The Enzymes, Vol XVIII: Control by Phosphorylation, Part B (Specific Enzymes), 3rd Edition

The Enzymes VOLUME XVIII

CONTROL BY PHOSPHORYLATION Part B Specific Enzymes (11) Biological Processes Third Edition

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THE ENZYMES Edited by Edwin G. Krebs Paul D. Boyer Department of Chemistry and Biochemistry and Molecular Biology Institute University of California Los Angeles, California

Howard Hughes Medical Institute and Department of Pharmacology University of Washington Seattle, Washington

Volume XVIII

CONTROL BY PHOSPHORYLATION Part B Specific Enzymes (II) Biological Processes THIRD EDITION

1987

ACADEMIC PRESS, INC . Harcourt Brace Jovanovich, Publishers

Orlando San Diego New York Austin Boston London Sydney Tokyo Toronto

C O P Y R I G H T 0 1987 BY ACADEMIC P R E S S . INC. ALL RIGHTS RESERVED. NO PART O F THIS PUBLICATION MAY BE REPRODUCED O R TRANSMITTED IN ANY FORM O R BY ANY MEANS, ELECTRONIC O R MECHANICAL, INCLUDING PHOTOCOPY. RECORDING. O R ANY INFORMATION STORAGE A N D RETRIEVAL SYSTEM. WITHOUT PERMISSION IN WRITING FROM T H E PUBLISHER.

ACADEMIC PRESS, INC Orlando. Florida 32887

United Kingdom Edition published by

ACADEMIC PRESS INC. (LONDON) LTD. 24-28 Oval Road. London NW I 7DX

Library o f Congress Cataloging i n Publication Data (Revised for vol. 18, part B) The Enzymes. Includes bibliographical references. 1. Enzymes-Collected works. I. Boyer, Paul D.,ed. [DNLM: 1. Enzymes. QU 135 B791eI QP601.E523 574.1’925 75-117107 (v. 18 alk. paper) ISBN 0-12-122718-9

PRINTED IN THE UNITED STATES OF AMERICA

87 88 89 90

9 8 7 6 5 4 3 2

I

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

xi

Section 1. Control of Specific Enzymes (Continued)

1. Enzymes of the Fructose 6-Phosphate-Fructose 1,6Bisphosphate Substrate Cycle

SIMONJ. PILKIS,THOMASH. CLAUS,PAULD. KOUNTZ, AND M . RAAFAT EL-MAGHRABI 1. Introduction ................ 11. Purification of Hepatic 6-Phosphofructo-2-Kinase-Fructose-2,6-

4

..........

5

111. Assay of 6-Phosphofructo-2-Kinase Activity . . . . . . . . . . . . . . . . . . . . . . . . IV. Assay of Fructose-2,6-BisphosphataseActivity . . . . . . . . . . . . . . . . . . . . . . V. Structural Properties ..................... VI. Catalytic Properties o .................... VII. Catalytic Properties of Rat Liver Fructose-2.6-Bisphosphatase ......... VIII. Evidence for Two Catalytic Centers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Regulation of 6-Phosphofructo-2-Kinase-Fructose-2,6Bisphosphatase by Low-Molecular-Weight Effectors . . . . . . . . . . . . . . . . . . . . . . . . . . X. Regulation of 6-Phosphofructo-2-Kinase-Fructose-2,6Bisphosphatase by Phosphorylation-Dephosphorylation . . . . . . . . . . . . . . . . . . . . . . . XI. 6-Phosphofructo-I-Kinase:Possible Role of Phosphorylation in the Control of Enzyme Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . XII. Fructose-l,6-Bisphosphatase:Possible Role of Phosphorylation in Control of Enzyme Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . XIII. Role of 6-Phosphofructo-2-Kinase-Fructose-2,6-Bisphosphatase in the Hormonal Control of Hepatic Gluconeogenesis and Glycolysis . . . . . . . . . . . . XIV. Summary and Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .......

V

7 9 14 18 21

22 27 32

37 40 41

vi

C 0N TEN TS

2. Pyruvate Kinase L . ENGSTROM,

P. EKMAN,E. HUMBLE, AND 0. ZETTERQVIST .................................... . . . .

47

11. Influence of Phosphorylation on the Kinetic Properties of Liver Pyruvate Kinase to Proteolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. The Reaction of Cyclic AMP-Dependent Protein Kinase with Liver Pyruvate Kinase as Substrate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Dephosphorylation of Liver Pyruvate Kinase with Phosphoprotein Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Acute Hormonal Regulation of Liver Pyruvate Kinase in Vivo and .............. in Intact Cells

....

51

....

55

....

59

....

62

....

65 68 71 72

....

VI11. Concluding Remarks References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.... ....

3. Pyruvate Dehydrogenase LESTERJ . REED AND STEPHEN J . YEAMAN I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

11. Mammalian Pyruvate Dehydrogenase Complex . . . . . . . . . . . . . . . . . . . . . . . . .

111. IV. V. V1.

Pyruvate Dehydrogenase Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pyruvate Dehydrogenase Phosphatase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of Mammalian Pyruvate Dehydrogenase Complex Comparison of Properties of Mitochondria1 a-Ketoacid Dehydrogenase Kinases and Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , . .

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

17 79 82 a4 86

.... ....

