237), 38, 44 Ray, P. M.,60 Ray, W.J., 232,233(94) Recondo, E.,33, 73, 74 Reddi, A. H.,569, 570(92), 572(92), 577 (92) Redfield, B., 144, 145, 146(84) Reed, G. H., 71, 295, 303(77a), 304, 362, 411, 412(94), 426(94), 437(94), 438 (941, 440, 441(94), 442(91), 449(94), 453(94) Reed, J., 255 Q Reed, L. J., 6(48, 49, 501, 7, 13(48, 49, 50), 17(48, 49), 18(48, 49), 286, 556, Qosba, P., 48 565 R Reed, W.D., 565 Rabin, B. R., 294, 431,432,438, 439(139), Reeves, R. E.,354 442( 139), 443( 139), 444( 139), 445 Regnouf, F.,460, 461(14), 462(14), 468 (14), 470, 478(14), 479(14), 482(14) (122), 446(120), 448(139), 449039) Reid, E., 56 Rabinowitz, M.,555, 557, 559(17) Reiff, D.P.,213 Rabussay, D., 47, 48(282, 283)

38), 464(31), 465, 466(17, 311, 467 (17,31, 581, 468(14, 30, 311, 469, 470 (65, 661, 471(31, 35, 47), 472(31, 35, 47, 7l), 473(35), 474(35, 471, 475 (35, 38, 71), 476(35, 47, 71), 477 (35), 478(14, 17, 29, 76), 479(14), 480(71, 76, 78), 481, 482(14, 35, 52, 71, 75, 78, 82, 83), 483(52, 78, 82), 484(52), 485(82) Pradet, A,, 281, 285 Prager, M. D.,356 Prakash, L.,551, 552 Prasblova, M. F.,59 Preddie, E. C.,9(88), 10, 13(88) Preiss, J., 9(107), 10, 13(107), 22(164), 24, 25(164), 31(202, 204, 210, 212), 32, 33, 75, 77(12), 78(15, 171, 81(18, 191, 83(17), 86(21, 22, 23, 24, 261, 87 (21, 22, 23), SS(22, 23), 89(23, 241, 90(24, 26), 93(23), 94(10, 11, 13, 47), 96(12), 97(11, 12, 13, 521, 99(11, 12, 13, 52), loO(12, 521, 101(57), laZ(l1, 57), 103(57), llO(11, 27, 52), 113(52, 56, 57), 114(11, 56, 57), 115(78), 117 (781,118(19, 73, 751, 119(89) Prescott, D. J., 166, 171, 173(33), 189, 190(82) Prescott, N.,29 Pressey, R., 56, 58(50) Pressman, B. C.,117 Price, N. C.,293, 368, 373, 378(124) Pricer, W. W.,Jr., 5, 31(12), 3202) Pringle, J. R., 241 (23), 242(23) Pritchard, J. G.,202 Prosen, R. J., 144 Pugh, E.L., 156,167, 176(9) Puig, J., 110 Pullman, M. E.,232, 233(103), 285 Purich, D.L.,252 Putman, E.,52, 70(4)

AUTHOR INDEX

Reim, M., 337 Reimann, E. M.,559, 561, 563(42), 568, 569(90), 570(89, 90), 571(89, 90),572 (89, 901, 573(35, 901, 574(35), 576 (89,901, 578(89) Reio, L.,82, 510 Remy, P.,365 Rendi, R., 310(35), 312, 318(35), 319 (351, 333(35) Resnik, H., 85 Reuben, J., 367, 368, 370(109), 381(109) Reynard, A. M.,250, 370, 376(114) Rhoads, D.G.,301, 302, 303 Rhodes, W.C.,12(128), 19, 20(128) Ribereau-Gayon, G., 75, 77(12), !M(12), 96(12), 97(12), 99(12), 100(12), 118 Rich, A.,359 Richards, H.H., 126 Richardson, C. C., 21, 44, 46(272, 2731, 47(272) Richardson, D. I., Jr., 227, 236 Richardson, J. P.,21,22(134) Richey, D.P.,491,502(31,38) Richman, D.J., 556, 574(14) Richterich, R., 384, 386 Richters, A. R., 287 Rickes, E. L., 144 Riepertinger, C.,18 Rigler, R., 9(85), 10, 13(85) Riley, G. A., 565 Riley, W.D.,558, 559(28), 560(29), 561 (391,563(28) Rinaudo, M.T.,55 Ritter, E., 158, 159(18), 160(18), 177, 178(55), 181(55), 182(55) Robbins, P. W., 35(228, 2291, 36, 54, 59, 62(98), 88(98), 68(98) Robbins, R. K., 573 Robert-Gero, M.,545(69), 551, 552 Roberts, R. B.,510 Roberts, R. M.,56, 59, 66(52) Robertson, J. H.,144 Robin, Y., 393, 394(57), 459, 460, 461 (231, 462(23, 35, 36, 38, 40), 463, 466(23), 467(23, 611, 468(23), 470 (40),471(35, 36, 401,472(35, 36), 473 (3.9, 474(23), 475(36, 381, 476(23, 36), 477(36), 482(23), 484(23) Robinson, J., 488, 489(6), 490(19), 491,

607 493(6), 494(6), 496(19), 498(6, 19), 500(6), 50103, 502(6), 504(6) Robinson, J. L.,183, 364, 380, 381(87) Robinson, L.,231 Robison, B., 309, 310(22), 313(22), 314 (22), 315(22), 316(22), 317(22), 319, 321(22), 322(22), 333(22) Robison, G. A.,278, 585 Robson, E.,283 Roche, J., 459, 461, 462(39), 463(39), 466(39), 467(39), 474(39) Rochovansky, O., 37(236, 2371, 38, 44 Rodbell, M.,26 Rodnight, R., 581 Rodwell, A. W.,492 Rohlfing, S. R., 108 Rominger, K. L.,6(47), 7, 13(47), 18 (47) Roncari, D. A. K., 169, 170 Rosalki, S., 386 Roschlau, P.,356, 358(13), 359(52a), 360 Rose, I. A., 4(8), 5, 241, 244, 252, 286, 364, 365, 380(86), 381(87) Roselino, E.,357, 374, 379(38, 135) Rosen, 0.M.,568, 570, 571(83), 572(83, 98), 577 Rosenberg, H., 410, 431, 461, 462(32), 464(32), 471(32, 34), 472(32, 34), 475 (32, 341, 476(32, 34), 477(32) Rosenberg, L. E., 145 Rosenbloom, F. M.,145 Rosenbusch, J. P.,519 Rosenthal, S. M.,124, 125(13, 171, 126 (13), 130(17) Rosett, T.,281 Roskoski, R., Jr., 6(40, 43), 7, ll(40, 43) Rosner, A., 548, 549, 550(74) Ross, P.D.,139 Rossiter, R. J., 461, 471(34), 472(34), 475(34), 476(34) Rossman, M.G., 454 Roth, J. R., 522, 538(42) Roth, R., 59,60(94), 61 Rouge, M.,9(87), 10, 13(87) Rouget, P.,9, 10(61), 13(61) Roustan, C.,433, 454(127), 479, 480(78), 481, 482(78, 821, 483(78, 82), 485(82) Rowland, L. P.,240, 241(9), 242(9), 249 (9), 257, 258(9), 262(9)

AUTHOR INDEX

608 Rowley, G. L., 403, 404(81), 405(81), 406 (81)

Rownd, R., 28, 29(194), 30(194) Roy, B. P., 384(12), 385, 400(12) Rozengurt, E., 356, 357, 372, 373(123), 374(50), 375, 376(123, 143), 377(123), 378(122, 123, 143), 379(50) Rubin, C. S., 568, 571(83), 572(83) Ruch, F. E., Jr., 178, 179(65), 180(65), 182(65), 183(65), 184(65) Ruddon, R. W., 580 Rudigier, J. F. M., 67 Rudolph, F. B., 249, 251(70), 252(70) Ruhn, C. S., 577 Ruppert, D., 261 . Rush, D., 491, 492(32) Russell, I. J., 52 Russell, P. J., 281, 282(28), 283, 289 (281, 291(28), 296(28), 297(28), 298 (29, 30), 299(29, 301, 300(29), 301 (30), 302(30), 303 Rutman, R. J., 2, 3(6) Rutter, W. J., 356, 379(19) Ruwart, M. J., 363,375(81)

5 Saari, J. C., 517, 518(29), 519(29), 520 (29), 521(29), 522(29), 523(29), 533 (291, 536 (29) Sabraw, A., 75, 77(12), 93, 94(12), 96 (12), 97(12), 99(12), 100(12), 110, 115(78), 117(78), 118(75) Sacher, J. A., 58 Sacktor, B., 493, 565 Sadowski, P., 45(276), 46, 47(276) Saenger, W., 213 St.. Liebe, G., 254 Sakamoto, Y., 14 Sakov, N. E., 269 Salamini, F., 56, 58(51) Salas, J., 274 Salas, M., 261, 263(123), 264, 266(123), 275(123) Salas, M. L., 240, 241(20), 242(20), 245 (20), 246(20), 247, 249(20, 521, 261, 263(62, 123), 264, 266(123), 274, 275 (1231, 570

Sala-Trepat, J. M., 552 Sallach, H. J., 351(21), 356, 374, 376(2) Saluste, E., 82, 510 Samamoto, J., 228 Sampath-Kumar, K. S. V., 555 Sampson, E. J., 202, 203(9), 204(9), 206 (9), 210(9), 225, 226(78), 229, 232 (78), 233(78) Samuel, D., 232, 233(104) Samuels, A. J., 393, 436 Sanadi, D. R., 309, 310(29, 34), 312, 324 Sanchez-Medina, F., 110 Sanders, M. M., 575 Sanes, J. R., 572, 573(105) Sanger, F., 555 Sanner, T., 271, 273(151) Sanno, Y., 492 Santarius, K. A., 91, 92(44, 45) Santi, D. V., 9(104), 10, 13(104) Sanwal, B. D., 286, 357, 359(30), 366(30), 374(30), 377 Sanwal, G. G., 77, 86(24, 261, 89(24), 90(24, 26) Sapico, V., 245, 25O(43), 280, 289(19, 20), 291(19, 201, 292(19, 201, 296(19, 201, 297(19, 20), 298(19), 299(19) Sargent, D., 61 Sarma, P. S., 555 Sarner, J., 491, 501(34) Sato, K., 286 Sato, T. R., 301 Sauer, F., 156, 167, 176(9) Sauer, H., 365 Sawas, C., 517 Sawicka, T., 54, 58, 66(86), 69(86) Sawula, R. V., 269 Schachner, M., 48 Schapira, F., 387 Scher, B. M., 74 Schevitz, R. W., 454 Schiller, K., 356, 379 Schirmer, H., 291, 292 Schirmer, I., 289(71), 291(71), 293(71), 295(71), 296(71), 297(71) Schirmer, R. H., 281, 284, 289(36), 291 (361, 292(36), 293(71), 295(71), 296 (36, 7 0 , 297(36, 71) Schlender, K. K., 574

AUTHOR INDEX

Schloen, C., 374 Schmidt, G., 21 Schmir, G. L., 422 Schmutzlor, R., 214 Schnaitman, C. A., 310(39, 40, 41, 431, 312, 313, 333(39, 40, 41, 43) Schneider, K. W., 284 Scholar, E. M., 309, 310(27), 312(56), 313(27), 315(27) Scholl, A., 387, 388 Schoner, W., 377 Schray, K. J., 221, 223(74), 225(74), 226 (74), 227(74), 232, 233(99), 241, 244, 381 Schrecker, A. W., 31(205), 32 Schuegraf, A., 2, 3(2), 275 Schultz, D. W., 275 Srhulz, G. E., 289(71), 291, 293(71), 295 (711, 296(71), 297(71) Schulz, H., 172 Schulze-Wethman, F. H., 566, 574(70) Schunn, R. A., 214, 215(49) Schwark, W. S., 377 Schwartz, E. R., 286 Schwarz, V., 574 Schweizer, E., 177, 178(55), 181(54, 55), 182(55), 186(54) Scopes, R. K., 337, 338, 339(13), 340(11, 13, 21), 341(30), 342(11, 131, 344 (30, 36), 345(30), 351(30), 384(11), 385, 388, 399 Scrutton, M. C., 353, 378(2) Sebald, M., 108 Secrist, J. A., 111, 300 Sedmak, J., 308, 309(9), 312(9, 511, 313, 315(9), 316(9), 319(9, 511, 320(9), 322(9), 324(51), 325(9), 326(9), 327 (91, 329(9), 331(9) Seeds, A. E., Jr., 508 Segal, A., 43 Segal, W., 491, 492(42) Segel, I. H., 36 Seifert, L. L., 52, 53, 55(16), 62(16), 63 (16), 65(16), 66(16), 67(16), 68(16) Seifert, W., 9, 13(80), 47, 48(282) Seitz, H. F., 488, 489(7), 493(7) Seliger, H. H., 19, 20(131) Sellinger, 0. Z., 384(19), 385

609 Sepulveda, L., 231 Setlow, B., 253, 269, 272(79), 273 Setondji, J., 365 Seubert, W., 377 Shanoff, M., 54 Shapiro, B. M., 41, 42(248), 43(255), 44 Shapiro, J. A., 60 Sharma, V. S., 429, 430(115) Sharman, G. B., 339 Shaw, W., 30 Shemanova, G. F., 491, 501(37) Shen, L. C., 31(210, 212), 32, 33, 94(10, ll), 97(11), 99(11), 102(11), 105, 106, 110(11), l14(11, 581, 286 Sherwin, A. L., 386, 394 Shield, B., 142 Shiio, I ., 357, 366 (36), 374 (36), 375 (361, 552 Shikamura, M., 59 Shimazu, T., 60 Shive, W., 518, 519 Shoemaker, C. B., 144 Shuey, E. W., 52 Siber, G. R., 386 Sibitani, A., 21 Siecrist, J. A., 245 Siekevitz, P., 302 Siess, E., 566, 574(70) Siewers, I., 225, 226(78), 232(78), 233(78) Sigal, N., 110 Sightler, J. H., 518 Sikes, S., 108 Siliprandi, N., 566, 580(67) Silver, B. C., 232, 233(104) Silver, S., 115 Simms, E. S., 312(53a), 313 Simon, L., 23 Simon, L. N., 573 Simon, W., 61 Simonarson, B. A., 384, 387, 388, 389(33), 395(33), 400(33), 420(33), 445(33), 448(33) Simoni, R. D., 158, 159, 172(13) Simplico, J., 362 Singer, M. F., 23 Singh, B., 56 Singh, S., 283 Singhal, R. L., 377

610 Singleton, R., Jr., 85, 372 Slatterly, P., 9(104), 10, 13(104) Slee, R. G., 374 Smellie, R. M. S., 22(175), 24, 25(174, 175) Smiley, I. E., 454 Smith, A. J., 118 Smith, C. P., 213 Smith, D., 281 Smith, D. H., 29 Smith, E., 460, 461(21), 462(21), 464(21), 465(21), 468(21), 472(21), 473(21, 72), 474(72), 475(72), 476(72), 482 (21, 721, 485(21), 486(21) Smith, E. C., 267, 374, 376(139) Smith, E. E. B., 33,52 Smith, E. L., 144 Smith, R. C., 135, 138(44) Smith, S., 160, 283 Smith, T. A., 148 Smyth, R. D., 144, 148 Soderling, T. R., 561, 563(42) Soling, H. D., 365, 566 Soll, D., 9, 13(81) Solomon, F., 252 Sols, A., 240, 241(20), 242(20), 245(20), 246(20), 247(20), 249(52), 261, 263 (52, 123), 264, 266(123), 274, 275 (1231, 278, 376, 492, 497(47) Somero, G., 357, 363, 374(47) Somerville, R., 312(54), 313 Sonico, F., 9, lO(66) Sorger, G. J., 366 Sorsoli, W. A., 141, 143(50) Sparks, J. R., 399 Spears, C., 145, 148(79) Spector, L. B., 342, 350(39), 483 Spiro, T. G., 230 Sportorno, G., 9(90), 10, 13(90) Sprinson, D., 365 Sprouse, H. M., 240, 241(12), 242(12), 259( 12) Spudich, J. A., 281 Staal, G. E. J., 354, 356, 374(4, 261, 375 (261, 376(26), 379(4) Stadtman, E. R., 4(10), 5, 40, 41, 42 (248), 43(242, 2561, 44(258), 240, 513, 514, 515(19), 533(19) Stadtman, T. C., 123, 124(4) Stafford, E’. E., 202

AUTHOR INDEX

Stahley, D. P., 551 Stam, M., 231 Stamm, N. B., 377, 379 Stammers, D. K., 344, 348(42), 359 Stanier, R. Y., 82, 108, 544, 545(66), 551 (661, 552(66) Starnes, W. L., 519 Steelman, V. S., 53, 59(20), 62(20), 66 (20) Steffens, J., 225, 226(78), 232(78), 233 (78) Steinberg, D., 574 Steinmetz, M. A., 358 Stekol, J. A., 135, 136(13) Stellwagen, E., 241(21), 242(21, 23), 247 (21), 248(56), 265(21), 267 Stephens, D. T., 248, 262(62), 266(62) Stephenson, M. L., 24 Stern, R., 9, 13(82), 574 Stevens, A., 21,22(155) Stewart, G. R., 374(39), 376(39) Stjernholm, R., 82, 510 Stocken, L. A., 579 Stocking, C. R., 56 Stone, D. B., 260, 265(104), 278(104) Stone, N., 9, lO(66) Storm, E., 376 Strand, M., 568 Stratman, F. W., 575 Stripe, F., 379 Strobel, G. A., 77, 86(25), 92(25) Strominger, J. L., 53 Stubbe, J., 365 Stulberg, M. P., 6(30), 7, 9(102), 10, 11 (30), 13(102) Stull, J. T., 558, 564(30) Stumpf, P. F., 491 Stumpf, P. K., 158, 159(13), 169, 170, 172 (13) Su, C. H., 141, 142(56) Su, S., 281, 282(28), 289(28, 291, 291(28), 296(28), 297(28), 298(30), 299(29, 30), 300, 301(30), 302(30), 303 Subbarow, Y., 458 Suda, M., 257 Sue, F., 356, 359(16), 360(16), 373, 374 (161, 376(16) Suelter, C. H., 85, 269, 356, 358(14), 359, 361, 362(71), 363(71), 366, 367, 369,

