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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. .

.

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.

51 55 62

. . . .

. . .

. .

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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. .

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. .

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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


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


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


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


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.


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


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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.

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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.


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


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


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


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


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


-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


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.


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.


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


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


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


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.


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


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.


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.


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


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


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.


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


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).


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


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.


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.


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.


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.


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).

92

JACK PREISS

$

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


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.


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


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)


ADPglucose


0.9 0.11 0.06 0.14


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


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.


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.


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.


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).

3.


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.

111


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


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.


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

JACK PREISS

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.


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).


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).

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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.


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.


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-


133

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

8. HARVEY MUDD

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).

4.


135

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 .


137

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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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).

4. THE


139

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


141

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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S. HARVEY MUDD

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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THE ADENOSTLTRANSFERASES

143

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.

S. HARVEY MUDD

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.


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).

146

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


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).


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.


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


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.


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.


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.


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.


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


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


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 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.


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.


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.


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.


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.


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.


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.


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


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.


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


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


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


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


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).

7.

PHOSPHOFRUCTOKINASE

255

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


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


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.

PHOSPHOFRUCTOKINASE

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


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.

P HOSP HOFRCCTOKINA8E

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


(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).

7.

PHOSPHOFRUCTOKINASE

267

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).

268

DAVID P. B M X H A M AND H E N R Y A. LARDY

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).


269

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).

270


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.

7.

PHOSPHOFRUCTOKINASE

271

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).

272

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).

7.

PHOSPHOFRUCTOKINASE

273

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).

274


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.

7.

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275

(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).

276


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).


277

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

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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).

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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).

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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).

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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).

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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).


Nucleoside Diphasflhokinases R . E . PARKS. J R . I . Introduction .

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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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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


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


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.


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.


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


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.


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).


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.


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


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.


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


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


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.


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


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


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.


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).


3-Phosphoglycerate Kinase R. K. SCOPES

.

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 . . . . . . . . .

.

.

.

.

. . . .

. .

.

. . . .

. . . . . .

.

.

. .

. . . . .

. . .

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

. . . . . . . .

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


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


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.


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


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 I120 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.


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

KINASE

369

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

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

The Enzymes: Group Transfer, Part B. Phosphoryl Transfer. One-Carbon Group Transfer. Glycosyl Transfer. Amino Group Transfer. Other Transferases

The Enzymes, Vol IX: Group Transfer, Part B (Phosphoryl Transfer, One-Carbon Group Transfer, Glycosyl Transfer, Amino Group Transfer, Other Transferases), 3rd Edition

Transfer

Transfer

Transfer

Transfer

Transfer

Transfer

Transfer

TRANSFER

Seelen-Transfer

Zeit-Transfer

Radiative Transfer

Seelen-Transfer

Zeit-Transfer

Seelen-Transfer

Atlantik Transfer

Zeit-Transfer

MANHATTAN TRANSFER

Manhattan Transfer

Heat transfer

Zeit-Transfer

Heat transfer

Heat Transfer

Seelen-Transfer

Seelen-Transfer

Heat Transfer

Seelen-Transfer

Zeit-Transfer

Atlantik Transfer

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

The Enzymes: Group Transfer, Part B. Phosphoryl Transfer. One-Carbon Group Transfer. Glycosyl Transfer. Amino Group Transfer. Other Transferases

The Enzymes, Vol IX: Group Transfer, Part B (Phosphoryl Transfer, One-Carbon Group Transfer, Glycosyl Transfer, Amino Group Transfer, Other Transferases), 3rd Edition

Transfer

Transfer

Transfer

Transfer

Transfer

Transfer

Transfer

TRANSFER

Seelen-Transfer

Zeit-Transfer

Radiative Transfer

Seelen-Transfer

Zeit-Transfer

Seelen-Transfer

Atlantik Transfer

Zeit-Transfer

MANHATTAN TRANSFER

Manhattan Transfer

Heat transfer

Zeit-Transfer

Heat transfer

Heat Transfer

Seelen-Transfer

Seelen-Transfer

Heat Transfer

Seelen-Transfer

Zeit-Transfer

Atlantik Transfer

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