92 93

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

97 100 103 112 118 119

.... ....

4. Branched-Chain Ketoacid Dehydrogenase PHILIP J . RANDLE, PHILIP

A.

PATSTON, AND

JOSEPH ESPINAL

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Animal Branched-Chain Ketoacid-Dehydrogenase Complex . . . . . . . . . . . . . . 111. Regulation by Reversible Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Biological Significance of Reversible Phosphorylation . . . . . . . . . . . . . . . . . . . V. Addendum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

5. Acetyl-Coenzyme A Carboxylase ROGERw. BROWNSEY AND

RICHARD

M. DENTON

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Structural Aspects and Regulation by Allosteric Effectors . . . . . . . . . . . . . . . . . . . . . 111. Short-Term Hormonal Regulation of Fatty Acid Synthesis

Associated with Persistent Changes in Acetyl-CoA Carboxylase Activity . . . . . . . . .

123 125 130

IV. Early Evidence for the Regulation of Acetyl-CoA Carboxylase

by Reversible Phosphorylation . . , . , . . , . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , , . . ,

134

CONTENTS

vii

V. Effects of Hormones on the Level of Phosphorylation of Acetyl-CoA Carboxylase within Intact Cell Preparations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Protein Kinases That Phosphorylate Acetyl-CoA Carboxylase . . . . . . . . . . . . . . . . . . VII. Protein Phosphatases That Act on Acetyl-CoA Carboxylase . . . . . . . . . . . . . . . . . . . . VIII. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References ............................................................

135 138 141 142 143

6. Hormone-Sensitive Lipase PETERSTRALFORS, HAKANOLSSON,AND PER BELFRAGE I. 11. 111. IV.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanism of Regulation of the Adipose Tissue Lipase Possible Role as a Hormone-Activatable, Multifunctiona

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

References

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

147 148 152 168 171 172

7. Hydroxymethylglutaryl-Coenzyme A Reductase DAVIDM. GIBSONAND REXA. PARKER I. 11. 111. IV. V. VI.

Introduction Topology ............................................................. Multivalent Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Reversible Phosphorylation in Vitro . . . . . . . . . . . . . . . Intracellular Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Reversible Phosphorylation and Degradation . . References ............................................................

180 180 185 195 199 206 210

8. Aromatic Amino Acid Hydroxylases SEYMOUR KAUFMAN

...

................................ .......................................... 111. Tyrosine Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I. Introduction .

IV. Tryptophan Hydroxylase References .....................

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

218 221 248 27 1 277

Section II. Control of Biological Processes

9. Phosphorylation of Brain Proteins S. IVAR WALAASAND PAULGREENGARD I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Protein Kinases in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

285 290

...

CONTENTS

Vlll

111. Phosphoproteins in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Protein Phosphatases in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

300 309 310 311

10. Regulation of Receptor Function JEFFREY L. BENOVICAND ROBERTJ. LEFKOWITZ I. Introduction and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. The P-Adrenergic Receptor . . . . . . . . . . . 111. Rhodopsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. The Nicotinic Acetylcholine Receptor . . .............................

V. The Receptors for EGF and Insulin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Other Membrane Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References .........................

319 320 328 329 329 330 331

11. Regulation of Ionic Channels SANDRAROSSIEAND WILLIAM A. CATTERALL I. 11. 111. IV. V. VI.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ................................. Calcium Channels . . . . . . . Potassium Channels . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . Acetylcholine Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sodium Channels ............ Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . .

335 337 342 347 351 354 354

12. Regulation of Protein Synthesis IRVINGM. LONDON,DANIEL H. LEVIN,ROBERTL. MATTS, N. SHAUNB. THOMAS,RAYMOND PETRYSHYN, AND JANE-JANE CHEN I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

11. Initiation of Protein Synthesis in Eukaryotic Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Role of eIF-2 in Eukaryotic Protein Chain Initiation

IV. V. VI. VII.

and the Effect of eIF-2a Phosphorylation Heme-Regulated eIF-2a Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . dsRNA-Dependent eIF-2a Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological Significance of HRI and dsI ........................ Guanine Nucleotide-Binding Proteins . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

360 360 362 369 371 373 376 377

13. Regulation of Contractile Activity JAMES R . SELLERSAND ROBERTS. ADELSTEIN I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Regulation of Vertebrate Smooth-Muscle Myosin by Phosphorylation

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

382 386

ix

CONTENTS 111. Role of Phosphorylation in Modulating Contractile Activity

of Striated Muscle Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV . Phosphorylation-Dependent Regulatory Systems in Invertebrate Muscles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V . Regulation of Cytoplasmic Myosins . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

404 406 406 412 413

14. Protein Phosphorylation in Prokaryotes and Single-Celled Eukaryotes

HOWARDV . RICKENBERCAND BEN H . LEICHTLING I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Protein Phosphorylation in Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Protein Phosphorylation in Single-Celled Eukaryotes ......................... IV . General Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