AUTHOR INDEX

371(70), 372, 373(70), 375(80, 81, 129), 378(129) Sugden, P.H.,266 Sugino, Y., 309, 310(30), 313(30), 315 (301, 321(30), 322(30), 325, 326(30), 329(30) Sugiyama, T., 57, 58 Sullivan, R. J., 384(19), 385 Sumi, T., 240(28), 241, 242(28) Sundararajan, T.A., 59, 355 Surganaryana-Murthy, P.,492 Susor, W.A., 356, 379(19) Sussman, M., 54, 55, 59, 60(94), 61(38), 62(22), 63(22), 65(22), 66(22), 70 (22) Sutherland, E.W.,26, 278, 556, 558, 565, 574 Sutherland, I., 144 Suyter, M., 488, 489(10), 490(10), 493 (lo), 495(10), 496(10), 497(10), 499 (lo), 501(10), 502(10) Suzuki, C., 27 Suzuki, I., 269 Suzuki, T., 14 Swanson, P. D.,385 Swiatek, K.R.,573 Swiatkowska, B.,60 Swick, R. W., 18 Szafraniec, L.J.,213 Szepesi, B.,379 Szorhnyi, E.T., 459, 460, 461(11, 12), 462 (11, 12), 466(12), 467(12) Szymczek, T., 60

611

331, 131(30), 132, 133, 134(30), 135 (20,30, 331, 136(20, 30, 33), 137(33), 138(30), 139(30), 142(20), 508 Talamo, B., 176, 183(50), 185(50) Talman, E.L.,67 Tanaka, H., 552 Tanaka, T., 257, 356, 359, 360, 373, 374 (161,376(16) Tanaka, Y., 57 Tanford, C.,119, 166, 168(32), 536 Tang, M.S.,143 Taniuchi, K.,356 Tanzer, M. L.,396, 405(66) Tao, M.,569, 570, 571, 572(91), 576(91), 577(91) Taqui Khan, M. M., 235 Tarnowski, W.,488, 489(7), 493(7) Tarui, S.,240, 241(19), 242(19), 252(19), 255(19), 257(191, 258(19), 270(19) Tarver, H.,128, 135(32), 138(32), 143 Tatum, E.L., 14 Taylor, B.,281 Taylor, C. B.,258, 373, 379 Taylor, J. S., 409, 440(89), 441(89, 143), 450(89) Taylor, K., 488, 489(17), 490(17), 491 (171,493(17) Teas, H.J., 509 Tejwani, G. A.,262(114), 264, 268, 276 Temkine, H., 354 Tena, G.M.,218 Tener, G. M.,9(96), 10, 13(96) Terenzi, H., 370, 374, 379(135) Terranova, T.,357 Terzi, M.,24,25(173) T Thain, E.M., 167 Tabor, C. W., 124, 125(13, 171, 126(13), Thang, M. N.,574 Theil, S.,488,489(4) 130(17, 18, 34), 136(34) Tabor, H.,124, 125(13, 171, 126(13), 129, Therriault, D.,371 Thiem, N. V., 459, 460, 461(22), 462(22, 130(17, 18, 341, 136(34) 39), 463(39), 466(22, 391, 467(22, 39), Takagi, T., 166, 168(32) 474(22, 39), 475(22), 478(22), 479 Takagi, Y., 310(38), 312 Takahashi, M., 518, 519(30), 523 (221,485(54) Thompson, A., 47 Takai, K.,27 Thompson, F.M.,332 Takasawa, S.,29 Thomson, A. R.,384(12), 385, 389(43), Takeda, F.,288 390(43, 53), 391(53), 392(45), 393, Takeda, M.,580 394, 400(12), 445, 469, 470‘37, 69) Takenaka, F., 377 Thornson. J. F.. 301 Takevama. S..577 Talalay, P., 125, 128, 129(33), 130(30, Thorner, ’J. W., 491, 494(28), 495, 496 I

I

I

612 (62), 497(28, 62), 498(62), 499(62), 500, 501(62), 502(62), 503(62) Thuma, E., 281, 284, 289(36, 711, 291(36), 292(36), 293(71), 295(71), 296(36, 71), 297(36, 71) Tietz, A., 232, 233(100), 355 Tills, D., 283 Timasheff, S. N., 255 Tincher, A., 551 Ting, W. K., 67 Tipper, D., 29 Tipton, K. F., 249, 252(65), 269 Todd, A., 144 Todd, J. K., 281 Tokushige, M., 259, 269(101) Tomino, S., 14 Toohey, J. I., 144, 148 Toomey, R. E., 193 Torres, H. N., 49 Totten, E. L., 240(37), 244 Tovey, K. C., 56, 66(52) Traugh, J., 574 Traut, R. R., 574 Travis, J., 7, 11(51), 20(51) Trayser, K. A,, 559, 560(34), 561(34), 562 (34), 564(34) Treble, D., 488, 489(11), 490(20), 493(11, 20), 498(11) Trivedi, B., 265 Trucco, R. E., 54, 245 Trueblood, K. N., 144 Truffa-Bachi, P., 512, 513(17), 515, 516, 517(25), 518(25, 29), 519(29, 34), 520 (29), 521(25, 29, 361, 522(29), 523 (28, 291, 524(28, 361, 525(25), 526 (25, 49), 527(28), 533(29), 535(36), 536(29), 537, 543 Trundle, D., 446 Tsai, C.-Y., 56, 58(51), 94 Tsai, M. Y., 258 Tsao, M. U., 263(121), 264 Tsay, S. S., 491, 492(33), 496 Tsolis, S. A., 215 Tsuboi, K. K., 53, 59, 62(21), 63(21), 65 (21), 66(21), 67(21), 69(21), 70(21) Tsuiki, S., 82, 286 Tsutsumi, E., 377 Tuominen, F. W., 356, 370, 374(113), 375 (113), 376(113), 379(29)

AUTHOR INDEX

Turkington, R. W., 567, 572(75) Turner, D. H., 62,66(123) Turner, J. F., 56, 58(48, 491, 62, 66(123), 262(116, 117, 118, 1191, 264, 266(116, 117, 1181, 311(49), 313, 315(49), 320 (491, 321(49), 329(49), 330(49) Tweedle, J. W., 36 Tweto, J., 175

U Uchino, H., 145 Uedi, A., 27 Ugi, I., 215 Uhr, M. L., 411, 412(93), 460, 461(20), 462(20), 465(20), 473, 484(20), 486 (20) Ui, M., 240(28), 241, 242(28), 265 Ukena, T., 204 Umbarger, H. E., 511(14), 512 Umbreit, W. W., 492 Umezawa, H., 29 Underhill, J. A., 400, 408, 409(87), 420 Underwood, A. H., 240(36), 244, 248, 259, 262(36, 611, 265(36), 266(36, 61), 267 (36), 275, 277(164) Ursprung, H., 388 Usher, D. A., 219,227,236 Utahara, R., 29 Utter, M. F., 311(46), 313, 353, 378(2) Uyeda, K., 241(17, 19), 242(17, 19, 271, 243(17), 244, 245, 246(47), 247(47), 249, 250, 252(19, 64), 253(27), 255 (19), 256(27), 257(19), 258(19), 260 (64),262(47), 263(27), 265, 266(47), 267(27), 270(19), 273

V Vagelos, P. R., 156, 158, 159(15), 160, 161, 162(28), 165, 166, 167, 168(36), 169, 170(39), 171(33), 173(33), 174 (46), 175(27, 481, 176(8), 179, 180, 183(50), 185(8, 501, 187, 188(1), 189, 190(81, 821, 193(51), 19461, 81), 195 (41, 81), 196(41), 197(41), 198(41) Valadares, J. R. E., 563 Valentine, W. N., 374

613

AUTHOR INDEX

VanAdrichem, M., 185 Vanaman, T. C., 158, 159, 166(16), 168 (161, 169, 170 Vnnbellinghen, P., 356 VanBerkel, T. J. C., 374 Van de Berg, J. L., 339 Vandemark, P. J., 491, 492(39) VandenBergh, S. G., 333 van den Bosch, H., 161, 162(28) Van der Voorn, P. C., 213, 215(44) Vanhembeeck, J., 9, lO(60) Van Milligen-Boersma, L., 356, 374(26), 375(26), 376(26), 379 Van Noort, D., 281 van Rapenbusch, R., 9(89), 10, 13(89), 516, 517(25), 518(25), 519(34), 520 (25), 521(25), 525(25), 526(25), 540, 542(60) Van Thoai, N., 391, 393, 394(57), 431, 432, 433, 434(128), 453(128), 454(128), 455(128), 459, 460, 461(14, 15, 17, 22, 23, 28, 29, 30), 462( 14, 15, 17, 22, 23, 28, 29, 30, 31, 35, 36, 38, 39, 401, 463 (391, 464(31), 466(17, 22, 23, 31, 391, 467(17, 22, 23, 31, 39, 57, 58, 59, 601, 468(14, 22, 23, 30, 31), 469(60), 470 (40, 65, 661, 471(31, 35, 36, 40, 471, 472(31, 35, 36, 47, 71), 473(35), 474 (22, 23, 35, 39, 60), 475(22, 28, 35, 36, 38, 47, 60, 711, 476(23, 28, 35, 47, 711, 477(36, 71), 478(14, 17, 22, 29, 60, 76), 479(14, 22), 480(71, 76, 781, 481, 482(14, 23, 35, 52, 71, 75, 78, 821, 483 (52, 78), 484(23, 521, 485(54, 82) Varmus, H. E., 492 Varvoglis, A. G., 202, 205(7), 206(7), 208, 209(26), 211(7) Veeger, C., 356, 374(26), 375(26), 376(26) Venkataraman, P. R., 24, 25(176) Venkitasubramanian, T. A., 491, 495(41), 496(41), 501(41) Venn-Watson, E. A., 356 Verachtert, H., 52 Vernon, C. A., 232, 233(92) Vernon, R. G., 488, 489(5), 490(5) Veron, M., 108, 517, 518(29), 519(29), 520(29), 521(29), 522(29), 523(29), 533(29), 536, 537(57), 539(57)

Viala, B., 394, 461, 462(40), 470(40), 471 (40) Vidra, J. D., 54 Vignais, P. V., 309, 310(15), 315(15), 316 (151, 319, 321(15), 322(15), 325(15, 591, 326(15), 329(15), 330(15) Vijayvargiya, R., 377 Vilkas, M., 203 Villar-Palasi, C., 52, 54, 55, 56(25), 60, 62(25), 66(25), 378, 556, 563, 574(14) Vinopal, R., 110, 118(73) Vinuela, E., 247, 249(52), 261, 263(52, 123), 264, 266(123), 275(123) Virden, R., 390, 395, 460, 461(16, 271, 462 (16, 27, 411, 463(27, 41), 464(16), 466 (41, 48), 467(41, 59), 468(16), 469, 471(16), 472(16, 48), 476(48), 478 (771, 479(48), 480(48, 77), 482(16), 483(48), 485(77) Viswanathan, P. N., 52, 56(8), 57(8) Vitols, E., 144, 145(75), 147(75), 148(75), 150(75), 151(75), 152(75) Volcani, B. E., 148 von Funcke, H. G., 566, 574(70) von Korff, R. W., 6, 12(18)

W Waddy, C. T., 572 Wadkins, C. L., 280, 281, 289(17), 291 (17), 292(17), 294(17), 296(17), 297 (17), 298(18), 299(18), 301(18), 302, 303(18), 309, 310(28, 421, 313(28), 314(28), 321(28), 325(28), 326(28), 329(28), 330(28) Wakid, N., 240, 241(12), 242(12), 259(12) Wakil, S. J., 6, 12(18), 156, 158, 159(16), 165, 166(16), 167, 168(16), 169(16), 170(16), 172, 176(9), 178, 179, 180 (63), 183(63), 186, 187(70), 188, 193 Wakisaka, G., 145 Wakselmann, M., 203, 222, 225(75), 226 (75) Waldenstrom, J., 9, lO(62, 671, 13(62, 67) WBlinder, O., 232, 233(106), 308, 309(6), 310(20), 312(6), 313(6), 314(6), 315 (6), 319(6, 18, 20, 21, 231, 320(18), 326(18), 329 Walker, D. G., 488, 489(5), 490(5)

614 Walker, G. A., 144, 145(75), 147(75, 831, 148(75), 150(75), 151(75), 152(75) Walker, I. O., 255 Walker, P., 357, 374(44), 376(44) Wall, R. E., 219 Wallace, A., 281 Walsh, B. T., 61 Walsh, C. T., 342, 350(39), 483 Walsh, D. A., 556, 559, 561, 563(42), 565, 566(12, 41), 567, 568(12), 569(89, go), 570(74, 89, 901, 571(89, 901, 572(12, 741, 573(12, 35, 901, 574(12, 35, 401, 576(89, 901, 577(74, 101, I N ) , 578 (74,891, 579,580 Walter, G., 48 Walter, U., 365 Walters, R. A., 579 Walton, G. M., 247, 263(54), 267(54), 286, 567, 574(79) Wampler, D. E., 512, 518(18), 519, 520 Ward, C., 57,61(74) Ward, J. B., 117 Ward, R., 370 Warner, R. C., 2, 3(2), 232, 233(102) Warren, S. G., 201 Wassef, M. K., 491, 501(34) Wasson, G., 178 Watanabe, M., 26 Watanabe, S., 340, 341(28), 342(28), 343, 347 (28) Watson, H. C., 341, 344(36), 345(43), 348(42), 350(43), 359 Watts, D. C., 294, 384, 386, 387, 388, 389 (8, 33), 390(8), 392, 393(8, 9 , 395 (33, 341, 396, 400(8, 9, 33), 408(47), 409(47, 65, 871, 410(65, 90a), 412(65), 416(65), 419(47), 420(33), 421(31, 651, 423(65), 424(65), 425, 427(65), 431, 432, 433(91), 434, 435(140d, 437 (65), 438(65), 439(139), 442(139), 443(139), 444(139), 445(122), 446 (120), 447, 448(65, 139), 449(65, 1 3 9 , 453(65), 460, 461(16, 27), 462(16, 27, 41), 463(27, 411, 464(16), 466(41), 467(41, 591, 468(16, 441, 469, 471(16), 472(16), 478(77), 479(62), 480(77), 482(16), 485(77) Watts, R. L., 387, 395(34), 466, 467(59) Waygood, E. B., 377

AUTHOR INDEX

Weaver, R. H., 308, 309, 312(5), 314(5), 321(5), 324(5) Webb, M., 115 Weber, G., 280, 290(15), 299, 300, 302, 365, 377, 379 Weber, K., 47, 48(284), 49(284), 119, 519 Webster, L. T., Jr., 9, lO(53, 54), 12(54) Wedding, R. T., 309, 312(19), 319(19), 325(19) Wei, S. H., 563, 574 Weinhouse, S., 286 Weiss, B., 44, 46(272), 47(272) Weiss, S. B., 21, 22(135) Weissbach, H., 123, 144, 145(74), 146(74, 84), 14804, 79), 149(74), 150 Weissman, C.,23 Wells, R. D., 311(47), 312(47), 313, 332 (47) Welton, R. F., 488, 490(21) Wendell, P. L., 341, 344(36), 345(43), 348 (42), 350(43), 359 Wensel, K.-W., 254 Westhead, E. W., 45, 512, 518(18), 519 (301, 520, 523 Westheimer, F. H., 201, 204, 216(1), 217, 219, 223 (141, 229, 234 Wheldrake, J. F., 35(223, 2241, 36 Whigham, W. R., 356 White, A,, 416, 417(105), 422(105), 428 (105), 429(105), 430, 485, 486(90) White, J. G., 144 Whitehouse, M., 6 Whitesides, G. M., 215 Wiame, J. M., 492 Wieker, H.J., 232, 233(109), 356, 358 (131, 361, 375, 378(142) Wieland, O., 488(1), 489(10), 490(10), 492, 493(10, 46), 494(46), 495(10, 461, 496(10, 46), 497(10), 499(10, 461, 501 (10, 46), 502(10, 46), 566, 574(70) Wiesendanger, S. B., 510(10), 511 Wiesmann, U., 384,386 Wildes, R. A., 362 Wiley, J. H., 56 Wilgus, H., 241(23), 242(23) Wilkie, N. W., 22(175), 24, 25(175) Willecke, K., 158, 159, 160(18), 177, 181 (54), 186(54)

615

AUTHOR INDEX

Williams, A., 213 Williams, J., 555 Williams, L. D., 217 Williams, R. J. P., 115 Williams-Ashman, H. G., 124, 569, 570 (92), 572(92), 577(92) Williamson, I. P., 183, 187(70), 188 Williamson, J. R., 161, 162(28), 275, 277 Wilson, L. G., 35(230), 36 Wilson, R. H., 361 Wilson, R. J. H., 358 Wilson, R. M., 148 Wilson, T. H., 492 Wilt, E. M., 47 Wimpenny, J. W. T., 115 Winder, F. G., 491, 492(44) Winton, B., 566, 574(70) Wirts, G. W., 12(130), 19 Wittkop, J., 77, 86(23), 87(23), 88(23), 89(23), 9303) Witsel, H., 232, 233(109) Wohlhueter, R. M., 44 Wolf, D., 43, 44 Wolf, G., 35(222), 36 Wollenberger, A., 56 Wong, K. K., 9, lO(56) Woo, s. L. c.,574 Wood, H. G., 2, 3(3), 6, 12(3), 18 Wood, T., 258, 375, 384(18), 385, 401(18) Wood, T. R., 144 Woods, A. E., 365 Woods, H., 374 Wool, I. G., 574 Wort, D. J., 56 Wosilait, W. D., 556 Wright, A., 54 Wright, B. E.. 54, 56, 57, 61(53, 74), 62 (27), 67(27), 249, 263(66), 264(66), 267(66 Wright, C. S., 454 Wright, N. G., 504, 510, 511, 512, 513, 533 Wright, R. S., 218 Wu, L., 308, 315(12), 320(12) WU, N.-C., 565 Wu, R., 262(115), 264, 275 Wulff, K., 41, 43(246) Wiister, K-H., 373, 375(127) Wyman, J., 267, 373, 526, 547(50)

Y Yagiri, Y., 145 Yamada, H., 82 Yamada, M., 14, 501 Yamada, T., 30 Yamamura, H., 570, 580 Yamashita, O., 55, 56(44) Yang, D. L., 6(47), 7, 13(47), 18(47) Yang, K. V., 202,231 Yang, P. C., 160 Yarus, M., 9, 10(59), 13(59) Yashikawa, H., 31(219), 35 Yates, R. A., 240 Yeh, J., 41 Yielding, K. L., 377 Yorke, R. E., 278 Yoshida, A., 340, 341(28), 342(28), 343, 347(28) Yoshida, M., 240(32, 33)) 243, 263(32, 33), 264(32, 33), 266(32), 267(32, 33) Yoshikawa, H., 340, 341(31) Yoshino, T., 145 Younas, M., 207, 209(32), 210, 212, 225 (32), 226(33) Younathan, E. S., 248, 255, 256(83), 270 (83), 271(144), 273(144) Young, D. L., 18 Yourno, J., 522, 538(42) Yphantis, D. A., 118,119 Yudelevich, A., 45(276), 46, 47(276) Yue, R. H., 232, 233(106), 308, 315(12), 316(11), 320(12), 384(17), 385, 389 (42), 390(42), 391(42), 392(42), 393 (42), 394(42), 395(61), 399(42), 401 (171, 402(17) Yugari, Y., 511(15), 512 Yunis, A. A., 564 Yurowitski, Yu. G., 357, 374(41)

Z Zachau, H. G., 9(85), 10, W85) Zahlten, R. N., 575 Zamecnik, P. C., 9, 13(83), 24 Zanean, G. T., 68 Zetsche, K., 56, 6064) Zetterqvist, b., 232, 233(106), 308, 309 (6), 312(6), 313(6), 314(6), 315(6), 319(6, 18), 320(18), 326(18)

616 Ziegler, H., 56 Zielke, C. L., 269 Zillig, W., 9, 13(80), 47, 48(282, 283) Zimmerman, G., 254 Zimmerman, S. B., 45(275), 46, 47(275)

AUTHOR INDEX

Zimmermann-Telschow, H., 356, 358 Ziter, F., 395, 399(63a), 400(63a), 401 (63a), 403(63a) Zon, G., 213 Zwaig, N., 492, 502(67), 503

Subject Index A Acetabularia, uridine diphosphoryl glucose pyrophosphorylase in, 60 Acetate activation of, 12 phosphate rster hydrolysis, ratc constants, 211, 212 Acetate ions, creatine kinasc and, 424425, 427

Acctoacetate formation of, 156-157, 165 vaccenate synthesis nnd, 157-158 Acetoacetyl acyl carrier protein synthesis, 194 assay of, 190 Acetyl acyl carrier protein, j3-ketoacyl acyl carrier protein synthetase and, 191, 193

Acetyl adenylate-enzymc complex, isolation of, 10 N-(N-Acetyl-/3-alanyl) cystearnine, malonyl coenzyme A-acyl carrier protein transocylase and. 180 Acetyl coenzyme A acyl carrier protein and, 156157, 164-

N-Arctylcysteamine, malonyl coenzyme A-acyl carrier protein transacylase and, 180 N-Ace tylimidazole arginine kinase and, 481 phosphofructokinase and, 273 Acetyl phosphate, hydrolysis and enzyme-catalyzed transfer reaction, 233

Acetylsalicylate, creatine kinase and, 427 0-Acetylserine, methionine adenosyltransferase and, 137 Acidic nuclear protein kinases, properties, 580 cis-Aconitate, phosphofructokinase and, 265

Actinomycin D, uridine diphosphoryl glucose pyrophosphorylase synthesis and, 61 Acyl adcnylate formation, evidence for, 8-10 hydrolysis, standard free energy, 4 Acyl carrier protein(s) amino acid composition, 158-159 distribution and intracellular loration, 158-164

165

formation, mechanism, 3-4, 6 /3-ketoacyl ncyl carrier protein synthetasc alkylation and, 194-195, 197 malonyl coenzyme A-ncyl carrier protein transacylase and, 180 pyruvatc formation from, 83 pyruvnte kinasc and, 377 Acetyl coenzyme A-ncyl carrier protein transacylase catalytic properties, 187-188 fntty acid biosynthesis and, 165 historical background, distribution and metabolic significance, 185-186 molecular properties, 186-187 617

fatty acid biosynthesis and, 164-165 function, 155-156 historical background, 156158 physical properties, 166 prosthetic group primary sequence and, 166-170 synthesis and turnover, 173-176 source, fatty acid products and, 172 structure-activity relationships, 170173

Acyl carrier protein hydrolasc, properties of, 174-175 Acyl phosphatase, phosphoryl transfer and, 233

618

SUBJECT INDEX

Adenine, protein kinase and, 577 Adenine myonic acid dinucleotide, synthesis of, 31, 33,35 Adenosine creatine kinase and, 407 phosphodiestcr derivatives, biosynthesis, 20-30 3-phosphoglycerate kinase crystals and, 344, 348 protein’ kinases and, 577 Adenosine diphosphate adenosine diphosphoryl glucose pyrophosphorylases and, 34, 74, 76 Aeromonas formicans, 109 Entner-Doudoroff pathway and, 7980

Escherichia coli, 99-103 Rhodospirillum rubrum, 81 Serratia marcescens, 107 spinach leaf, 88-89 adenylate kinase and, 299 arginine kinase and, 476, 482, 483-484, 485 creatine kinase and, 407, 408, 409, 411, 412-419, 422, 424-426, 433, 435, 436, 437, 439, 441, 449, 450, 453 glycerol kinase and, 501 hydrolysis, rate, 204 lombricine kinase and, 476 metal ion complexes, 235 methionine adenosyltransferase and. 135 nucleoside diphosphokinases and, 323 oscillations in concentration, 276 phosphofructokinase and, 250-252, 254, 259, 261-263, 264, 266, 271, 275 3-phosphoglycerate kinase and, 336, 337, 346, 348-351 phosphoryl transfer and, 233 pyruvate kinase and, 360, 361, 364, 368, 369, 371, 375, 380, 382 synthesis of derivatives adenine-myonic acid dinucleotidr and adenylyl diphosphoglycerate, 33-35 adenosine diphosphoglucose, 32-33 general features, 30-32 Adenosine diphosphoryl glucose adenosine diphosphoryl glucose pyro-

phosphorylase and, 118 activator effects, 97-98 Aeromonas formicans, 109 Chlorella pyrenoidosa, 90 Entner-Doudoroff pathway and, 79 green leaves, 86-90 Serratia marcescens, 107 starch synthesis and, 57-58 synthesis of, 31, 32-33 Adenosine diphosphoryl glucose pyrophosphorylase activator-inhibitor interaction, 99-100 activators, kinetic parameters and, 9799 classification of, 75-77 compounds causing no stimulation, 95 energy charge and, 104-107 mutants, 113-114 kinetic properties Aeromonas formicans, 108-109 Enterobacteriaceae, 94-107 Escherichin coli mutant, 109-117 general effects of activators, 77-78 organisms using Entner-Doudoroff pathway, 78-81 plants and algae, 86-94 Rhodospirilliim rubrum, 81-86 Serratia marcescens, 107-108 nonchlorophyllous plant tissue, 93-94 physical properties, 117-119 reaction catalyzed, 74 regulation of, 74 substrate and activator kinetic constants, inhibitors and, 101-102 Adenosine 5’-hypophosphate, pyruvate kinase and, 365 Adenosine monophosphate adenosine diphosphoryl glucose pyrophosphorylase, 34, 74, 75, 76 activator interaction, 99-100 Entner-Doudoroff pathway and, 7980 manganese ions and, 103 mutants and, 112-114, 116-117 Rhodospirillum rubwm, 81 Serratin marcesceiis, 107 substrate and activator kinetir constants, 101 adcnylate kinase and, 298-300

SUBJECT INDEX

creatine kinase and, 407 enzyme control by, 286 exchange into adenosine triphosphate, 8 glycerol kinase and, 503, 504 methionine adenosyltransferase and, 135 phosphofructokinase and, 254, 258, 259, 261-263, 266, 271, 272, 275 3-phosphoglycerate kinase and. 348 phosphorylasc and, 557. 558 protein kinase and, 577 pyruvatc kinasc and, 370, 374 Adenosine T.5’-monophosphate analogs, protein kinases and, 573 hydrolysis, free energy, 27 mechanism of action artivation mechanism, 570-572 kinetic observations, 568-570 phosphofructokinase and, 259, 260-263, 266, 271, 272, 274 glycolysis and. 278 protein kinases and. 556, 561, 565 5’-Adenosinemonophosphate aminohydrolase, see Adenylate deaminasc Adenosine 5’-monophosphoramidate, formation and function, 48 Adenosine 5’-phosphosulfate. synthesis of, 35-37 Adenosine tctraphosphate, methionine adenosyltransferase and, 135 Adenosine triphosphatase methionine adenosyltransferase and. 128, 129 phosphoryl transfer and, 233 vitamin BIZ. adenosyltransferase and, 146 Adenosine triphosphate adenosine diphosphoryl glucose pyrophosphorylase, 118 activator effects, 97-99 Aeromonas formicans, 109 Entner-Doudoroff pathway and. 78. 79, 81 inhibitor effects, 100, 101, 102 manganese ions and, 103-104 mutants and, 115 plant tissues, 93 Rliodospirillum riibrum, 81, 85-86

619 Serratia marcescens, 107 spinach leaf, 87-90 adenylate kinase and, 295, 299 analog, binding by aspartokinase I and, 524-525 arginine kinase and, 476, 482, 485 argininosuccinate synthesis and, 37-39 nspartokinases and, 512, 520, 525, 541, 543, 545, 550, 551, 552, 553 creatine kinase and, 395-396, 406, 407, 410, 412419, 421, 422, 424-426, 427, 433, 437, 441, 443, 449, 450, 453 deoxyribonucleic acid ligase and, 45-47 energy charge and, 105 enzyme control by, 285-286 glycerate kinase and, 507 glycerol kinase and, 498501 gramicidin synthesis and, 14-15 guanosine monophosphate synthesis and, 3940 hydrolysis rate, 204 standard free energy, 2 4 labeled, nucleoside disphosphokinase and, 315, 319 lipoatc activation and, 17-18 lysine-sensitive aspartokinase and, 45 methionine adenosyltransferase and, 123, 125, 130, 134-135, 136, 138, 140 methylene analogs, methionine adenosyltransferase and, 135 nucleoside diphosphokinase kinetics and, 322, 327, 328, 331 oscillations of concentration, 276 phosphofructokinase and, 241, 243, 246, 247, 248, 249-252, 258, 259-260, 261-263, 264, 265, 266, 267, 269, 27C272, 273, 274, 275 3-phosphoglycerate kinase and, 336, 337, 339, 341, 346-347, 34M51 phosphorylase kinase and, 557, 560561, 563-564, 565 phosphoryl transfer and, 233 protein kinases and, 569, 571, 572, 576, 577 pyruvate kinase and, 360, 363, 364, 368, 369, 375-376, 379-380, 381 sulfate activation and, 35-37

620

SUBJECT INDEX

vitamin BIZ. adenosyltransferase and,

carboxyl group activation and, 6-20 functions of, 5-6 Adenosine triphosphate(cytidine triphos- Adipose tissue glycerol kinase in, 489-490, 493 phate) : transfer ribonucleic acid nuphosphofructokinase of, 262 cleotidyltransferase, adenylyl group Aerobacter aerogenes transfer by, 25 adenosine diphosphoryl glucose pyroAdenosine triphosphate :glucose-l-phosphosphorylase of, 75, 76 phate adenylyltransferases, regulaglycogen synthesis, regulation of, 34 tion of, 33, 34 phosphofructokinase of, 245, 250 Adenosyl compounds, nomenclature, 123 Aerobacter cloacae BAdenosylhomocysteine, methyl transadenosine diphosphoryl glucose pyrofer and, 124 phosphorylase of, 75, 76 S-Adenosylmethionine glycogen synthesis, regulation of, 34 cystathionine-y-synthaee and, 142 Aeromonads, characteristics of, 108 methionine adenosyltransferase kinetAeromonas formicans ics and, 132, 135, 137-139 adenosine diphosphoryl glucose pyrotripolyphosphataae and, 130-131, 133 phosphorylase of, 75, 76, 108-109 ultraviolet absorption, 129 glycogen synthesis, regulation of, 34 Adenylate deaminase Age, methionine adenosyltransferase adenylate kinase assay and, 301 and, 142 kinetics of, 269 Agrobacterium tumefaciens nucleoside diphosphokinase assay and, adenosine diphosphoryl glucose pyro325 phosphorylase of, 76, 78-81 Adenylate kinase glycogen synthesis, regulation of, 34 catalytic properties Alanine assay, 300 aspartokinase and, 547 equilibrium constants, 302 pyruvate kinase and, 372, 376-377 mechanism, 302-305 p-Alanine metal requirement, 297-298 activation of, 12 nucleotide specificity, 298-300 acyl carrier protein and, 158, 159, 161, distribution, 280-282 167 function, 285-288 Aldolase, phosphofructokinase assay genetics and disease, 282284 and, 244 historical background, 280 Algae molecular properties adenosine diphosphoryl glucose pyrocomposition, 291-293 phosphorylase of, 76, 77 physical properties, 295-297 starch synthesis, regulation of, 34, 74 preparation and purity, 288-291 Alkaline phosphatase, phosphorothioates reactive groups, 293-295 and, 213 nucleoside diphosphokinase assay and, Alloxan 324 methionine adenosyltransferase and, reaction catalyzed, 279 143 Adenyl cyclase phosphofructokinase and, 269 adenylyl transfer and, 26-27 Allylglycine epinephrine and, 561 aspartate semialdehyde synthesis from, Adenylyl-2,3-diphosphoglycerate, synthe513 sis of, 31, 35 aspartokinase and, 547 n-N-Amidinoproline, creatine kinase Adenylyl group transfer adenosine triphosphate and, 2-3 and, 405, 406-407 145-147, 149, 150

SUBJECT INDEX

L-N-Amidinoproline, creatine kinase and, 405, 406-407 Amino acids activation of, 6, 12-13 acyl carrier protein composition, 158159

adenylate kinase composition, 291-293 arginine kinase composition, 469470 aspartokinase I composition, 520-521 aspartokinase I1 composition, 542 aspartokinase I11 composition, 543 bacillus aspartokinase composition, 548 creatine kinase composition, 390392 muscle-type, 389 brain-type, 390 glycerol kinase composition, 494 phosphofructokinase composition, 255256, 257

3-phosphoglycerate kinase, 338, 340, 343

pyruvate kinase composition, 358 Amino acyl transfer ribonucleic acid, synthesis, 10-11 1-Aminocyclopentane carboxylate, methionine adenosyltransferase and, 136137

Amino glycoside antibiotics, adenylylation of, 27-30 2-Amino-4-hexenoic acid, methionine adenosyltransferase and, 136 2-Amino-4-hexynoic acid, methionine adenosyltransferase and, 136 Amino peptidase, malonyl coenzyme A-acyl carrier protein transacylase peptide and, 182 2-Amino-9-p-D-ribofuranosylpurine-5’triphosphate, phosphofructokinase and, 245 pdminosalicylate, creatine kinase and, 427

Ammonia, guanosine monophosphate synthesis and, 39 Ammonium chloride glycerol kinase and, 501 protein kinase and, 576 Ammonium ions aspartokinases and, 546, 549 glutamine synthetase and, 43 methionine adenosyltransferase and, 134

phosphofructokinase and, 248, 261, 276 tripolyphosphatase and, 134 pyruvate kinase and, 366,367 vitamin BIZ. adenosyltransferase and, 151-152

Anaerobiosis, phosphofructokinase and, 274-275

Androgen, methionine adenosyltransferase and, 143 Anemias, pyruvate kinase and, 354 Anions, creatine kinase and, 396, 423-427. 449

Annelida, guanidino kinases of, 461, 462 Antibiotics, resistance factors and, 27-30 Antibody adenylate kinase and, 283 creatine kinase and, 394, 436, 438 guanidino kinases and, 470-471 phosphofructokinase and, 258, 268 Antigens microbial, synthesis of, 59, 67-68 Arabinitol diphosphate, adenosine diphosphoryl glucose pyrophosphorylase and, 94-95 Arabinose 5-phosphate, phosphofructokinase and, 251-252 Arginine, binding by arginine kinase, 486

o-Arginine, arginine kinase and, 474 Arginine kinase amino acid composition, 469-470 sequences, 432 bivalent metal ions and, 472 carboxymethyl, arginine binding by, 480

creatine kinase hybrid, 387, 395 dead-end complexes, 482 difference spectra, 433, 434 discovery and isolation, 45-60 distribution of, 461463 equilibrium, 486 essential thiol groups of, 478, 479, 480 a-helix in, 394 immunological properties, 470, 471 molecular weights, 466, 467 nucleotide specificity, 476 partial reactions, 483-484 peptide, reactive thiol group and, 470 reaction mechanism, 482-484