420 421 429 450 451

AuthorIndex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

451

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

503

Subjecrlndex

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Preface Over the past two decades there has been a remarkable increase in the recognition of the salient importance and the wondrous complexity of the control of enzyme catalysis. The modulation of enzymic and other protein-dependent processes by protein phosphorylation or dephosphorylation has emerged as the most widespread and important control achieved by covalent modification. So much information has emerged that adequate coverage in two volumes (XVII and XVIII) was a challenging task. The editors are gratified that the contributing authors have commendably met this challenge. The first portion of Volumes XVII and XVIII concerns the “machinery” of control by protein phosphorylation and dephosphorylation and includes coverage of the major types of protein kinases and of phosphoprotein phosphatases. The central core of the volumes presents chapters on the control of specific enzymes. This is followed by a substantial final section on the control of biological processes. The selection of authors for various chapters was a rewarding experience, but made somewhat difficult because for most topics there was more than one wellqualified potential author. The quality of the volumes was assured by the welcome acceptance of the invitation to participate by nearly all of the invited authors. The reversible covalent modification of enzymes and of proteins with other functions is now known to occur in all types of cells and in virtually all cellular compartments and organelles. Enzymes as a group constitute those proteins whose function and control are best understood in molecular terms. The treatment of enzymes gains additional importance because their regulation provides prototypic examples to guide investigators studying less well-defined and often less abundant proteins. The versatility of protein control by phosphorylation finds expression in ion channels, hormone receptors, protein synthesis, contractile processes, and brain function. Chapters in these areas point the way for future exciting developments. Although the breadth of coverage is in general regarded as satisfying, there are other topics or areas that may have warranted inclusion. These include the xi

xii

PREFACE

developing knowledge of the control by phosphorylation of histones of the nucleus and the messenger-independent casein kinases, whose role is not as clear as that of the major protein kinases that respond to regulatory agents. The quality of the volumes has been crucially dependent on the editorial assistance of Lyda Boyer and the fine cooperation provided by the staff of Academic Press. We record our thanks here. As readers of this Preface have likely discerned, it is a pleasure for the editors to have volumes of high quality to present to the profession. Paul D. Boyer Edwin G. Krebs

Section I

Control of Specific Enzymes (Continued)

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Enzymes of the Fructose 6Phosphate-Fructose 1 6Bisphosphate Substrate Cycle SIMON J. PILKIS* THOMAS H. CLAUSt M. RAAFAT EL-MAGHRABI*

PAUL D. KOUNTZ"

*Department of Molecular Physiology and Biophysics Vanderbilt University School of Medicine Nashville, Tennessee 37232 fAmerican Cyanamide Co. Medical Research Division Lederle Laboratories Pearl River, New York 10965

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ase- Fructose-2,6-

4

11. Purification of He

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

5

111. Assay of 6-Phosp

V. Structural Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Catalytic Properties of Rat Liver 6-Phosphofructo-2-Kinase . . . . . . . . . . . . . A. Phosphoryl-Acceptor Specificity .................... B. Phosphoryl-Donor Specificity ...................... C. Studies on Reaction Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Catalytic Properties of Rat Liver Fructose-2,6A. Substrate Specificity . . . . . . . . . . . . . . . . . . B. Product-Inhibitor Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I 9 9 12 12 14

14 I5

3 THE ENZYMES,Vol. XVlII Copyright Q 1987 by Academic Press, Inc. All rights of repruductiun in any lorn reserved.

4

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

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

C. Substrate Inhibition . . .

15

.. VIII. Evidence for Two Catalytic Centers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Effect of Thiol-Group Modification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

16

..................... B. Effect of Limited Proteolysis C. Effect of Adenine Nucleotide Analogs . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Effect of Histidyl Residue Modification . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of 6-Phosphofructo-2-Kinase-Fructose-2,6Bisphosphatase by Low-Molecular-Weight Effectors . . . . . . . . . . . . . . . . . . . Regulation of 6-Phosphofructo-2-Kinase-Fructose-2,6Bisphosphatase by Phosphorylation-Dephosphorylation . . . . 6-Phosphofructo-I-Kinase: Possible Role of Phosphorylation in the ................................... Control of Enzyme Activity A. Liver . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B . Skeletal Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Heart .......................... D. Ascaris ............................ Fructose- 1,6-Bisphosphatase: Possible Role of Phos Control of Enzyme Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ............................................... B. Yeast . . . . . , . . , . . . . , . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Role of 6-Phosphofructo-2-Kinase-Fructose-2,6-Bisphosphatase in the Hormonal Control of Hepatic Gluconeogenesis and Glycolysis . . . . . . . . Summary and Overview . . . . . . . . . . . . . . . , . . . . , . . . . . . . . . . . . . . . . . . . . . ................ References . . . . . . . . . . . . . .

20 21 21

D. Studies on the Reaction

IX. X. XI.

XII.

XIII. XIV.

1.

18 18

21 22 21 21 30 31 31

32 32 36

31 40 41

Introduction

The two enzymes responsible for the interconversion between fructose 6phosphate (Fru-6-P) and fructose 1,6-bisphosphate (Fru- 1,6-P,) in many cell types are 6-phosphofructo- 1-kinase (ATPm-fructose 6-phosphate 1-phosphotransferase, EC 2.7.1.11) and fructose-l,6-bisphosphatase (EC 3.1.3.11). 6Phosphofructo-1-kinase catalyzes the transfer of the terminal phospho group of ATP to the C-1 hydroxyl of Fru-6 P as shown in Eq. (1) while fructose-1,6bisphosphatase catalyzes the hydrolysis of Fru- 1,6-P, to yield Fru-6-P and Pi as shown in Eq. (2). Mg*

Fru-6-P

+

+ ATP @ Fru-1,6-P2 + ADP

FIX- I ,6-P2

+ H20 -+

FIX-6-P

+ P,

(2)

The 6-phosphofructo-1-kinase reaction represents the first committed unique step in glycolysis while the fructose- 1,6-bisphosphatase reaction represents an important step in the gluconeogenic pathway. Both enzymes are subject to a multiplicity of control mechanisms including reciprocal regulation by a number of allosteric effectors, changes in enzyme amount, and covalent modification. Re-

1.

5

FRU-6-P-FRU-I ,6-P* SUBSTRATE CYCLE

ciprocal regulation of these enzyme activities in liver has been shown to be mediated by fructose 2,6-bisphosphate (Fru-2,6-P2) (1-7). Furthermore, the synthesis and degradation of this sugar diphosphate is catalyzed by a unique bifunctional enzyme which is also subject to regulation by low-molecular-weight ligands, changes in enzyme amount, and covalent modification (5-7). 6-Phosphofructo-2-kinase-fructose-2,6-bisphosphatase (EC 2.7.1.105 and EC 3.1.3.46) catalyzes both transfer of the terminal phospho group of ATP to the C-2 hydroxyl of Fru-6-P as shown in Eq. (3) and the hydrolysis of Fru-2,6-P2 to Fru-6-P and Pi as shown in Eq. (4). F~x-6-P+ ATP

Mg'+ F t ~ - 2 , 6 - P *+ ADP

*

Fn1-2.6-P~+ H 2 0 + Fru-6-P

+ Pi

(3)

(4)

The activities of this enzyme determine the steady-state level of Fru-2,6-P, and ips0 fucto glycolytic and gluconeogenic flux in liver. It is the purpose of this chapter to review the regulation of these four enzyme activities, with particular emphasis on the role of phosphorylation. Many of the general regulatory properties of 6-phosphofructo-1-kinase and fructose- 1,6-bisphosphatase have been discussed before and the reader is referred to a number of excellent reviews (8-13). Some of the regulatory properties of the bifunctional enzyme are summarized here as a preface to reviewing its regulation by phosphorylation.

II. Purification of Hepatic 6-Phosphofructo-2-KinaseFructose-2,6-Bisphosphatase

6-Phosphofructo-2-kinase-fructose-2,6-bisphosphatase has been purified to homogeneity only from rat (14-16) and bovine (17) liver. The bifunctional enzyme has been detected only in the cytosol fraction from liver extracts and there is no evidence for particulate forms. Purification of the rat liver enzyme has depended on the ability to elute it specifically with substrate either from phosphocellulose (14, 15) or from a Fru-6-P-Sepharose affinity column (16). When measured at pH 7.4 and 30" and under optimal conditions the V,,, values of the kinase and bisphosphatase reactions are both about 60 nmol/min/mg. From this value the turnover number (kcat) can be calculated to be 0. l/s. This is one of the lowest turnover numbers known, indicating that it takes 10 s for the enzyme to turnover one time. The specific activities of the hepatic enzyme are 2-3 orders of magnitude less than that of most other phospho-group transferring enzymes. For example, rat liver 6-phosphofructo- 1-kinase has a specific activity of about 100 kmol/min/mg while rat liver fructose-l,6-bisphosphatasehas a specific activity of about 20-40 pmol/min/mg. Either the bisphosphatase or kinase activity has been detected and/or partially purified from other tissue sources including plants

6

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

(18, 19), yeast (20), rat heart (21), rat kidney, and bovine neutrophils (T. Chrisman and S . J. Pilkis, unpublished data). In most of these cases the bifunctionality of the protein has not been definitively established. However, both activities from neutrophils, rat kidney, and plants (18, 19) copurified, suggesting that the enzyme is probably bifunctional in these tissues. It is interesting to note that the reported specific activities of the plant enzyme (19) are orders of magnitude higher than those for the enzvme from mammalian liver. The rat liver enzyme is stable when stored in the presence of KCl and an appropriate reducing agent (14-16). Routine conditions for storage include 100 mM KCI, 0.5 mM dithiothreitol, 20% glycerol, and 50 mM Tris-HC1, pH 7.4. Under these conditions the enzyme does not lose activity for up to 3 months when stored at -70".

111. Assay of 6-Phosphofructo-2-Kinase Activity - Two methods have been used to measure 6-phosphofructo-2-kinase activity.

In the first method (22), the enzyme is incubated with Fru-6-P and ATP and the reaction terminated by the addition of 0.25 N NaOH followed by heating at 90" for 30 min. Fru-2,6-P2 is stable in hot alkali while Fru-6-P and other sugar monophosphates are destroyed (23). The pH is readjusted to neutrality with acetic acid, and the amount of Fru-2,6-P2 formed is determined by a 6-phosphofructo 1-kinase activation assay employing either rat liver 6-phosphofructo 1 kinase (22), skeletal muscle 6-phosphofructo 1-kinase (16), or the pyrophosphate-dependent enzyme from potato tubers (24). The potato enzyme has been reported to be an order of magnitude more sensitive to activation by Fru-2,6-P2 than most other 6-phosphofructo 1-kinases and is now available commercially. In the second method, Fru-6-P is incubated with Mg[y3,P]ATP and 6-phosphofructo-2-kinase in order to generate [2-32P]Fru-2,6-P, (7, 2 5 ) . To stop the reaction the sample is made 0.25 N in NaOH and heated at 90" for 30 min. Excess [y3,P]ATP is removed by charcoal treatment and the 32P-radioactivityin Fru-2,6-P2 counted. This assay can only be employed with purified enzyme or if 6-phosphofructo-1 -kinase has been removed.