622

SUBJECT INDEX

stability, 468-469 substrate specificity, 474, 475 subunits, 468 Arginine residues, phosphofructokinase, 255, 256, 257

Argininosuccinate biosynthesis, acceptor molecule and, 11

synthesis of, 38-39 Arsenate, creatine kinase and, 426 Arsenicals, pyruvate kinase and, 361 Arthrobacter ncyl carrier protein amino acid composition, 159 4’-phosphopantetheine peptide of. 169, 170

Arthrobacter crystallopoietes, phosphofructokinase of, 263, 264 Arthrobacter vi.~cosus adenosine diphosphoryl glucose pyrophosphorylase of, 76, 78-81 glycogen synthesis, regulation of, 34 Arthropoda, guanidino kinase of, 462 Asrites tumor phosphofructokinase, properties of, 242, 262

Aspartate aspartokinase I and, 520, 527, 529, 532 aspartokinase I1 and, 541 aspartokinase I11 and, 543 aspartokinases and, 545, 547, 549, 550, 551, 552

homoserine production from, 510 Aspartate-p-semialdehyde aspartokinase I and, 514-520 aspartokinase I1 and, 541 aspartokinases and, 551, 552 homoserine biosynthesis and, 510 synthesis of, 513 Aspartate semialdehyde dehydrogenase, aspartokinase assay and, 512 Aspartokinase ( 8 ) historical background, 509-511 lysine-sensitire, adenylylation of, 4445

methods of assay, 512-513 other coliform bacteria, 544 reaction catalyzed, 511-512 regulated by concerted feedback inhibition

Bacilli, 54&551 other genera, 552 other nonsulfur photosynthetic bacteria, 545-546 pseudomonads, 551-552 Rhodopseudomonas capsulatus, 544545

synthesis, repression of, 513-514 three isofunctional enzymes, 513-515 Aspartokinase I chemical properties amino acid analysis, 520-521 identity of subunits, 622 partial sequences, 521-522 sulfhydryl groups, 521, 525-526 conformational changes circular dichroism and optical rotatory spectra, 535-536 difference spectra, 526-527 ligand effects on fluorescence, 527533

relaxation studies, 533-535 distribution of two activities on polypeptide chain, 536-540 extinction coefficient, 517 homoserine dehydrogenase I and, 515516, 524-525

kinetic parameters, 519-520 ligand binding adenosine triphosphate analog, 524525

pyridine nucleotide, 523-524 threonine, 523 molecular weight, 517-518 purification and criteria of homogeneity, 516 stability, 517 tetrameric structure, 518-519 Aspartokinase I1 amino acid composition, 541-542 extinction coefficient, 541 kinetic parameters, 541 molecular weight, 541 purification and criteria of homogeneity, 540 stability, 540 subunit structure, 541 Aspartokinase I11 amino acid comDosition. 543 extinction coefficient, 542

SUBJECT INDEX

Benzimidazolecobamide, vitamin B,, adenosyltransferase 'assay and, 148, 151 Benzoate activation of, 6, 12 creatine kinase and, 427 Bicarbonate creatine kinase and, 425, 426 pyruvate kinase and, 364, 382 Bimolecular mechanisms, enzymic phosphoryl transfer and, 235-238 Biotin, attachment to enzymes, 6, 13 Biotin activating enzyme, adenylyl group transfer, 18 Birds creatine kinases of, 387-388 nucleoside diphosphokinases of, 311 Bis-2,4-dinitrophenyl phosphate, hydrolysis of, 231 Blue Dextran, pyruvate kinase and, 356 Borate ions, creatine kinase and, 426 Borofluoride ions, creatine kinase and, B 426 Bacillus brevas, peptide synthesis by, 11, Boroliydride, vitamin Bls. and, 147, 148 14-17 Brain Bacillus cereus, aspartokinase of, 551 creatine kinase, 386 Bacillus licheniformis, aspartokinase of, amino acid composition, 390 551 nucleoside diphosphokinases of, 310, Bacillzts polymyxa 317 aspartokinase phosphofructokinase, properties of, 242, inhibition, 546-547 249, 258, 262, 265, 266, 275 purification and properties, 547-548 protein kinase in, 567, 569, 581 subunits, 548 Brevibacterium fEavum, aspartokinase of, Bacillus stearothermophilus, aspartoki552 nase of, 550-551 Brevibacterium liquefaciens, phosphoBncillus subtilis fructokinase of, 259 aspartokinases of, 548550 Bromide ions, creatine kinase and, 411nucleosidc diphosphokinase of, 312, 412, 425, 426, 452 315. 316. 319. 322-323, 329 Brussels sprouts, phosphofructokinase of, Bacteria 247, 262 coliform, aspartokinases of, 544 Buffer glycogen synthesis by, 74 adenosine diphosphoryl glucose pyropyruvate kinases of, 357, 366, 374 phosphorylase and, 87-88 Barium ions phosphofructokinase and, 253, 265 ndenylate kinase and, 298 n-Buty lamine creatine kinase and, 450 phosphate ester hydrolysis, rate conB d e y leaf, adenosine diphosphoryl g 1 ~ stants, 211 cose pyrophosphorylase of, 90 Butyrate, creatine kinase and, 427 Benzene sulfonate. creatine kinase and, Butyryl acyl carrier protein, fatty acid 427 biosynthesis and, 165 inhibition of, 544 kinetic parameters, 543 molecular weight, 543 purification and criteria of homogeneity, 542 P-Aspartyl phosphate homoserine biosynthesis and, 510 synthesis of, 553 Associative mechanisms, enzymic phosphoryl transfer and, 235-238 Aurocyanide, 3-phosphoglycerate kinase crystals and, 344345 Avocado acyl carrier protein, amino acid composition, 159 8-Azaguanosine diphosphate, nucleoside diphosphokinase and, 323, 324 6-Azauridine diphosphate. nucleoside diphosphokinase and, 323 Azotobacter, aspartokinases of, 552

624

SUBJECT INDEX

C Cadmium ions 3-phosphoglycerate kinase nnd, 349 pyruvate kinase and, 370 Calcium ions adenosine diphosphntc complex, 235 adenylate kinase and, 298 arginine kinases and, 471, 472 creatine kinase and, 409-412, 416, 417, 429, 430, 450 glycerate kinase and, 508 glycerol kinase and, 501 nucleoside diphosphokinases and, 329 phosphofructokinase and, 247 3-phosphoglycerate kinase and, 349 phosphorylase kinase and, 562, 564 protein kinrlses and. 573 pyruvate kinase and, 377 Canavanine, arginine kinase and, 474 0-Carbamylserine, methionine adenosyltransferase and, 137 Carbon dioxide fixation, 3-phosphoglycerate kinase and. 337 glycogen accumulation and, 82 Carboxyl groups acyl carrier protein, modification of, 173 Carboxyl group activation Rdenylyl group transfer acylation of Nc-lysyl residues of enzymes, 17-18 adenylation of luciferin and dehydroluciferin, 19-20 fatty ncid and amino acid activation. 6-11 nucleic acid-independent peptide synthesis, 11-17 Carboxymethylcysteine, P-ketoacyl acyl carrier protein synthetnse and, 194195 1-Carboxymeth yl-2-iminohexahydropyrimidine, creatine kinase and, 405, 406 l-Carboxymethyl-2-iminoimidazolidine, creatine kinase and, 405, 406 Carboxypeptidase, creatine kinase and, 393

Carboxypeptidase A . . acyl carrier protein and, 171 aspartokinase I and, 522 2-Carboxyphenyl phosphoramidate. hydrolysis of, 221 Cnrnosinc. synthesis of, 6, 12 Carp muscle, creatine kinase of, 389 Carrots, phosphofructokinase of, 262 Casein phosphorylase kinase and, 558 protein kinase and, 566, 574, 575, 576, 577, 580 Cntecholnmines, glycolysis and, 278 Cations methionine adenosyltrnnsfernsc and, 133-134 monovalent, pyruvate kinase and, 354, 366-368, 372 phosphofructokinnse nnd, 247-248 uridine diphosphoryl glucose pyrophosphorylase and, 62, 70-71 Cellulose synthesis, uridine diphosphoryl glucose pyrophosphorylase and, 57 Cell walls polymers, synthesis of, 59,60 Cetyltrimethylammonium bromide, phosphate ester hydrolysis and, 231 Chicken muscle, creatine kinnse of, 389 Chlamydomonas reinhardii, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Chlorella pyrenoidosa, adenosine diphosphoryl glucose pyrophosphorylase of, 90. 92 Chlorelln vulgaris, adenosine diphosphoryl glucose pyrophosphory1nse of. 90 Chloride ions arginine kinase and, 485 creatine kinase and, 411-412, 415, 416, 417, 423-427, 436, 438, 452 Chloroacetate, creatine kinase and. 427 Chloroazanil, uridine diphosphoryl glucose pyrophosphorylase and, 68 p-Chloromercuribenzoate, glycerol kinase and, 496

SUBJECT INDEX

malonyl coenzyme A-acyl carrier protein transacylase and, 183, 184 nucleoside diphosphokinase and, 330 3-phosphoglycerate kinase and, 338, 346, 350 7-Chloro-4-nitrobenzo-2-oxa-l,3-diazole, adenylate kinase and, 293-294 p-Chlorophenylalanine, pyruvate kinase and, 377 Chloroplasts, uridine diphosphoryl glucose pyrophosphorylase in, 56 Cholate, activation of, 6, 12 Chordata, guanidino kinases of, 463 Chromatography, adenylate kinase assays and, 300301 Chromosome, 3-phosphoglycerate kinnse gene location, 339-340 Chymotrypsin. aspartokinase I and, 522, 536

Circular dichroism, aspartokinase I, 535536 Citrate concentration, fatty acids and, 277 phosphofructokinase and, 258, 261-263, 265-266, 272, 275, 278 3-phosphoglycernte kinase and. 340 pynivate kinase and. 363 Citrobnctei freundii adenosine diphosphoiyl glucose pyrophospliorylase of, 75, 76, 96 glycogen synthesis, regulation of, 34 Clostridium butyriciim ncyl carrier protein, amino acid composition, 159 Clostiidium pasteuriairum phosphofructokinase, properties of, 242, 253, 256, 263, 267 Clostridium tetanomorphum, vitamin BIZ, adenosyltransferase of, 144, 145 Cobalamins, nomenclature, 122 Cobalt ions adenosine diphosphoryl glucose pyrophosphorylase and, 102-103 adenylate kinase and, 298 creatine kinnse and, 409 glycerol kinase and, 501 guanidino kinases and. 472 nucleoside diphosphokinases and. 329 phosphofructokinase and, 247 3-phosphoglycerate kinase and, 349

625 protein kinases and, 573, 578 pyruvate kinase and, 369 Coelenterata, guanidino kinases of, 462 Coenzyme A acyl carrier protein and, 171, 173-174, 175 carboxyl group activation and, 12 distribution in cells, 164 luciferase inhibition and, 20 Copper ions, pyruvate kinase and, 376 Corn, phosphofructokinase of, 262, 266 Creatine active form of, 414 arginine kinase and, 474 binding to creatine kinase, 436, 437, 439, 451452 Creatine kinase activating metal ion, 409-412 adenylate kinase assay and, 301 brain-type, other, 402-403 catalytic site, formation and topography, 439442 conformational changes, substrate-induced, 436-438 dead-end complexes, 416, 418, 423, 425, 436, 441, 444, 449, 453 distribution of, 461463 “essential” thiol group effects of substrates, 448-451 importance for catalytic activity, 443448 structural involvement, 4 4 U 4 3 general considerations, 384-385 hybrid, preparation of, 403 kinetic constants, 405 kine tics active forms of substrates, 412414 anion effects, 423427 equilibrium, 428431 nucleotides as inhibitors, 422 substrate binding, 414-420 temperature effects, 420422 mechanism of transphosphorylation, 451455 metal-bound nucleotide and. 303 muscle assay and specific activity, 400 purification, 400 stability, 401

626

SUBJECT INDEX

ox brain assay and specific activity, 401 purification, 401 stability, 401402 peptide, reactive thiol group and, 470 phosphoryl transfer and, 233 rabbit muscle assay and specific activity, 395-398 purification, 395 stability, 398-399 reactive groups essential for activity cysteine residues and number of catalytic sites per molecule, 431432

histidine, 434 lysine, 432434 tyrosine. 434-436 structure nmino acid composition, 390-392 isoenzymes, interspecific hybrids, conformers and genetic variants, 386390

molecular weight, 395 primary structure, 392-393 secondary and tertiary structure, 393-394

subunit shape and organization, 394395

substrate specificity guanidine substrates and organization of the creatine binding site, 403407

nucleotide substrates and related inhibitors, 407409 Crotonyl acyl carrier protein, fatty acid biosynthesis and, 165 Cyanogen bromide, acyl carrier protein and, 171, 172 Cyclic 3',5'-nucleotide phosphorodiesterase, phosphorothioates and, 213 Cycloheximide, uridine diphosphoryl glucose pyrophosphorylase synthesis and, 01 Cystathionase, mutants and, 141 Cystathionine-y-synthase, mutants and, 141, 142

Cysteine creatine kinase assay and, 396, 398 glycerol kinase and, 496

methionine adenosyltransferase and, 137

pyruvate kinaae and, 372 Cysteine residues acyl carrier proteins, 158, 159 crcatine kinase, 392, 431432, 440 effect of substrate, 448451 importance for catalytic activity, 443-448

structural involvement, 442-443 giianidino kinases, 469-470, 477-480 P-ketoacyl acyl carrier protein synthetase, 195, 197-198 phosphofructokinase, 272 3-phosphoglycerate kinase, 342, 343, 344, 350

pyruvate kinase, 360-361 Cystine, sulfate activating enzyme and, 37

Cytidine diphosphate arginine kinase and, 476 rreatine kinase and, 408, 409, 416, 417 nucleoside diphosphokinases and, 323 pyruvate kinase and, 364 Cytidine diphosphoryl glucose formation of, 68 synthesis of, 32 Cytidine triphosphate adenosine diphosphoryl glucose pyrophosphorylase and, 81 adenylate kinase and, 299 glycerol kinase and, 498, 499 methionine adenosyltransferase and. 135

nucleoside diphosphokinases and, 322 phosphofructokinase and, 246, 247, 271 protein kinases and, 572 vitamin BIs8adenosyltransferase and, 150-151

Cytochronie oxidase, aeromonads and, 108

Cytosol, protein kinase in, 567

D Deamido nicotinamide adenine dinucleotide activation of. 13 synthesis of, 31-32 Decanoyl acyl earrier protein, p-keto-

SUBJECT INDEX

627

acyl acyl carrier protein synthrsis diphosphoryl glucose pyrophosand, 191, 193 phorylase and, 34, 76, 78 cis-3-Decenoyl acyl carrier protein, p- Dephosphocoenzyme A, synthesis of, 31 ketoacyl acyl carrier protein syn- Diacetyl-a-naphthol reagent, guanidino thetase and, 191, 193 kinase assay and, 465 Dehydroluriferin, adenylylntion of, 19- m-Diaminopinielate, aspartokinase and, 20 549 Drhydrolurifrryl-adenylate Dibenzylphosphate, dibcnzylphosphohydrolysis. free energy, 19-20 enolpyruvate hydrolysis and, 223 Denervation Dibenzylphosphoenolpyruvate,hydrolysis muscle, creatine kinase and, 386-387 of, 223 Diclyostelium discoideum Deoxyadrnosine diphosphate phosphofructokinase of, 249, 263, 264 adenylate kinase and, 299 uridine diphosphoryl glucose pyrophosnrginine kinase and, 476 phorylase of, 54, 60-62, 63 rreatine kinase and, 408, 409 Diet lombricine kinase and, 476 adenylate kinase and, 286-287 nucleoside diphosphokinase and, 324 methionine adenosyltransferase and, pyruvate kinase and, 364 142-143 Deoxyadenosine monophosphate, ndenylDiethylpyrocarbonate ate kinase and, 299 nrginine kinase and, 482, 484 Deoxyadenosine triphosphate rreatine kinase and, 434, 453 adenylate kinase and, 299 P,P,-Diethylpyrophospha te arginine kinase and, 476 hydrolysis, rate, 204-205 Inethioninr adenosyltransferase and. Diethylstilbestrol, pyruvate kinase and, 135 377 phosphofructokinase and, 246 1.5-Difluoro 2,44initrobenzene, creatine 3-phosphoglywrate kinase and. 348 kinase and, 393, 436 Deoxycytidine diphosphate Dihydroxyacetone, glycerol kinase and, nucleoside diphosphokinase and, 323, 497498 324 2,3-Dimercaptopropan-l-o1,creatine kipyruvate kinase and, 364 nase and, 431 Deoxyrytidine triphosphate, nucleoside 1-Dimethylamino naphthalene-hulfonyl diphosphokinases and, 323 chloride Deoxyguanosinr diphosphatc arginine kinase and, 481, 483-484 rreatine kinase and, 409 creatine kinase and, 433, 446 nucleoside diphosphokinasrs and, 323. N,N-Dimethylhydroxylamine, ppruvate 324, 326-327 kinase and, 365 Deoxyguanosinr triphosphate Dimethyl suberimidate adenylate kinase and, 299 aspartokinase I and, 519 nucleoside diphosphokinase and, 323, glycerol kinase and, 495 326-327 Dimethylsulfoxide, pyruvate kinase and. 3-phosphoglycerate kinase and, 348 363 Deoxyribonucleic acid ligase 2,4-Dinitro-l-fluorobenzene,see Fluoroadenylyl transfer functions of, 45-48 dini trobenzene phage induced, adenylyl donor, 47 2.4-Dinitrophenyl-l,3,2-dioxaphosphoriDeoxyribonucleic acid polymerase, nunane-2-oxides, pyridines and, 210cleosidr diphosphokinase activity of, 211 332 2.4-Dinitrophenylhydrazine,pyruvate ki2-Deoxyribose 5-phosphate, adenosine nase assay and, 371