IV. Assay of Fructose-2,6-Bisphosphatase Activity Four methods have been used to measure fructose-2,6-bisphosphataseactivity. In the first method (14, 1 3 ,disappearance of the substrate, Fru-2,6-P2, is measured with the 6-phosphofructo l-kinase activation assay (16, 2 2 , 2 4 ) . This assay is convenient for monitoring fructose-2,6-bisphosphataseactivity in crude tissue extracts or during purification.

1.

FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

7

In the second method (7, 25), the formation of 32Pifrom [2-32P]Fru-2,6-P, is measured using a DEAE-Sephadex column to separate 32Pi from unhydrolyzed substrate. Because Fru-6-P acts as a potent noncompetitive inhibitor, a Fru-6-P depleting system is needed to determine initial velocities (26). The third method takes advantage of the fact that the enzyme is phosphorylated on incubation with [2-32P]Fru-2,6-P, to form a phosphoenzyme intermediate which is labeled on a histidyl residue (27).Under these conditions the amount of [32P]E-P formed is directly proportional to the amount of enzyme protein. This is a very specific assay for the enzyme because, as far as is known, no other proteins form such a covalent linkage upon incubation with Fru-2,6-P2. The method is also very sensitive, permitting the detection of as little as 5 ng of enzyme protein and is useful for determining the amount of fructose-2,6bisphosphatase protein in crude extracts or during purification. The fourth method involves measuring the rate of formation or breakdown of the phosphoenzyme intermediate, which has been shown to be kinetically competent (26). The rate of formation of [32P]E-P from [2-32P]Fru-2,6-P, is monitored with a flow-quench instrument. The rates of dephosphorylation are slow enough to be determined by hand. In order to use many of the above assay procedures it is necessary to have [2-32P]Fru-2,6-P, of high-specific activity. Labeled fructose 2,6-bisphosphate can be prepared by first converting carrier-free 32P-inorganic phosphate to [y3,P]ATP enzymically (28).The labeled substrate is then prepared by incubating Fru-6-P and [y3,P]ATP with a homogeneous preparation of rat liver 6phosphofructo-2-kinase (25).

V. Structural Properties The subunit molecular weight of rat and beef liver 6-phosphofructo-2-kinasefructose-2,6-bisphosphatasehas been reported to be 50,000-55,000 by the criteria of SDS-gel electrophoresis (14-17). Only a single protein band is seen on both SDS one-dimensional (14-1 7) and two-dimensional gels (29, 30) provided the enzyme is either totally phosphorylated or dephosphorylated. The isoelectric point is 6.4 for the phosphorylated form and 6.6 for the dephosphorylated form (29,30). The apparent molecular weight of the native protein obtained either by gel filtration (14, 15, 17) or sucrose-gradient density centrifugation (this laboratory, unpublishedresults) is 100,000-1 10,000.The Stokes radius is 47 A. These results suggest that the enzyme is a dimer. Consistent with this notion, 2 mol of 32P are incorporated/mol of dimer upon incubation of the enzyme with CAMP-dependent protein kinase and [y3,P]ATP (14, 17, 31). Similar stoichiometry is observed when the enzyme is incubated with [2-32P]Fru-2,6-P, (26). These results also suggest, but do not prove, that the two subunits are identical.

8

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI TABLE I AMINO ACID

COMPOSITION OF

RAT LIVER6 - P H O S P H o r R U C T o - 2 - K I N A S E - ~ K U ~ r ~ ) S E - 2 ,

6-BISPHOSPHATASE" EquivalentslSubunit (55 kDa) Residues

24 h

48 h

72 h

96 h

Average

Aspartic acid Threonine Serine Glutamic acid Glycine Alanine Valine Methionine Leucine Isoleucine Tyrosine Phenylalanine Histidine Lysine Arginine Proline Tryptophanc Cysteine Total

41.2 22.2 24.9 62.4 23.1 27.8 27.3 5.8 42.0 26.8 27.9 12.8 13.4 23.1 32.2 18.5 -

42.3 21.2 21.6 65.0 19.7 27.8 31.9 6.1 42.3 27.2 28.6 13.4 13.6 23.4 35. 18.3

38.6 19.4 18.1 61.6 23.0 27.8 30.5 6.2 42.6 27.0 28.3 12.6 13.6 23.2 32.2 20.2

39.5 17.4 14.7 61.1 22.1 27.8 30.5 5.9 41.7 26.5 27.5 12.7 13.5 22.9 31.4 15.7

41

11.8

12.2

-

-

-

13.1

10.9

24" 28" 63 22 28 30 6 42 27 28 13 14 23 33 18 5 12 457

"Amino acid composition of 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatasewas determined with the Waters Pic0 Tag Amino Acid Analysis System after hydrolysis with 6 N HCI and a crystal of phenol at 115°C. "Extrapolated to zero time hydrolysis. Determined spectrophotometrically.

The amino analysis of the enzyme is given in Table I. The amino acid composition previously reported for the enzyme had a higher proportion of glycine and serine (15).The amino-terminal residue is blocked (31)and -His-Tyr are the carboxyl-terminal residues (27). Peptide mapping of the trypsin-treated enzyme using high-pressure liquid chromatography yielded the expected number of peptides (this laboratory, unpublished results) given that the enzyme contains 33 arginine and 23 lysine residues per subunit. Cyanogen bromide cleavage yielded fragments of 21,500,12,500,8200,5200,4100,1850, and 1850 daltons and their sum is close to the subunit molecular weight obtained by SDS-gel electrophoresis (this laboratory, unpublished results). The enzyme is likely a dimer of identical chains with a molecular weight approximately 1 10,000. The sedimentation coefficient is 58 k 0.2 sec-l (27). It has not yet been possible to dissociate the enzyme into monomers and retain activity.

9

I . FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

VI. Catalytic Properties of Rat liver 6-Phosphofructo-2-Kinase

A.

PHOSPHORYL-ACCEPTOR SPECIFICITY

Table I1 shows the sugar phosphate-specificity of 6-phosphofructo-2-kinase and the structure of a number of the analogs is shown in Fig. 1. 6-Phosphofructo-Zkinase appears to have a strict specificity for D-fructose 6-phosphate (I, see Fig. 1) as substrate (32). The only other sugar monophosphates that acted as substrate were epimers of the natural substrate, and of those only L-sorbose 6phosphate (11, Fig. 1) showed significant activity. The data suggest that while it is necessary to maintain the proper orientations at both the C-3 and C-4 hydroxyl groups for maximal activity, the orientation at C-4 is most important. The retention of significant activity with L-sorbose 6-phosphate (11, Fig. 1) suggests that the negatively charged group can still interact with the enzyme to some extent even though the phosphonoxymethyl portion of the moiety is in the opposite orientation. In Fig. 1 the predominant anomeric form of each sugar monophosphate is given. It is possible that differences in rates of phosphorylation may be accounted for by different proportions of anomeric forms, even though the rates of spontaneous anomerization between the a-and p-forms are probably substantially greater than the rate of the reaction (turnover number = 0.1/s). However, it would appear that differences in the proportion of anomeric forms cannot completely explain the differences in rate since both D-tagatose 6-phosphate (111, Fig. 1) and D-fructose 6-phosphate are predominantly in the p-form in solution but their K,,, values for the enzyme differ by 400-fold. TABLE 11 SUGAR

PHOSPHAK

Sugar phosphate o-Fructose 6-phosphate L-Sorbose 6-phosphate o-Psicose 6-phosphate D-Tagatose 6-phosphate a- and P-methyl+ fructofuranoside 6-phosphate I-0-methylfructose 6-phosphate 2,5-Anhydro-o-mannitol 6phosphate o-Arabinose 5-phosphate

SPECIFICITY OF 6-PHOSPHOFKUCTO-2-KINASE

K,, analog K,, (d) Log K,,, Fm-6-P

0.035 0.175 7.4 15.0

0 0.7 2.3

2.6

Relative V,,,, I .o 1.1

0.42 0 . I5

Catalytic efficiency Mimin X 10-3 171

38 0.34 0.06

8-D-FRUCTOFURANOSE 6-P

8-0- TAGATOFURANOSE 6-P

a -L- SOR 80FURANOSE 6-P

a-D-PSICOFURANOSE 6 - P

on

(I) 1-0 METHYL DFRUCTOFURANOSE 6 - P

8-D-RlBOFURANOSE 5-P

B-D-ARABINO FURANOSE 5 - P

pocQHHOH 0-H

OH

OH

.

(nr)

(P) 2.5 ANHYDRO-D-

MANNITOL 6-P

(a+B)METHYL D-FRUCTO FURANOSIDE 6 - P

FIG. 1. Structure of epimers and various substrate analogs of D-fructose 6-phosphate.