628 2,4-Dinitrophenyl methyl phosphate, pyridines and, 210 2,4 (2,6)-Dinitrophenylphosphates hydrolysis of, 205, 231 pyridines and, 209, 210 Dioldehydrase, vitamin BIZ. adenosyltransferase assay and, 148 P,P‘-Di-y-phenylpropyl pyrophosphate hydrolysis, rate, 204 1,3-Diphosphoglycerate, 3-phosphoglycerate kinase and, 336, 347, 349 2,3-Diphosphoglycerate, phosphofructokinase and, 258, 266 Dipicolinic acid, aspartokinase and, 549, 551 Disulfide bonds, aspartokinase I, 521, 536 5.5’-Dithiobis(2-nitrobenzoate) arginine kinase and, 479 aspartokinase I and, 521, 525-526 creatine kinase and, 447 glycerol kinase and, 497 glycocyamine kinase and, 479 p-ketoacyl acyl carrier protein synthetase and, 190 phosphofructokinase and, 270, 271 3-phosphoglycerate kinase and, 338, 350 pyruvate kinase and, 360, 361 Dithiothreitol creatine kinase and, 398, 446 p-ketoacyl acyl carrier protein synthetase and, 190, 194 malonyl coenzyme A-acyl carrier protein transacylase and, 183 nucleoside diphosphokinase and, 330, 331 phosphofructokinase and, 253-254, 257, 271 Dodecanoyl acyl carrier protein, p-ketoacyl acyl carrier protein synthetase and, 191, 193 cis-5-Dodecenoyl acyl carrier protein, P-ketoacyl acyl carrier protein synthetase and, 191, 193 Dodecyl sulfate adenosine diphosphoryl glucose pyrophosphorylase and, 119 aspartokinase I and, 518-519, 522, 525 aspartokinases and, 548 creatine kinase and, 394

SUBJECT INDEX

glycerol kinase and, 495 P-ketoacyl acyl carrier protein synthetase and, 189-190 nucleoside diphosphokinase and, 330 phosphofructokinase and, 254, 255, 256, 270 3-phosphoglycerate kinase and, 342 phosphorylase kinase and, 559, 562 pyruvate kinase and, 359 Dogfish muscle, creatine kinase of, 389

E Echinodermata, guanidino kinases of, 462463 Echinoidea, guanidino kinases of, 461463 Electron paramagnetic resonance creatine kinase, 437438, 440441, 450 pyruvate kinase, 367-369 Energetics, methionine adenosyltransferase, 139-141 Energy charge adenosine diphosphoryl glucose pyrophosphorylase and, 104-107 mutants, 113-114 adenylate kinase and, 285, 286 definition of, 105 nucleoside diphosphokinases and, 332 phosphofructokinase and, 276 Enterobacteriaceae adenosine diphosphoryl glucose pyrophosphorylases, 94-97 activator effects on kinetic parameters, 97-99 activator-inhibitor interaction, 99100 energy charge and, 104-107 inhibitor effects on kinetic constants, 101-102 manganese ion effects, 104-107 Entner-Doudoroff pathway, adenosine diphosphoryl glucose pyrophosphorylase and, 75, 78-81 Enzymes Nc-lysyl residues, acylation of, 17-20 Epinephrine phosphorylase and, 558, 561 pyruvate dehydrogenase and, 566

629

SUBJECT INDEX

Equilibrium constants adenylate kinase, 299, 302 creatine kinase, 428431 nucleoside diphosphokinase, 320 Erythrocytes adenylate kinase in, 282 nucleoside diphosphokinases of, 308309, 310-311, 313-314, 316, 319, 322-323, 32C327, 329, 331 phosphofructokinase, properties, 242, 249, 257-258, 262 3-phosphoglycerate kinase, 339 amino acid composition, 343 kinetic constants, 347 molecular weight, 342 pyruvate kinase of, 354, 359

Erythrose 1,4diphosphate, adenosine diphosphoryl glucose pyrophosphorylase and, 95, 97 Escherichia aureseens adenosine diphosphoryl glucose pyrophosphorylase of, 75, 76 glycogen synthesis, regulation of, 34 Escherichia coli acetylcoenzyme A-acyl carrier protein transacylase of, 186-188 acyl carrier protein amino acid composition, 159 amino acid sequence, 168-169 4’-phosphopantetheine peptide of, 169, 170 physical properties, 166 adenosine diphosphoryl glucose pyrophosphorylase, 75, 76 activator specificity, 95 physical properties, 118-119 aspartokinases, 513-543 deoxyribonucleic acid ligase of, 47 fatty acid synthesis in, 161 glutamine synthetase, regulation of, 40-44

glycerol metabolism in, 492 glycogen synthesis, regulation of, 34 P-ketoacyl acyl carrier protein synthetase of, 189-199 lysine-sensitive aspartokinase of, 4445

malonyl coenzyme A-acyl carrier protein transacylase of, 178-185

methionine adenosyltransferase of, 141142

mutants, adenosine diphosphoryl glucose pyrophosphorylases of, 10% 117

phosphofructokinase, properties of, 242, 246, 247, 253, 256-257, 265 pyruvate kinase, 359

263,

uridine diphosphoryl glucose pyrophosphorylase in, 60, 67-68 Estradiol, methionine adenosyl transferme and, 143 l,N’-Ethenoadenosine triphosphate adenylate kinase and, 300 phosphofructokinase and, 245 Ethionine methionine adenosyltransferase and, 13.5136, 143

resistance to, 142 Ethoxyformic anhydride, phosphofructokinase and, 272 N-Ethyl-N-amidinoglycine, creatine kinase and, 4051 O-2’-(Ethyl 2diazomalonyl) cyclic adenosine monophosphate, phosphofructokinase and, 274 E thylenediaminetetraacetate creatine kinase and, 396 glycerate kinase and, 507, 508 glycerol kinase and, 495, 496, 500 p-ketoacyl acyl carrier protein synthetase and, 190 methionine adenosyltransferase and, 128

phosphofructokinase and, 253, 257 pyruvate kinase and, 363 vitamin Bas adenosyltransferase and, 151

Ethylene hydrogen phosphate, hydrolysis of, 217 N-Ethylglycocyamine, creatine kinase and, 403 N-E thylmaleimide acetyl coenzyme A-acyl carrier protein transacylase and, 187, 188 arginine kinase and, 479 aspartokinase I and, 523 creatine kinase and. 443 glycerol kinase and, 497

630

SUBJECT INDEX

p-ketoacyl acyl carrier protein synthctase and, 194 malonyl coenzyme A-acyl carrier protein transacylase and, 183, 184 Ethyl mercuric phosphate, 3-phosphoglycerate kinase and, 345 N-Ethylmorpholine, guanidino kinases and, 484 Euglena gracilis, acyl carrier protein in, 160-161

F Fatty acids activation of, 6, 12 glycolysis and, 277 unsaturated, biosynthesis of, 191 Fatty acid synthetase acyl carrier protein and, 157, 164-165 labeled acetyl peptides from, 186 labeled malonyl peptjde from, 177-178 Fatty acyl coenzyme A acyl carrier protein transacylation, pketoacyl acyl carrier protein synthetase and, 196-198 Fatty acyl thioesters, acetyl coenzyme A-acyl carrier protein transacylasc and, 187 Ferredoxin, pyruvate formation and, 83 Ferricyanide, phosphofructokinase and, 269

Ferrous ions aspartokinase and, 512 glycerate kinase and, 508 guanidino kinases and, 472 Filter paper, pyruvate kinase and, 357358 Fish, creatine kinases of, 388 Flavin adenine dinucleotide synthesis of, 31 vitamin B1?.adenosyltransferase and, 145

Flnvobacterium thermophilum, phoaphofructokinase of, 243, 249, 263, 264 Fluorescence aspartokinase I coenzyme binding, 524 ligand binding, 527-533 creatine kinase assay and, 396

Fluoride ions creatine kinase and, 425, 426, 452 methionine adenosyltransferase and, 134

phosphate ester hydrolysis, rate constants, 211, 212 phosphofructokinase activation and, 243, 259, 261

pyruvate kinase and, 364, 382 Fluoroacetate, creatine kinase and. 427 Fluorodini trobenzene creatine kinase and, 431, 443 phosphofructokinase and, 271 Fluorophosphate, pyruvate kinase and, 364

5-Fluorouridine triphosphate, nucleoside diphosphokinase and, 323 Formate ions, creatine kinase and, 425426

Fructose, pyruvate dehydrogenase and, 566

Fructose diphosphatase, phosphofructokinase assay and, 244 Fructose diphosphate adenosine diphosphoryl glucose pyrophosphorylase, 34, 75, 76, 118 Aeromonas jormicans, 109 energy charge and, 106 enterobacteriaceae, 94-97 inhibitor interaction, 99-100 kinetics and, 97-98, 101-102 manganese ions and, 103-104 mutants and, 112-113, 115-117 spinach leaf, 86 glycerol kinase and, 502-503 phosphofructokinase and, 250-252, 259. 260, 261-263, 266, 270, 273, 275, 276

pyruvate kinase and, 356, 361, 362-363, 369, 370, 373-375, 376, 377

n-Fructose 1-phosphate, phosphofructokinase and, 244-245, 250 Fructose 6-phosphate adenosine diphosphoryl glucose pyrophosphorylase and, 75, 76, 78-79, 80-81, 86, 93, 109 phosphofructokinase and, 244, 249-252, 254, 259, 261-263, 264, 265, 266268, 270-271, 272, 273, 275, 276, 278 Fumarate, creatine kinase and, 427

SUBJECT INDEX

631

Glucose-6-phosphate dehydrogenase G adenylate kinase assay and, 301 Galactosamine, uridine diphosphoryl creatine kinase assay and, 396 glucose pyrophosphorylase and, 67 guanidino kinase away and, 465 Galactose nucleoside diphosphokinase assay and, metabolism, uridine diphosphoryl glu324, 325 cose pyrophosphorylase and. 5% 3-phosphoglycerate kinase assay and, 59, 67 341 Galactosemia, defect in, 68-69 uridine diphosphoryl glucose pyrophosP-Galactosidase, aeromonads and, 108 phorylase assay and, 52 Gel electrophoresis Glucuronides, synthesis of, 59 adenylate kinase, 282-283 Glutamate aspartokinase I and, 516, 518-519 glutamine synthetase and, 43 phosphofructokinase, 255 isomerization of, 144, 148, 151 3-phosphoglycerate kinases, 338-339, Glutamate residues 340, 342 acyl carrier protein, 168 Gentamicin, adenylylation of, 30 glycerol kinase, 495 Glucagon, histone kinase and, 579 D-GlUCarate Glutamine metabolism, glycerate kinase and, 505 carboxyl group activation and, 13 Glucitol 1,6diphosphate, adenosine diguanosine monophosphate synthesis phosphoryl glucose pyrophosphoryland, 39 ase and, 95 Glutamine synthetase, adenylylation and Glucocorticosteroid, methionine adenodeadenylylation of, 40-44 syltransferase and, 143 Glutamine synthetase: adenylyltransferGluconeogenesis. pyruvate kinase and, ase, molecular weight and subunits, 353 4u4 Glucose Glutathione adenylate kinase assay and, 301 glycerol kinase and, 496 metaholism, phosphofructokinase regumethionine adenosyltransferase and, lation and, 264 128, 134 phosphofructokinase and, 269 Glucose 1,6diphosphate, pyruvate kinase and, 374 Glyceraldehyde, glycerol kinase and, 497498 Glucose 1-phosphate ndenosinc diphosphoryl glucose pyro- Glyceraldehydephosphate dehydrogmase phosphorylase, 79, 81, 85-86, 88, 93, content of muscle, 338 107 3-phosphoglycerate kinase assay and, activator effects, 97-98 336-337, 341, 346 inhibitor effects, 100, 101-102 uridine diphosphorylglucose pyrophosmutants and, 115 phorylase assay and, 52 hydrolysis and enzyme-catalyzed D-Glycerate kinase transfer reaction, 233 catalytic properties, 507-508 phosphofructokinase and, 244, 245 rnetaholic role, 505-506 uridine diphosphoryl glucose pyrophosmethods of assay and distribution, phorylase and, 66, 67 504-505 Glucose 6-phosphate molecular properties glycolysis control and. 278 purification and state of purity, 506hydrolysis and enzyme-catalyzed trans507 fer reaction, 233 phosphofructokinase and, 275 stability, 507

632

SUBJECT INDEX

Glycerol glycerol kinase configuration and, 497, 502 methionine adenosyltransferase and, 128

pyruvate kinase and, 363 Glycerol 1,34iphosphate, adenosine diphosphoryl glucose pyrophosphorylase and, 95, 97 Glycerol kinase catalytic properties product inhibition, 501-502 substrate specificity and kinetics, 497-501

thermodynamics, 502 distribution, 48-90, 491492 metabolic role, 492-493 methods of assay, 490-491 molecular properties rhemical modification, 496497 composition, 494 purification and state of purity, 493494

size and subunit structure, 494-495 stability, 495496 regulation mammals, 504 microorganisms, 502-503 Glycerol 3-phosphate glycerol kinase and, 502, 504 phospholipid synthesis and, 165 Glycerol 3-phosphate dehydrogenase glycerol kinase assay and, 490 glycerol metabolism and, 492, 493 a-Glycerophosphate dehydrogenase, phosphofructokinase assay and, 244 Glycine, aspartokinase and, 545, 547 Glycocyamine, creatine kinase and, 403, 404,405, 406 Glycocyamine kinase amino acid composition, 469 bivalent metal ions and, 472 discovery and isolation, 460-461 distribution of, 462 essential thiol groups of, 478, 479 molecular weight, 467 stability, 468 substrate specificity, 474, 475 Glycogen accumulation of, 82, 106

Escheiichia coli mutants, 110-111, 114, 115

aeromonads and, 109 cell content, uridine diphosphoryl glucose pyrophosphorylase and, 55 phosphorylase kinase and, 559, 563 synthesis, 33 adenosine diphosphoryl glucose and, 74

uridine diphosphoryl glucose pyrophosphorylase and, 57 Glycogen synthetase, protein kinases and, 574, 575 Glycolate creatine kinase and, 427 metabolism, glycerate kinase and, 505, 506

Glycolipids, synthesis of, 59 G1ycolysis adenosine diphosphoryl glucose pyrophosphorylase and, 75 rontrol, phosphofructokinase and, 274278

hormonal control, 277-278 rate in muscle, 338 Glycosides, synthesis of, 59 Glyoxylate metabolism, glycerate kinase and, 505 Gramicidin S, biosynthesis, 11, 14-17 Growth hormone, methionine adenosyltransferase and, 143 Guanidinium chloride ncyl carrier protein and, 166 adenosine diphosphoryl glucose pyrophosphorylase and, 119 aspartokinase I and, 518. 519, 536 aspartokinase I1 and, 541 creatine kinase and, 386, 394, 403 glycerol kinase and, 494-495 /3-ketoacyl acyl carrier protein sgnthetase and, 189 phosphofructokinase and, 255, 256, 257, 270

pyruvate kinase and, 358 Guanidinoacetate, methyl transfer to, 126

Guanidino kinases, see also Phosphagen kinases catalytic properties activation by metal ions, 471-473

633

SUBJECT INDEX

function of amino acid residues, 477482 p H optimum, 476-477 substrate specificity, 473476 catalytic reaction, 459 determination of enzymic activity chemical stop methods, 464-465 continuous recording methods, 465466 isotopic methods, 465 distribution and function, 461464 equilibrium. 4&486 immunological reactions, 470-471 reaction mechanism, 482485 Guanosine diphosphatc adenosine diphosphoryl glucose pyrophosphorylase and, 79, 80 arginine kinase and, 476 creatine kinase and, 408, 409, 416, 417 nucleoside diphosphokinase and. 323 phosphofructokinase and, 252 3-pliosphoglyceratr kinase and. 348 pyruvate kinase and, 364 Guanosine diphosphoryl glurose, formation of, 68 Guanosinc 3’,5’-mononucleotide, protein kinase and, 578 Guanosine monophosphate adenosine diphosphoryl glucose pyrophosphorylasc and, 79, 80 nucleoside diphosphokinase and, 327328 synthesis of, 39-40 Guanosine triphosphate ndenylate kinase and, 299 arginine kinase and, 476 glycerol kinase and, 499 methionine adenosyltransferase and, 135, 138 nucleoside diphosphokinases and, 322 phosphofructokinase and, 246, 247, 271 3-phosphoglycerate kinase and, 348 protein kinases and, 572

H Heart nucleoside diphosphokinases of, 310, 317 phosphorylase kinase of, 564

a-Helix acyl carrier protein and, 166 arginine kinase, 481-482 aspartokinase I, 535 creatine kinase and, 393 3-phosphoglycerate kinase, 344, 346 Hrmolytic anemia, 3-phosphoglycerate kinase deficiency and, 340 Heparin phosphorylase kinase and, 564 synthesis of, 59 cis-9-Hexadecenoyl acyl carrier protein. P-ketoacyl acyl carrier protein synthctase and, 193 Hrxokinase adenylate kinase assay and, 301 creatine kinase assay and, 396 guanidino kinase assay and, 465 inhibition of, 278 nurlcoside diphosphokinase assay and, 324, 325 3-phosphoglycerate kinase assay and. 34 1 Histidine, carboxyl group activation and, 12 Hiatidine residues adenylate kinase, 291, 293, 294-295 nrgininc kinase, 479, 482, 484, 485 creatine kinase, 434, 443, 452, 453-454 nucleoside diphosphokinase. 309, 319, 329 phosphofructokinase, 272 Histone, protein kinase and, 568, 571, 574,575, 576 Histone kinases, properties, 579-580 Holo-acyl carrier protein synthetase, properties of, 174 Homoarginine, arginine kinase and. 474 Homocysteine, methionine adenosyltransferase and, 135, 137 Homoserine growth requirement for, 509 methionine adenosyltransferase and, 137 Homoserine dehydrogenase aspartokinase assay and, 512 assay of, 513 Homoserine dehydrogenase I, distribution on polypeptide chain, 536, 538

634 Homoserine dehydrogenase 11, aspartokinasc I1 and, 540 Hormones glycolysis and, 277-278 methionine adenosyltransferase and, 143 Human muscle, creatine kinase of, 389 Hybrids, creatine kinase, 386-387, 403 Hydrogen peroxide, phosphofructokinase and, 269 Hydrophobic regions, adenylate kinase,

304

SUBJECT INDEX

phoryl glucose pyrophosphorylaw and, 78 Hypophosphite ions, creatine kinase and, 426 Hypotaurocyamine kinase bivalent metal ions and, 472 distribution of, 462 immunological properties, 471 molecular weight, 467 purification of, 461 substrate specificity, 474, 475

I

Imidol adenylate Hydroxide synthesis of derivatives, 37 phosphate ester hydrolysis, rate ronargininosuccinate, 38-39 stants, 211, 212 guanosine monophosphate, 39-40 Hydroxocobinamide, vitamin B,,, adeIndoleacetate, cell wall synthesis and, nosyltransferase and, 151 59-60 P-Hydroxyacyl acyl carrier protein deInhibition constants, uridine diphoshydrase, specificity of, 193 phoryl glucose pyrophosphorylases, P-Hydroxybutyrate, creatine kinase and, 65,67 427 P-Hydroxybutyryl acyl carrier protein, Inosine diphosphate arginine kinase and, 476 fatty acid biosynthesia and, 165 creatine kinase and, 408, 409, 439 0-Hydroxydecanoyl thioester dehydrase, nucleoside diphosphokinase and, 323 unsaturated fatty acid synthesis and, 3-phosphoglycerate kinase and, 348 191 pyruvate kinase and, 364 Hy droxylamine Inosine diphosphoryl glucose, synthesis acetyl acyl carrier protein and, 186 of, 32 acyl hydroxamate formation from, 8 Inosine trjphosphate aspartokinase assay and, 512 adenylate kinase and, 299, 300 dibenzylphosphoenol pyruvate hyarginine kinase and, 476 drolyis and, 223 glycerol kinase and, 499 guanidino kinases and, 482 methionine adenosyltransferase and, malonyl coenzyme A-acyl carrier pro135 tein transacylase and, 181, 184-185 nucleoside diphosphokinases and. 322 monobenzyl phosphoenol pyruvate hyphosphofructokinase and, 246, 247-248, drolysis and, 221-222 260, 267, 271 phosphate ester hydrolysis, rate con3-phosphoglycerate kinase and, 348 stants, 212 protein kinases and, 572 pyruvate kinase and, 364, 382 Insect p-Hydroxymercuribenzoate flight muscle, glycerol kinase in, 489, adenylate kinase and, 293 493 glycerate kinase and, 507 Insulin phosphofructokinase and, 254 adenylate kinase and, 287 pyruvate kinase and, 380 glycolysis control and, 277 p-Hydroxymercuriphenylsulfona te, pyruvate dehydrogenase and, 566 glycerol kinase and, 497 uridine diphosphoryl glucose pyroHydroxypyruvate. adenosine diphosphosphorylase and, 60

SUBJECT INDEX

Intestinal mucosa, nucleoside diphosphokinase of, 310 Intestine, glycerol kinase in, 489 Iodide ions, creatine kinase and, 426, 446 Iodine arginine kinase and, 481 methionine adenosyltransferase and, 13k135 phosphofructokinnse and, 269 Iodoace tamidc acetyl coenzyme A-acyl carrier protein transacylase and, 187, 188 arginine kinase and, 479 creatine kinase and, 304, 425, 426, 427, 431432, 438, 442-443, 444445,448, 449,450,453 glycerol kinase and, 496-497 P-ketoacyl acyl carrier protein synthetase and, 194, 197, 198 malonyl coenzyme A-acyl carrier protein transacylase and, 183 phosphofructokinase and, 270-271 pyruvate kinase and, 361 Iodoacetate adenylate kinase and, 294 aspartokinase I and, 521, 536 creatine kinase and, 393, 399, 431, 433, 436, 437, 438, 439, 442, 443, 445446, 449-450 glycerate kinase and, 507 glycerol kinase and, 497 malonyl coenzyme A-acyl carricr protein transacylase and, 179 phosphofructokinase and, 270 Ionic strength, protein kinases and, 576 DL-Isoaminobutyrate, creatine kinase and, 427 Isocitrate, phosphofructokinase and, 265, 266 Isoelectric focusing, nucleoside diphosphokinases, 313-314 Isoelectric points nucleoside diphosphokinases, 316-318 protein kinases, 578 Isoenzymes adenylate kinase, 282-284 arginine kinase, 460 nucleoside diphosphokinase, 309, 313314

635 phosphofructokinase, 257-258 Isoleucine aspartokinase and, 514, 544, 545, 547, 552 biosynthesis, 511 J

Jejunal mucosa, phosphofructokinase of, 262, 275-276

K Kanamycin, adenylylation of, 30 a-Ketoacid dehydrogenase, lipoate activating enzyme and, 17, 18 P-Ketoacyl acyl carrier protein reductase acyl carrier protein peptide and, 172 specificity of, 193 p-Ketoacyl acyl carrier protein synthetase srctyl acyl carrier protein and, 186 ncyl carrier protein peptide and, 172 catalytic properties assay, 190 mechanism, 194-199 p H optimum, substrate specificity and kinetics, 190-194 historical background, distribution and metabolic significance, 188-189 molecular properties, 189-190 a-Ketobutyrate, adenosine diphosphoryl glucose pyrophosphorylase and, 78, 81 2-Keto3-deoxy-phosphogluconate, adenosine diphosphoryl glucose pyrophosphorylase and, 78 a-Ke toglutarate glutamine synthetase and, 43 phosphofructokinase and, 266 a-Ketoglutarate-succinate thiokinase, nucleoside diphosphokinase assay and, 324 Kidney glycerol kinase in, 489 nucleoside diphosphokinases of, 310, 316, 318 phosphofructokinase of, 262, 266 Kinase activating factor, phosphorylase kinase and, 562

636

SUBJECT INDEX

Kinetic constants adenosine diphosphoryl glucose pyrophosphorylase, 78-117 adenylate kinase, 299 aspartokinase I, 519-520 aspartokinase 11, 541 nspartokinase 111, 543 creatine kinase, 405, 407, 408, 411, 415, 419, 421

glycerate kinase, 507 glycerol kinase, 497, 499 nucleoside diphosphokinases, 322323 phosphorylase kinase, 563-564 protein kinases, 572 uridine diphosphoryl glucose pyrophosphorylases, 66 Kinetics pyruvate kinase inhibitors, 375-379 substrates and activators, 372-375

1 Lactate, creatine kinase and, 427 Lactate dehydrogenase adenylate kinase assay and, 301 aspartokinase assay and, 512 creatine kinase assay and, 396 glycerate kinase assay and, 505 glycerol kinase assay and, 490 guanidino kinase assay and, 464 kinetics, 269 nucleoside diphosphokinase assay a.nd, 321. 324

phosphofructokinase assay and, 244 pyruvate kinase assay and, 371 Loctobacillzis cnsei. phosphofructokinase of, 249,263, 264 Loctobacillzis plantarum, phosphofructokinase of, 250,263, 264 Lactose, synthesis of, 59 Laurate, pyruvate kinase and, 377 Leaves, adenosine diphosphoryl glucose pyrophosphor~lasesof, 89-90 Lens adenylate kinase, physical properties, 296

phosphofructokinase of, 249, 262, 264 Leucine, nspartokinase I11 and, 544, 547 Lipoate, attachment. to enzymes, 6, 13

Lipoate activating enzyme, adenylyl group transfer and, 17-18 Lithium chloride, glycerol kinase and, 501

Liver ncy1 carrier protein, 4’-phosphopantetheine peptide of, 170 adenylate kinase, 280, 282, 287 amino acid composition, 292 physical properties, 296 purification, 289 substrate specificity, 299 glycerol kinase in, 489 nucleoside diphosphokinases intracellular distribution, 310, 314 physical properties, 317-318 phosphorylated enzyme, 319, 320 substrate specificity, 322-323 phosphofructokinase, properties, 242, 258, 262, 266

phosphorylasc kinase of, 565 protein kinase in, 567 pyruvate kinase of, 359 regeneration, histone kinase and, 579 uridine diphosphoryl glucose pyrophosphorylase amount present, 55 molecular weight, 62-63 purification from, 53-54 Liver fluke. phosphofructokinase of, 260261

Lombricine kinase amino acid composition, 469 bivalent metal ions and, 472 discovery and isolation, 461 distribution of, 462 cssential thiol groups of. 478, 479 immunological properties, 470-471 molecular weight, 467 nucleotide specificity, 476 peptide, reactive thiol group and. 470 reaction merhanism, 483 stability, 468 substrate specificity, 475, 476 Luciferase, inhibition of, 20 Luriferin activation of, 12 adenylylntion of, 19-20 T,ung, nuclroside diphosphokinasr of. 310. 317

637

SUBJECT INDEX

Lysine aspartokinase and, 512, 513, 544, 545546, 547, 549, 550. 551. 552 hiosynthesis of, 511 Lysine residues acyl carrier protein, 172-173 adenosine diphosphoryl glucose pyrophosphorylase and, 97 adenylate kinase, 294. 295 arginine kinasr, 481 creatine kinase, 432-431, 452, 454 deoxyrihonucleic acid ligase, ndenylylation of, 46-18 Nf-group, carboxyl activation and, 13, 17-20

nudeosidr diphosphokinases and, 319. 329

phosphofructokinase, 255, 256, 257 pyruvate kinase. 360, 382 Lysophosphatidic acid, biosynt hesis of, 165

M Magncsium ions adenosine diphosphate complexes, 235 ndenosine diphosphoryl glucose pyrophosphorylase and, 79, 85-86, 87, 93, 100, 101, 115

adenylate kinasc and, 298, 302. 303-304 aspartokinase and, 512, 548 creatine kinase and, 396, 407, 409-414, 417, 428, 429430, 439, 449. 450

glutamine synthetase and, 42 plycerate kinase and. 508 glyrerol kinasr and, 498, 500-501 guanidino kinases and, 471. 472, 486 metabolic control and, 286 incthionine adenosyltransfernse and, 134, 135

nucleoside diphosphokinases and, 329330

]-)liospliofructokinnsc and, 241, 243. 247-248, 261

3-phosphoglycerate

kinase and, 336,

346-348. 349

phosphorylase kinase and, 563 phosphoryl transfer reactions and. 229230

protein kinases and, 572, 573, 576, 578

pyruvate kinase and, 360,361, 368-370, 371, 373, 375-376,380

standard free energy of hydrolysis of adenosine triphosphate and, 2-3 tripolyphosphatase and, 134 Maize starchdeficient mutants, 94 starch synthesis in, 58 tissues, adenosine diphosphorylglucose pyrophosphorylasee of, 93-94 Malute, phosphofructokinase and, 265, 266

Maleic anhydride, phosphofructokinase and, 256, 272-273 Malonate, creatine kinase and, 427 Malonyl acyl carrier protein, decarboxylation, 6-ketoacyl acyl carrier protein synthetase and, 198-199 Malonyl coenzyme A, acyl carrier protein and, 156-157 Malonyl coenzyme A-acyl carrier protein trnnsacylase catnlytic properties assays, 179-180 mechanism, 180-185 pH optimum, substrate specificity and kinetics, 180 fatty acid hiosynthesis and, 165 historical background, distribution and metabolic significance. 176-178 molecular properties, 178-179 Mammals acyl carrier protein in, 160 me thionine adenosyltransfernse age and, 142 dietary factors, 142-143 hormonal effects, 143 nucleosidr diphosphokinases of, 310311, 316318

Mammary gland glycerol kinase in, 489 protein kinase in, 567 uridine diphosphorylglucose pyrophosphorylase of, 63 Manganese ions adenosine diphosphate complexes, 235 adenosine diphosphorylglucose pyrophosphorylase and, 102-104 adenylate kinase and, 295, 298 aspartokinase and, 512

SUBJECT INDEX

creatine kinase and, 407, 409412, 416, 417, 426, 428, 429-430, 437, 439440 glycerate kinase and, 508 glycerol kinase and, 501 guanidino kinases and, 472 methioninc ndenosyltransfernse and, 134 nucleoside diphosphokinases and, 329 phosphofructokinase and, 247 3-phosphoglycerate kinase and, 349 protein kinnses and, 573, 578 pyruvate kinase and, 362, 367-369, 370. 373, 377, 381 vitamin BIZsadenosyltransferase and, 145, 151 Membrane proteins, protein kinase and, 574, 575 Mcrcaptoethanol creatine kinase and, 398, 401 glycerol kinase and, 496 p-ketoacyl acyl carrier protein synthetase and, 190, 194 methionine adenosyltransferase and. 128 phosphofructokinase and, 257 vitamin BIZ. adenosyltransferase and, 149, 150 2-Mercaptoethylamine, acyl carrier protein and, 167 6-Mercapto-9-p-~-ribofuranosylpurine 5’triphosphate aspartokinase I and, 524-525 phosphofructokinase and, 245 p-Merruribenzoate nspartokinase I and, 525 phosphofructokinase and, 270, 271 Mercuric ions 3-phosphoglycerate kinase crystals and, 344 phosphoenolpyruvate hydrolysis and, 380-381 Metal ions, catalysis of phosphoryl transfer, 227-231, 234-235 Mctaphosphate mechanism enzymic phosphoryl transfer and, 233235 hydrolysis of acyclic phosphate esters. 202-206

Methioninc adenosine triphosphate :sulfate adenylyltransferase synthesis and, 37 analogs, methionine adenosyltransferase and, 135-137 aspartokinase and, 513, 515, 545, 547 biosynthesis of, 511 growth requirement for, 509-510 methionine adenosyltransferase, 138 regulation of, 141 Methionine adenosyltransferase catalytic properties activators and pH effects, 133-135 assay, 129-130 energetics, 139-14 1 kinetics, 137-139 reversibility, partial reactions and mechanism, 130-133 substrate specificity and inhibition by analogs, 135-137 net reaction, 125-127 physical properties, 128-129 purification, 127 regulation and genetics mammals, 142-143 microorganisms, 141-142 significance and distribution, 123-125 Methionine residue acyl carrier protein, 173 ndenylate kinase, 294 3-phospl~oglycerate kinase, 343 Methionine sulfone, methionine adenosyltransferase and, 136 Methionine sulfoxide, methionine adenosyltransferase and, 136 o,L-N-Methyl-N-amidinoalanine,creatine kinase and, 405 N-Methyl-N-amidino-p-alanine,creatine kinase and, 405, 406 N-Methyl-N-amidinoaminomethylphosphinic acid, creatine kinase and, 405, 406 N-Me t hyl-N-amidinoaminomethylphosphonic acid, creatine kinase and, 405, 406 N-Methylamidino-N-me thylglycine. creatine kinase and, 405, 406 a-Methyl aspartate, pyrophosphate exchange and, 38

SUBJECT INDEX

P-Methylaspartate, formation of, 144 Methyl 2-carboxyphenyl phosphatr, hydrolysis of, 220 Methylene blue adenylate kinase and, 295 phosphofructokinase and, 272 Methyl ethylene phosphate, hydrolysis of, 217 Methyl hydrogen N-methyl-N-amidinoaminomethylphosphate, crcatine kinase and. 405, 406 N-Mcthylhydroxylamine, pyruvate kinase and, 365 Methylmalonatc, excretion of, 145 Mcthylmalonyl coenzyme A, isomerization of, 144 Methylmalonyl coenzyme A-pyruvate carboxytransferase, biotin activating rnzyme and, 18 Methyl phosphate hydrolysis, metal ions and, 230-231 Micrncoccus glutnmicua, aspnrtokinaw of, 552 Microorganisms glpccrol kinase in, 491-492 mrthionine adcnosyltransferase, regulation and genetics, 141-142 nucleosidr diphosphokinases of, 312 Microsomes protein kinase in, 567 iiridinr diphosphoryl glucose pyrophosphorylase in, 56 Mitochondria adenylate kinase, 280, 281-282, 287-288 physical properties, 296. 297 purification of, 289 substrate specificity, 299 creatine kinase in, 385 glycerol kinas? in, 491 nucleosidc diphosphokinases of, 310, 311, 312, 314, 316, 322-323, 3 2 9 330, 331 protein kinaae in, 567, 580 serinc-bound phosphate in, 566 Molecular weight argininr kinases. 466. 467 aspartokinase I. 517-518 nspartokinase 11, 541 aspartokinase 111, 543 bacillus aspartokinases, 548, 549, 550

639 creatine kinase, 395 glycerol kinase, 494-495 nucleoside diphosphokinases, 316, 318 3-phosphoglycerate kinase, 342 pliospliorylase kinase, 559 protein kinases, 577-578 pyruvate kinase, 358,359 uridine diphosphoryl glucose pyrophosphorylases, 62-63 Mollusca, guanidino kinase of, 462 Monkey muscle, creatine kinase of, 389 Monobrnzyl phosphoenolpyruvate, hydrolysis of, 221-222 Muscle adenylate kinase, 280, 287 amino acid composition, 292 mechanism, 303 physical properties, 296 purification of,289 substrate specificity, 299 rrratine kinase, 386-387 amino acid composition, 389 glycerol kinase in, 489 heart, phosphofructokinase of, 242, 249, 259, 260, 262, 265, 266, 269, 270, 272 nucleoside diphosphokinases of, 310 3-phosphoglycerate kinase, 337-338. 339 amino acid composition, 343 kinetic ronstants, 347 molecular weight, 342 X-ray crystallography, 345-346 pyruvate kinase purification of, 355, 356 structure, 358-359 skeletal phosphofructokinase of. 242, 246, 247, 249, 253-256, 257-258, 259, 262, 265, 269, 270, 272 phosphorylase kinase of, 557-564 Muscular dystrophy adenylate kinase and, 284 ereatine kinase and, 386-387 Mutants aspartokinase I and, 515-516. 536-537 nspartokinase I1 and, 540 glycerol kinase, 503 Mycobncterium phlei acyl carrier proteins