11

I . FRU-6-P-FRU- I ,6-P2 SUBSTRATE CYCLE

Modification of either of the moieties at C-1 or C-2 of D-fructose 6-phosphate also resulted in loss of activity. 1-0-methyl-D-fructose 6-phosphate (V, Fig. 1) was not phosphorylated by 6-phosphofructo-2-kinase, and absence of the hydroxymethyl group, as with D-arabinose 5-phosphate (VI, Fig. 1 ) or D-ribose 5phosphate (VII, Fig. l), also resulted in complete loss of activity. The lack of activity with D-arabinose Sphosphate, which otherwise resembles the natural substrate, strongly points to the requirement of the hydroxymethyl group in the substrate. The failure of either 2,5-anhydro-~-mannitol6-phosphate (VIII, Fig. 1) or (a P)-methyl-D-fructofuranoside6-phosphate (IX, Fig. 1) to act as a substrate for the enzyme suggests the importance of the free anomeric hydroxyl group. The importance of this group is certainly not unexpected since it is the site to which phosphate is transferred from ATP. Thus, the substrate specificity of 6phosphofructo-2-kinase requires a 2-hydroxymethyl-2,3,4-trihydroxy-5-phosphonoxymethyl tetrahydrofuran structure, with the P-hydroxyl group at C-3 cis to the p-anomeric hydroxyl group. This same orientation is preferred for the phosphonoxymethyl moiety at C-5, while the opposite orientation is required for the hydroxyl group at C-4. Modification at each carbon results in loss of activity but inversion of the phosphonoxymethyl moiety at C-5 has the least effect on binding. The substrate specificity for the liver 6-phosphofructo-2-kinase is more strict than that for 6-phosphofructo- 1-kinase from muscle. Muscle 6-phosphofructo- 1kinase can phosphorylate a number of sugar phosphates including D-fructose 1phosphate which is phosphorylated at the C-6 hydroxyl (33), D-glucose l-phosphate (34,D-sedoheptulose 7-phosphate ( 3 9 , D-fructose 6-sulfate (36),as well as L-sorbose 6-phosphate (37). The relative rates of phosphorylation of the epimers suggest that the substrate specificity of muscle 6-phosphofructo-1-kinase requires a 2-hydroxymethyl-3,4-dihydroxy 5-phosphonoxymethyl tetrahydrofuran structure with both the hydroxyl group at C-3 and the phosphonoxymethyl group at C-5 oriented cis to the p-anomeric hydroxyl group. The hydroxyl group

+

TABLE 111 INHIBITION OF 6-PHOSPHOPRUCTO-2-KINASEB Y

Sugar monophosphate 2,5-Anhydro-~-mannitol 6-phosphate o-Ribulose 5-phosphate o-Ribose 6-phosphate o-Arabinose 5-phosphate (a+P)Methyl-o-fructofuranoside 6phosphate I-0-Methyl-o-fructose 6-phosphate

SUGAR MONOPHOSPHATES

K,

Inhibition type Competitive

0.096(mM)

Competitive Competitive Competitive none

2.5

-

none

-

8.7 10

12

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

at C-4 can be either cis or trans to the p-anomeric hydroxyl group. However, the anomeric hydroxyl group is not required in order for the sugar phosphate to be a substrate for 6-phosphofructo 1-kinase since 2,5-anhydro-~-mannitol 6-phosphate is an excellent substrate (38). Thus, there are striking differences between substrate specificities of the two kinases. As shown in Table 111, the only effective inhibitor of 6-phosphofructo-2kinase from among these sugar phosphate analogs was 2,5-anhydro-~-mannitol 6-phosphate with a K , of about 100 pM. The inhibition was competitive, indicating that 2,5-anhydro-~-mannitol 6-phosphate can bind at the kinase active site, but this binding is nonproductive because of the lack of the C-2 hydroxyl group. There is evidence that inhibition of the 6-phosphofructo-2-kinase by 2 3 anhydo-~-mannitol6-phosphate occurs in the intact cell under certain conditions (39, 40).

B . PHOSPHORYL-DONOR SPECIFICITY The rat liver 6-phosphofructo-2-kinase reaction utilizes ATP and to a lesser degree GTP (22, 23) as phosphoryl donors but the phosphoryl-donor specificity is very different from 6-phosphofructo- 1-kinase from rat liver or muscle which can use a wide variety of nucleoside triphosphates as phosphoyl donors in the catalytic reaction. Furthermore. in contrast to the muscle and liver 6-phosphofructo-1-kinase, which exhibit allosteric inhibition by ATP, increasing concentrations of ATP above the catalytic optimum does not cause inhibition of 6phosphofructo-2-kinase (15, 16, 22). C. STUDIESON REACTION MECHANISM The enzyme has been shown to catalyze hydrolysis of ATP in the absence of added sugar phosphate or sugar diphosphates (25, 41). The existence of an ATPase activity suggests, but does not prove, that the reaction mechanism of the 6-phosphofructo-2-kinaseinvolves a two-step transfer mechanism that includes a phosphoenzyme intermediate. Support for the existence of a phosphoenzyme intermediate was provided by the discovery that the enzyme catalyzed exchange reactions between ADP and ATP and between Fru-6-P and Fru-2,6-P2 (25, 42). The adenine nucleotide exchange reaction occurs in the presence of a Fru-6-P trapping system at a rate which was 20% that of the kinase reaction. The sugar phosphate exchange occurs at a rate nearly 20% that of the kinase or bisphosphatase reaction and is not affected by inclusion of glucose and hexokinase to trap ATP. The sugar phosphate exchange reaction is almost completely dependent on the presence of Pi (42). The existence of these exchange reactions is the only direct evidence for covalent catalysis in the kinase reaction.

1.