640

SUBJECT INDEX

amino acid composition, 159 physical properties, 166 primers for, 160 Myocardial infarction, adenylate kinasc and, 284 Myokinasr, see Adenylate kinase Myristate, pyruvnte kinase and, 377

N Negmine, creatine kinase and, 405 Neuvospora crassa methionine adenosyltransferase of, 141142

phosphofructokinase of, 263 Neurotubule protein, protein kinase and, 574 Nickel ions glycerol kinase and, 601 3-phosphoglycerate kinase and, 349 pyruvate kinase and, 369 Nicotinamide adenine dinucleotide adenylate kinase assay and, 301 deoxyribonucleic acid ligase and, 4547

glycerate kinase assay and, 504, 505 P-ketoacyl acyl carrier protein synthetase assay and, 190 malonyl coenzyme A-acyl carrier protein, transacylase assay and, 179 nucleotide diphosphokinase assay and, 321, 324

phosphofructokinase and, 266, 276 assay, 243-244 3-phosphoglycerate kinase assay and, 336,337,341,346

pyruvate kinase assay and, 371 ribonucleic acid modification and, 4849

uridine diphosphoryl glucose pyrophosphorylase assay and, 52, 53 Nicotinamide adenine dinucleotide phosphate ndenosine diphosphoryl glucose pyrophosphorylase and, 75, 76. 94-95, 96, 97, 109 energy charge and, 106 kinetic effects, 97-98, 100 manganese ions and, 103 mutants and, 112, 115

adenosine diphosphate glucose pyrophosphorylase and, 34 adenylate kinase assay and, 301 aspnrtokinase assay and, 512 aspnrtokinase I and, 518, 519, 523-524, 532, 534, 535

nspartokinase I1 and, 541 fatty acid biosynthesis and, 165 uridine diphosphoryl glucose pyrophosphorylase assay and, 52 Nicotinate ribonucleotide, deamidonicotinamidc adenine dinucleotide synthesis and, 31-32 Ninhydrin, creatine kinase assay and, 396

Nitrate ions, creatine kinase and, 405, 406, 411, 423, 425-426, 435, 436, 438, 452 Nitritc ions, creatine kinase and, 425426

p-Nitrophenyl acetate, creatine kinase and, 432, 433,438 2-Nitrophenylsulfenyl chloride, aspartokinase I and, 521 Norleucine, methionine adenosyltransferase and, 136 Norvaline nspnrtokinase and, 547 methioninc adenosyltransferase and, 136

Nuclear magnetic resonance phosphoranes, 215 pyruvate kinase, 366-368, 373 Nuclei, adenylate kinase in, 281, 282 Nucleophilic reactions. acyclic phosphorus and, 208-214 Nucleoside diphosphokinases catalytic properties assay, 321, 324-325 conformational changes, 331 kinetics and catalytic mechanism. 326-329

metal requirements, 329-330 reaction catalyzed, 320 specificity, 320-321, 322-323 sulfhydryl groups, 330-331 distribution, 309-313 functions in the cell, 331-333 historical development, 307-309 mechanism, 308-309

64 1

SUBJECT INDEX

molecular properties isozyme occurrence, 313-314 phosphorylated enzyme, 315-320 physical properties, 315-318 purification procedure, 314, 318-320 phosphoryl transfer and, 233 rextion catalyzed, 307-308 Suclcosidr t riphosphate :deoxyribonucleic acid nuclrotidyl transferasc, function of, 26 Nucleotide diester analog. phosphoryl transfer and, 233

0 Octanoate, pyruvate kinase and, 377 Oleate, pyruvate kinase and, 377 Oligoribonucleotides, polynucleotide phosphorylase and, 23 Ophelinc kinasc bivalent metal ions and, 472 distribution of, 462 immunological properties, 470-471 purification, 461 substrate specificity, 475, 476 Optical rotatory dispersion, aspartokinase I, 535-536 Orchiectomy, mcthionine adenosyltransfcrase and, 143

ox

hrain-type creatine kinases of, 390 muscle, creatine kinase of, 389 Oxalate, activation of, 6, 12 Oxygen transfer, adenylyl group transfer and, 8-9 N (1-Oxyl-2,2,5,5-tetramethyl-3-pyrrolidinyl) iodoacetamidr, creatine kinas? and, 440441,450 Oxy phosphoranc p hcnan threncquinone-triisopropy 1 phosphite adduct, structure of. 214

P Palmitate biosynthesis of, 191-192 pyruvate kinase and, 377 Palmitoleate, biosynthesis of, 191-192 Palmityl acyl carrier protein, biosynthesis of, 165

Palmityl thioesterase, fatty acid biosynthesis and, 165 Pancreas regeneration, histone kinase and, 579 Pante tlieine acrtyl coenzyme A-acyl carrier protein transacylase and, 187 adenylylation of, 31, 32 gramicidin synthesis and, 15-16 malonyl coenzyme A-acyl carrier protcin transacylase and, 180 tyrocidinc synthesis and, 17 Pantoate, activation of, 12 Pantothenate, formation of, 6 Pasteur effect, phosphofructokinase and, 274276 Pea seed nucleoside diphosphokinase of, 311.314, 316, 319 phosphofructokinase of, 262, 264 3-phosphoglycerate kinase, kinetic constants, 347 Penicillium chrysogenum, adenosine triphosphate :sulfate adenylyltransferase of, 3s37 Prntacovalency, pseudorotation and, 214219 1.5-Pcntanediol diphosphate, adenosine diphosphorylglucose pyropliosphorplasc. and, 95 Pepsin acyl carrier protein peptides, 167 malonyl coenzyme A-acyl carrier protein transacylase peptides and, 177-178 Peptidasc. methionine adenosyltransferase preparations and, 127 Peptide nucleic acid-independent synthesis, adenylyl group transfer and, 11, 14-17 Peptide sequences, creatine kinase, 392 Performic acid fatty acid synthetase and, 177, 178, 186 malonyl coenzyme A-acyl carrier protein transacylase and, 181 Prriplasmic space, 5’-nucleotidase in, 161 PH adenosine diphosphoryl glucose pyro-

642

SUBJECT INDEX

phosphorylases and, 81, 84, 86, 8788

aspartokinases and, 549, 550 creatine kinase and, 397, 398, 399, 402, 412413, 418, 419, 422, 428, 429, 442-443 glycerate kinase and, 508 glycerol kinase and, 495496, 498, 499, 501, 503 guanidino kinases and, 476-477

methionine adenosyltransferase and, 133-134

nucleoside diphosphokinase and, 328329

phosphofructokinase and, 247, 248, 259, 265

phosphorylase kinase and, 560 protein kinases and, 576 standard free energy of hydrolysis of adenosine triphosphate, 2-3 tripolyphosphatase and, 134 uridine dipliosphoryl glucose pyrophosphorylases and, 62 Phenylacetate, activation of, 6, 12 Pheny lalanine aspartokinase nnd, 547 aspartokinase I11 and, 544, 545 gramicidin synthesis and, 14-15 pyruvate kinase and, 372, 373, 376-377 tyrocidine biosynthesis and, 17 Phenylethylbiguanide, pyruvate kinase and, 377 Phenyl N-(glycy1)-phosphoramidate, intramolecular exocyclic displacement reaction of, 229-230 Phenylhydrazine, glycerate kinase assay and, 505 Phenylisothiocyanate, creatine kinase and, 393 Phenylmethanesulfonylfluoride, malonyl coenzyme A-acyl carrier protein transacylase and, 183 Phenyl phosphate hydrolysis, rate constant, 206 y-Phenylpropyl di(tri)phosphate hydrolysis, rates, 204 y-Phenylpropyl triphosphate, pyrophosphate formation from, 229 Phenylthiophosphate monoanion hydrolysis, rate, 206

Phormia regina muscle, phosphorylase kinase of, 565 Phosphagen kinases, see also Guanidino kinases amino acid composition, 391 historical background, 457459 Phosphate acyclic di- and triesters, hydrolysis, 207-208

adenosine diphosphoryl glucose pyrophosphorylase and, 74, 76, 77, 79, 80, 81, 86,88-90, 91-92, 93, 99, 100, 101, 103

creatine kinase and, 407, 423, 426 glycerol kinase and, 504 inorganic, adenosine diphosphate glucose pyrophosphorylases and, 34 methionine adenosyltransferase and, 138

mitochondria1 adenylate kinase and, 287-288

phosphofructokinase and, 262-263, 264, 265, 266, 275

uridine diphosphoryl glucose pyrophosphorylase and, 66, 67 Phosphate diester monoanions, amines and, 209 Phosphate esters hydrolysis of acyclic esters di- and triesters, 207-208 metaphosphate mechanism for monoesters, 202-206 Phosphate monoester dianions, amines and, 209 Phosphate triesters, oxyanions and, 209 Phosphoarginine binding by arginine kinase, 485 creatine kinase and, 403 isolation of, 458 Phosphocreatine nctive form of, 414 phosphofructokinase and, 266, 275 phosphoryl transfer and, 233 Phosphodiesterase(s), phosphorothioates and, 213 Phosphoenol-3-bromopyruvate, pyruvate kinase and, 365 Phosphoenol-3-fluoropyruvate, pyruvate kinase and, 365

SUBJECT INDEX

Pliosphoenol-a-ketobutyrate, pyruvate kinase and, 365 Phosphoenol-a-ketocaproate, pyruvate kinase and, 365 Phosphoenol-u-ketovalerate, pyruvate kinase and, 365 Phosphoenolpyruvate adenosine diphosphoryl glucose pyrophosphorylase and, 34, 76, 79, 8081, 86, 95, 97, 107 adenylate kinase assay and, 301 nspartokinase assay and, 520 glycerate kinase assay and, 504 glycerol kinase assay and, 490 hydrolysis of, 221, 222 nucleoside diphosphokinase assay and, 324 phosphofructokinase and, 266 phosphoryl transfer and, 233 pyruvate kinase and, 360, 361, 363, 365, 367, 370, 372, 373, 37S376, 379380,382 Phosphoenolpyruvate carboxykinase, gluconeogenesis and, 354 Phosphoenolpyruvate synthase, reaction catalyzed, 83 Phosphofructokinase assay of activity, 243-244 catalytic properties cation requirement, 247-248 isotope exchange studies, 252 kinetic studies of mechanism, 248252 phosphoryl acceptor specificity, 244245 phosphoryl donor specificity, 245-247 glycolysis control hormonal control, 277-278 Pasteur effect, 274-276 pyridine nucleotide oscillations, 276 levels, pyruvate kinase and, 374 properties of, 242 purification, 241-243 reaction catalyzed, 240 regulatory properties, 261-269 role of specific groups in activity histidine, 272 other functional groups, 272-274 thiols, 269-272 structural properties

643 chemical modification, 260-261 Clostridium pasteuriunum, 256 Escherichia coli, 256-257 isoenzymes, 257-258 molecular weight, 253-254 rabbit erythrocyte, 257 rabbit muscle, 2562.56 reversible inactivation by dilution, 259-260 Phosphoglucomutase phosphofructokinase and, 245 phosphoryl transfer and, 233 uridine diphosphoryl glucose pyrophosphorylase assay and, 52 6-Phosphogluconate, phosphofructokinase and, 266 3-Phosphoglyceraldehyde, adenosine diphosphoryl glucose pyrophosphorylase and, 34, 76, 95, 97 2-Phosphogly cerate adenosine diphosphoryl glucose pyrophosphorylase and, 34, 95, 97 phosphofructokinase and, 266 3-Phosphoglycerate adenosine diphosphoryl glucose pyrophosphorylase and, 34, 76, 77, 8687, 88-90, 93, 96, 118 physiological significance of activation, 91-92 phosphofructokinase and, 266 3-Phosphoglycerate kinase biological occurrence, 337-338 historical background, 335-336 molecular properties molecular weight, 342 primary structure, 342-343 secondary structure, 344 tertiary structure, 344-346 nucleoside diphosphokinase assay and, 324 purification procedures, 340-341 reaction catalyzed, 336-337 reaction kinetics back and forward reactions, 346-348 metal ion specificity, 349 nucleotide specificity, 348 postulated mechanism, 349-351 species variation and genetics, 338-340 uridine diphosphoryl glucose pyrophosphorylase assay and, 52

644

SUBJECT INDEX

Phosphoglycocyamine, occurrence of, 459 Phosphoglycolate, pyruvate kinase and, 367-368

Phosphohistidine, phosphoryl transfer and, 233 Phosphohypotaurocyamine, occurrence of, 459 Phospholactate, pyruvate kinase and, 367-368, 382

Phospholipid, biosynthesis of, 165 Phospholombricine, occurrence of, 459 Phosphonium esters, inversion of configuration, 213 Phosphoopheline, occurrence of, 459 4’-Phosphopante theine acyl carrier protein and, 158, 160. 165, 167

fatty acid synthesis and, 169 turnover of, 175-176 Phosphoramidates amines and, 209 hydrolysis of, 203-204, 206 pyridines and, 209 Phosphoranes geometry, 214 pseudorotation of, 215 5-Phosphoribosyl-l-pyrophosphate, adenosine diphosphoryl glucose pyrophosphorylase and, 95 Phosphorothioates alkaline phosphatase and, 213 cyclic 3’,5‘-nucleotide phosphorodiesterase and, 213 phosphodiesterases and, 213 ribonuclease and. 213 Phosphorylase site phosphorylated, amino arid sequence, 557 Phosphorylase kinase phosphorylation of, 558 protein kinases and, 574575 skeletal muscle kinetic properties, 560-564 purification and molecular properties, 559 reaction catalyzed and metabolic significance, 557-558 specificity, 558-559 N-Phosphoryl creatine monoanion hydrolysis, rate of, 204

0-Phosphorylhydroxylamine, formation of, 364 Phosphorylimidazolium ion, phosphoryl transfer to pyridine-2-carbaldoxime, 228

N-Phosphoryl imidazolium monanion, reactivity of, 204 N-Phosphoryl-N’-methylimidazolium ion, hydrolysis, rate of, 204, 206 Phosphoryl transfer adenosine triphosphate and, 2-3 catalysis intramolecular, 219-227 metal ion, 227-231 enzymic, 225, 227 enzymic catalytic mechanisms, 232-233 bimolecular or associative, 235-238 metaphosphate, 233-235 Phosphorus acyclic, nucleophilir reactions at, 2 0 s 214

Phospho taurocyamine creatine kinase and, 403 occurrence of, 458-459 Phosvitin, protein kinase and, 574, 575 Phosvitin kinases, properties, 580-581 Photosynthesis, phosphate levels and, 91 pH-stat adenylate kinase assay and, 301 creatine kinase assay and, 396, 401, 409-410

guanidino kinase assay and, 466 pyruvate kinase assay and, 371 Physnrzim polycephalum ndenylate kinase of, 281 protein kinase of, 578 Pituitary gland, protein kinasc in, 567 Plants adenosine diphosphoryl glucose pyrophosphorylase of, 76, 77, 86-90 energy charge in, 285 nucleoside diphosphokinases of, 311312

starch synthesis, regulation of, 34 uridine diphosphorylglucose pyrophosphorylase in, 56 Plasma membrane, aryl carrier protein and, 164 Platelets, phosphofructokinase of. 258

SUBJECT INDEX

Platyhelminthes, guanidino kinases of, 462

&Pleated sheet aspartokinase I and, 535-536 creatine kinase and, 394 Polyadenylate polymerase. adenylyl transfer by, 22, 25-26 Poly-P-hydroxybutyrate, accumulation of, 82 Polynucleotide adenylyltransferases, adenosinr triphosphate specific, 2426

Polynucleotide phosphorylase adenylyl group transfer and, 22-24 function of, 24 protein kinase and, 574 Potassium chloride aspartokinase I and, 517, 518, 523, 524, 527, 532, 533-534 glycerol kinase and. 501, 503 protein kinases and, 576 Potassium ions aspartokinases and, 546, 549 glycerate kinase and, 508 methionine adenosyltransferase and, 134

phosphofructokinase and, 248 3-phosphoglycerate kinase and, 346 phosvitin kinase and, 581 pyruvate kinase and, 360, 361, 366-368, 370 tripolyphosphatase and, 134 vitamin BIZ, adenosyltransferase and, 145, 151

Proline. tyrocidine synthesis and, 17 Proline residues, guanidino kinases and, 469-470

Pronase, malonyl coenzymr A-acyl carrier protein transacylase peptide and, 182

Propionate, creatine kinasr and, 427 Propionibacterium shermaiaii, vitamin BIZ. adenosyltransferase of, 146-147, 154 N-Propyl-N-amidinoglycine,creatine kinase and, 405, 406 Propylphostonic acid methylester. hydrolysis of, 216-217 Protamine phosphorylation of, 566

protein kinases and, 571, 574, 575, 576, 580

Protease, phosphorylase kinase and, 562 Protein (s) functional groups, adenylylation of, 40-49

heat stable, protein kinases and, 577, 579

protein kinase substrate specificity and, 573-576

regulatory, glutamine synthetase and, 43

Protein kinases cyclic adenosine monophosphatedependent mechanism of action of nucleotide, 568-572

nomenclature, 566 properties, 572-578 purification, 568 tissue and subcellular distribution. 567-568

historical background, 555-557 substrate specific phosphorylase kinase, 557-565 pyruvate dehydrogenase kinase, 565566

nonclassified, 578-581 Proton transfer, phosphate ester hydrolysis and, 202-203 Pseudomonads, aspartokinases of, 551552

Pseudorotation pentacovalency and, 214-219 preference rules, 215-216 Puccinin striijormis, starch accumulation and, 92 Pyridine phosphate ester hydrolysis, rate constants, 211, 212 Pyridine-2-carbaldoxime, phosphoryl transfer to, 228 Pyridoxal phosphate adenosine diphosphoryl glucose pyrophosphorylases, 34, 75, 76, 94-95, 96, 97, 109

energy charge and, 106 inhibitor effects, 100, 102 kinetic effects, 97, 98

646

SUBJECT INDEX

manganese ions and, 103 mutants and, 112, 115, 117 phosphofructokinase and, 273 pyruvate kinase and, 355, 360, 370 4-Pyridoxic acid 5-phosphate, adenosine diphosphoryl glucose pyrophosphorylase and, 94-95, 97 Pyrophosphatase methionine adenosyltransferase and, 128, 129

acetyl coenzyme A formation and, 3, 4

adenosine diphosphoryl glucose pyrophosphorylase and, 79, 87, 88 activator effects, 97-98 argininosuccinate synthesis and, 38-39 creatine kinase and, 407 exchange with adenosine triphosphate,

guanidino kinase assay and, 464 molecular properties chemical modification, 360-361 composition, 358 conformational change, 361-364 purification, 356358 structure, 358-359 nucleoside diphosphokinasc assay and, 321, 324

phosphofriictokinase assay and, 244 phosphoryl transfer and, 233 pyridine nucleotidc oscillations rind, 276

Pyruvate oxime hydroxamate, formation of, 221-222, 2!23 Pyruvate phosphate dikinnsc. pyruvate kinase nnd, 354

8

methionine

adenosyltransferase

Q

and,

125, 131, 135, 138, 153

Quinones, phosphofructokinase and, 269

uridine diphosphoryl glucose pyrophosphorylase and, 65-67, 70 vitamin B13, adenosyltransferase and, 146, 152

Py ruvat e adenosine diphosphoryl glucose pyrophosphorylase and, 34, 75, 76, 7879, 80, 81-83, 84-86, 118

formation from acetate, 82-83 pyruvate kinase and, 364, 370, 380 Pyruvate carboxylase, gluconeogenesis and, 354 Pyruvate dehydrogenase kinase, properties, 565-566 Pyruvate kinase adenylate kinase assay and, 301 aspartokinase assay and, 512, 520 catalytic properties assay, 371 control, 378-379 kinetics, 372-377 mechanism, 379-382 stoichiometry and specificity, 364371

thermodynamics, 371-372 rreatine kinase assay and, 396 general considerations, 353-355 glyccratr kinase assay and, 504 glycerol kinase assay and, 490

R Rabbit brain, creatine kinase of, 390 muscle, creatine kinase of, 388, 391 Red bone marrow. nucleoside diphosphokinase of, 310 Rhamnose, synthesis of, 59 Rhodopseudomonas cnpsulntn adenosine diphosphoryl glucose pyrophosphoryl of, 76, 7 8 8 1 aspartokinase of, 544-545, 546 glycogen synthesis, regulation of, 34 Rhodopseudomonas spheroides, aspartokinase of, 552-553 Rhodospirillicm rubrum adenosine diphosphoryl glucose pyrophosphorylase, 75, 76, 81-83 physical properties, 118 reaction mechanism, 86 temperature effects, 83-86 aspartokinase of, 545 glycogen synthesis, regulation of, 34 Hhodospirillum tenue, aspartokinase of, 545

Ribonuclease phosphorothioates and, 213 phosphoryl transfer and, 233, 236

647

SUBJECT INDEX

Ribonucleic acid polyadenylate polymerase and, 25 synthesis, adenylyl group transfer and, 20-26 Ribonucleic acid polymerase activity, determination of, 54-55 adenylyl group transfer and, 21-23 covalent modification of, 48-49 protein kinase and, 574 Ribose 5-phosphate adenosine diphosphoryl glucose pyrophosphorylase and, 34, 76, 78 Ribosome proteins, protein kinase and, 574. 575 Ribulose diphosphate carboxylase, 3phosphoglycerate kinase and, 337 Rice leaf, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Rubidium ions, methionine adenosyltransferase and, 134

s Saccha~omycescerevisiae aspartokinase of, 553 mcthionine adenosyl transferase of, 141 Salicylate, creatinc kinasr and, 427 Salicyl phosphate triesters. hydrolysis of, 224 Salicyl phosphntr dianion, hydrolysis of, 220-221 Salicyl thiophosphate, hydrolysis of, 221 Salmonelln typhirnu&m ndenosine diphosphoryl glucose pyrophosphorylase of, 74, 75, 76 nspartokinases of, 544 glycogrn synthesis, regulation of, 34 uridine diphosphoryl glucose pyrophosphorylase of, 63, 65 Sceitedesmirs obliquus, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Sedoheptulose diphosphate, adenosine diphosphoryl glucose pyrophosphorvlnsr and. 94-95, 97 Scdoheptulose 7-phosphate, phosphofructokinase and, 244 Selenomethionine, methionine adenosyltransferase and, 136

Serine methionine adenosyltransferase and, 137 pyruvate kinase and, 372 Serine phosphate, phosphoryl transfer and, 233 Serine residues acyl carrier protein, prosthetic group and, 167, 173 malonyl coenzyme A-acyl carrier protein transacylase, 178, 182 phosphorylase kinase, 559-580 protein kinases and, 555, 557, 575, 580 Serratia marcescens adenosine diphosphoryl glucose pyrophosphorylase of, 75, 76, 77, 107108 glycogen synthesis, regulation of, 34 Sex, methionine adenosyltransferase and, 143 Silkworm, uridine diphosphoryl glucose pyrophosphorylase in, 55 Slime mold, uridine diphosphoryl glucose pyrophosphorylase in, 55 Sodium acetate, protein kinases and, 576 Sodium chloride glycerol kinase and, 501 protein kinases and, 576 Sodium dodecyl sulfate, see Dodecyl sulfate Sodium ions 3-phosphoglycerate kinase and, 346, 348 phosvitin kinnse and, 581 Sorghum leaf, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Spectrophotometry, vitamin BIZ,adenosyltransferase assay and, 148 Spectinomycin, adenylylation of, 28-30 Spermatozoa glycerol kinase in, 490 phosphofructokinase of, 262, 266 Spermidine synthesis, S-adenosylmethioninc and, 124 Spinach acyl carrier protein amino acid composition, 159 4'-phosphopantetheine peptide of, 169, 170

648

SUBJECT INDEX

adenosine diphosphoryl glucose pyrophosphorylasc of, 86-90, 118 Spleen, nucleoside diphosphokinases of, 310, 311, 317

Stability constants, crcntine kinase and, 430

Stannous ions, guanidino kinases and, 472

Starch synthesis, 33 adenosinr diphosphoryl glucosc and. 74

uridine diphosphoryl glucose pyrophosphorylase and, 57-58, 91-92 Stereospecificity, pyruvate kinasc, 365366

Streptomycin. adcnylylation of, 2 8 3 0 Strontium ions, creatine kinase and, 450 Subunits arginine kinase, 468 aspartokinase I, 518-519, 522 aspartokinase 11, 541 bacillus aspartokinase, 548 creatine kinase, 394-395, 432 glycerol kinase, 495 nucleoside diphosphokinases, 331 phosphofructokinase, 254-257 phosphorylase kinase, 559, 560-561 protein kinases, 570-571, 577-578 pyruvate kinase, 358-359 Succinate glycogen labeling by, 82 phosphofructokinase and, 265-266 Succinate thiokinase, nucloside diphosphokinase activity of, 332 Succinic anhydride, phosphofructokinase and, 273 Succinyl coenzyme A formation from methylmalonyl cocnzyme A, 144 pyruvate kinase and, 377 Sucrose synthetase, starch synthesis and, 5758

Sugar beet leaf, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Sulfate activation of, S 3 7 adenosine diphosphoryl glucose pyrophosphorylase and, 79

creatine kinase and, 420-421, 423, 426 glycerol kinase and, 504 Sulfhydryl groups acyl carrier protein, 166-167 adenylate kinase, 284, 293-294 aspartokinase I, 521 P-ketoacyl acyl carrier protein synthetase. 190, 194 malonyl coenzyme A-acyl carrier protein transacylase, 182-183 nucleoside diphosphokinasc, 329, 330331

phosphofructokinase and, 269-272 3-phosphoglycerate kinase, 345, 350 pyruvate kinase, 360 uridine diphosphoryl glucose pyrophosphorylase and, 62

T n-Tagatose 6-phosphate, phosphofructokinase and. 244 Taurine, acyl mrrier protein and, 158. 159

Taurocyaminr kinase amino acid composition, 469 bivalcnt metal ions and, 472 discovery and isolation, 460 distribution of, 462 essential thiol groups of, 478 immunological properties, 470-471 molecular weight, 467 reaction mechanism, 483 stability, 468 substrate specificity, 474, 475 Temperature adenosine diphosphoryl glucose pyrophosphorylase kinetics and, 83-86 aspartokinasc I and, 517 creatine kinase and, 399, 400, 402, 420422

glycerate kinase and, 507 glycerol kinase and, 495 nucleoside diphosphokinase and, 329 phosphofructokinase and, 243 protein kinases and, 576 pyruvate kinase and, 361-362, 363 Tetradecanoyl acyl carrier protein, Pketoacyl acyl carrier protein synthetase and, 191, 193

SUBJECT INDEX

cis-Tetrahydrofuran-4-01-3-pl1enylphosphate, phosphoryl transfer and, 233 Trtranitromethane arginine kinase and, 481-482 cwatine kinase and, 455 Tetrathionate creatinc kinase and, 433, 434 guanidino kinases and, 481, 482 Tetrazolium dyes, nucleoside diphosphokinase drtrction and, 325 Thallium ions. pyrur-ate kinase and, 366367, 370 Thrrmodyn:imics glywrol kinase, 502 py ruva te k inase, 371-372 Thcrmolysin, malonyl coenzyme A-acyl carrier protein transacylase peptidr and, 182 Thiocynnnte ions, crrntine kinasc and, 426 Thiol groups, see Sulfhydryl groups Thiophosphinatc esters. inversion of configuration, 213 Thiolphosphonothiona te hydrolysis, racemization and. 213-214 Threoninr aspartokinases and, 512, 513, 518, 523. 525, 526, 545-546, 547, 549. 550. 551, 552, 553 circular dichroism, 535-536 difference spectra, 526-527 fluorescence, 527-532 relaxation studies, 533-535 biosynthesis of, 511 growth requirement for, 509, 510 Thymidine diphosphate nucleoside diphosphokinasc and, 323, 324, 325, 327, 328, 329 Thymidine diphosphoryl glucose formation of, 68 uridine dipliosphoryl glucose pyrophosphoryluse and, 67 Thymidine diphosphoryl rhamnose, uridine diphosphoryl glucose pyrophosphorylase and, 68 Thymidinr triphosphate glycerol kinase and, 499 nucleosidc diphosphokinases and, 322 phosphofructokinase and, 246 protein kinases and, 572

649 Thymus, nucleoside diphosphokinases of, 310, 323 Thyroid, phosphofructokinase of, 258 Tissues, uridine diphosphoryl glucose pyrophosphorylase in, 55 Tobacco leaf, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Tomato leaf, adenosine diphosphoryl glucose pyrophosphorylase of, 90 Transfer ribonucleic acid(s) riirboxyl group activation and, 12-13 enzyme caonformation and, 11 Transphosphorylation nonenzymic. metal ions and, 297-298 Trehalose, synthesis of, 59 Trichloroacetate, creatine kinase and, 427 Tricthylenetetramine cobalt ion, methyl phosphate hydrolysis and, 230-231 Trifluoromethylmethionine, methionine adenosyltransferase and, 136 Triglyeeride lipase, protein kinases and, 574 Triiodothyronine, uridine diphosphoryl glucose pyrophosphorylase synthesis and. 56 Trimetaphosphate, vitamin By,, adrnosyltransferase and, 152 Trinitrobenzene sulfonate, pyruvatr kinase and, 360, 370, 376, 382 Triose phosphate dehydrogenase, nucleoside diphosphokinase assay and, 324 Triosephosphate isomerase phosphofructokinase assay and, 244 separation from 3-phosphoglyrerate kinase, 341 Tripolyphosphatase, methionine adenosyltransferase and, 134, 136 Tripolyphospha te creatine kinase and, 407 mrthionine adenosyltransferase and, 130-132, 134, 135, 138, 140-141, 153 vitamin adenosyltransferase and, 146-147, 149-150, 152, 153 Troponin phosphorylase kinase and, 558-559, 564 Drotein kinases and. 574

650 Trypsin ncyl czrrier protein activity and, 171172, 173 aspartokinase I and, 521422, 536 aspnrtokinase I1 and, 541 creatine kinases and, 393, 394, 432. 433 glycerol kinasc and, 495 guanidino kinases and, 469, 470 phosphofructokinnse and, 255, 256, 257, 274 phosphoprotamine and, 575-576 phosphorylase and, 557 pyruvate kinase and, 359 Tryptophan, aspartokinase and, 547 Tryptophan residues ndenylate kinase, 291 aspartokinase I, 521, 528, 532, 536 pyruvate kinase, 361462, 373 Tubercidin 3’,5’-monophosphate, protrin kinase and, 573 Tumor tissue, uridine diphosphoryl glucose pyrophosphorylase in, 55 Tyrocidine hiosynthesis of, 17 composition of. 14 Tyrosine residues ncyl carrier protein, 173 arginine kinase, 481482 aspartokinase I, 528-529, 532 creatine kinase, 434-436, 454455 glutamine synthetase, ndenylylation of, 41 phosphofructokinase, 273

U Urea nrginine kinase and, 468-469 nspartokinase I and, 525 creatine kinase and, 388, 434 gunnidino kinases and, 478 innlonyl coenzymr A-acyl carrier protein transacylase and, 181, 184-185 nucleoside diphosphokinase and, 331 phosphofructokinase and, 253, 254, 255, 256, 270 pyruvate kinase and, 358, 369 Uridine 2’,3’-cyclic phosphorothioate, ribonuclease and, 236 Uridine diphosphate arginine kinase and, 476

SUBJECT INDEX

creatine kinasc and, 408, 409, 416, 417 nucleoside diphosphokinase and, 323 pyruvate kinase and, 364 uridine diphosphoryl glucose pyrophosphorylase and, 66, 67 Uridine diphosphoryl galactose, formation of, 68 Uridine diphosphoryl glucosamine, synthesis of, 54 Uridine diphosphoryl glucose photosynthesis and, 92 synthesis of, 32-33 uridine diphosphoryl glucose pyrophosphorylasc :md, 65-67, 70 Uridine diphosphoryl glucose dehydrogenase, uridine diphosphoryl glucose pyrophosphorylase assay and, 53 Uridine diphosphoryl glucose pyrophospholylase nnalytical nnd synthetic applications, 54-55 function, 51-52 measurrment of activity, 52-53 metabolic function cytology. 55-56 metabolism, 57-59 regulation, 59-62 properties kinetics, 65-68 mechanism, 69-71 optima, 62 specificity, 68-69 structure, 62-65 purification, 53-54 sources of, 53-54 Uridine diphosphoryl mannosc, formation of, 68 Uridine diphosphoryl xylose, formation of, 68 Uridine t riphosphate ndenylate kinase and, 299 glutamine synthetase and, 43 glycerol kinasc and, 498, 499 methionine adenosyltransferase and, 135 nucleoside diphosphokinases and, 322 phosphofructokinasc and, 246, 247, 267 3-phosphoglycerate kinase and, 348 protein kinases and, 572

651

SUBJECT INDEX

uridine diphosphoryl glucose pyrophosphorylase and, 65. 67, 70

V Vaccenate, formation of, 157 Valine, aspartokinase and, 547, 552 Vanadate ions, creatine kinase and, 426 Vitamin BIZ,nomenclature, 122 Vitamin B,2. rrductase, vitamin BIZ, ndenosyltransferase and, 147 Vitamin B,?, reductase, adenosyltransferase and, 145 Vitamin B1,,adenosyltrnnsferase catalytic properties activators and inhibitors, 151-152 assay, 148-149 kinetics and sribstrntr specificity, 150-151 reversibility, partial reactions and mechanism, 149-150 net reaction, 145-147 purification and physical properties, 147-148 significancr and distribution, 144-145 Vitamin E deficienry. crentinc kinase and, 386387

W Wnter tritiated, pyruvate kinase and, 380 Wheat leaf, starch synthesis in, 92 White blood cells, phosphofructokinase of, 257-258

X Santhosine diphosphnte, creatine kinase and, 409 Xanthosine inonophosphatc, guanosine monophosphnte synthesis from 3940

X-rays histone kinase and, 579 phosphofructokinase and, 271 X-ray crystallography 3-phosphoglycerate kinase, 341, 344346 pyruvate kinase, 359 Sylitol 1,5-diphosphate, ndenosine diphosphoryl glucose pyrophosphorylase and, 95

Y Yeast ncyl carrier protein amino acid composition, 158-180 4’-phosphopantetheine peptide of, 170 adenylate kinase mechanism, 303 physical properties, 296. 297 purification of, 289 substrate specificity, 299 nucleoside diphosphokinase of, 308, 314, 315, 316, 319-320, 322-323 phosphofructokinase, properties of, 242, 246, 247, 249, 261, 263, 265. 266, 267, 268, 275 3-phosphoglycerate kinase, 337-338 amino acid composition, 343 kinetic constants, 347 molecular weight, 342 X-ray crystallography, 344-345 pyruvate kinase purification of, 355-356 structure, 359

Z Zinc ions nucleoside diphosphokinases and, 329 3-phosphoglycerate kinase and, 347, 349 phosphoryl transfer and, 228-230 pyruvnte kinase and, 369-370

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