FRU-6-P-FRU-I ,6-Pz SUBSTRATE CYCLE

13

In contrast, Kitajima et al. (43) could not detect sugar phosphate exchange. Moreover, the adenine nucleotide exchange has been shown to be stable to a variety of protein-modifying reagents that affect kinase activity, suggesting that the exchange may be unrelated to the kinase or that the modifications affected the sugar phosphate site (42).That the ADP-ATP exchange may be unrelated to the kinase reaction is further suggested by the observation that Fru-6-P is a weak rather than potent inhibitor of exchange (43).While it is clear that the ADP-ATP and Fru-6-P-Fru-2,6-P2 exchange reactions are authentic exchange reactions, it is not clear whether the phosphoenzyme intermediates of the exchange reactions are involved in the net reactions. It has not been possible to isolate a phosphoenzyme intermediate upon incubation of the enzyme with [y-32P]ATP(25). The question of whether intermediates are involved in the normal reaction mechanism of the kinase can best be resolved by following the stereochemical course of the reaction (44). Preliminary work on the question has been in progress. With [y-(S)- l6O, l7O, I80]ATP as substrate, [2-160, 170, 180]Fru-2,6-P, has been produced by the kinase reaction, the chiral phospho group transferred to 1,3-butanediol by alkaline phosphatase (the stereochemistry of which is known), and the configuration about the phosphorus determined. It was found that the reaction proceeded by net inversion of the configuration at phosphorus. The most reasonable interpretation of this data is that the reaction proceeds via a single inline displacement and does not involve a phosphoenzyme intermediate, though it is not possible to rule out any odd number of multiple single displacements that might include formation of a phosphoenzyme. There have been a number of steady-state kinetics studies on the enzyme. The kinase reaction is inhibited by both of its products (5, 6, 25, 43). ADP is a competitive inhibitor with respect to ATP with a K i of 0.6 mM (5, 25, 43, 45). All other inhibition patterns including ADP versus Fru-6-P (43) and Fru-2,6-P, versus Fru-6-P or ATP are noncompetitive (5, 25, 43, 45). The apparent K i for Fru-2,6-P, is about 0.2 mM (45).The product inhibition pattern is not consistent with a ping-pong mechanism, nor is it consistent with any other straight-forward reaction mechanism. For example, it has been postulated that the pattern is consistent with a sequential-ordered mechanism (43),with ATP binding first and then Fru-6-P. However, the existence of the ADP-ATP exchange reaction precludes such a mechanism with the current data. If the exchange takes place at the active site, then covalent catalysis and thus a different mechanism is implicated. However, if the exchange occurs at a separate (allosteric) site, then any steadystate kinetics analysis to determine the reaction mechanism would be difficult to interpret. This may be the case with the 6-phosphofructo-2-kinase. The 6-phosphofructo-2-kinase reaction has been shown to be reversible and Pi was found to stimulate the reversal of the kinase reaction (42, 4 3 ) . When the enzyme was incubated with [2-32P]Fru-2,6-P, and ADP, the rate of [y-32P]ATP

14

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

production was only about 2% of the forward reaction. Addition of a Fru-6-P depleting system or Pi increased this rate by 2-fold and 7-fold, respectively. In the presence of both, the reverse reaction was stimulated by more than tenfold to a rate equal to 30% of the forward reaction (42). Inorganic phosphate has been shown to be an activator of the 6-phosphofructo-2-kinase reaction with the activation being characterized by an increase in affinity for Fru-6-P (5, 6, 17, 22, 23, 46). The concentration of Pi necessary to elicit these effects is about 0.5 mM (45).Since the reverse kinase reaction and sugar phosphate exchange are almost totally dependent on Pi, it raises the question of whether Pi directly participates in the reaction. Moreover, after alkylation of the enzyme by iodoacetamide, the forward kinase reaction appears to be almost entirely dependent on Pi (45).Carboxamidomethylation of 2 cysteine residues per enzyme subunit caused a 12-fold stimulation of the kinase V,,, but this effect was only seen when the kinase was assayed in the presence of Pi. The effects of Pi on the forward and reverse reactions of the kinase and on the sugar phosphate exchange all appear to involve enhanced affinity for sugar phosphate, while the 12-fold increase in the alkylated kinase V,,, suggests that Pi may also influence the turnover number per se. Laloux et al. (47) claimed that the native rat hepatic 6-phosphofructo-2-kinase is totally dependent on Pi and that the enzyme from yeast and spinach leaves has a “nearly complete phosphate dependency.” However, the liver enzyme preparation used was only 10% pure and had an ATPase activity equal to 50% of the kinase activity. This apparent phosphate dependency may be related to Pi effects on contaminating activities and, in fact, it is not observed with a homogeneous preparation of the enzyme (P. D. Kountz, M. R. El-Maghrabi, and S. J . Pilkis, unpublished results).

VII. Catalytic Properties of Rat liver Fructose-2,6Bisphosphatase

A.

SUBSTRATE SPECIFICITY

So far as is known fructose-2,6-bisphosphatase is absolutely specific for Fru-2,6-P2. Other sugar disphosphates that have been tested as substrates for the rat liver fructose-2,6-bisphosphatasewith negative results include Fru- 1,6-P,, sedoheptulose-1,7-P,, arabinose- 1,5-P,, ribose-1,5-P2, g1ucose-l,6-P2, sorbose-2,6-P2, psicose-2,6-P2, and tagatose-2,6-P2 (S. J. Pilkis, unpublished results). This apparent absolute specificity is in contrast to the mammalian liver fructose-1,6-bisphosphatase which acts on sedoheptulose- I ,6-P, (48)with nearly the same V,,, as Fru-1,6-P2, although the affinity for the higher homolog is considerably lower than for Fru-l,6-P2.

1.

B.

FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

15

PRODUCT-INHIBITOR SPECIFICITY

It has been demonstrated that Fru-6-P is a potent noncompetitive inhibitor of the fructose-2,6-bisphosphatase(2, 4-6, 14, 17, 25, 26, 42, 43, 45, 49). The structural requirements for sugar monophosphate inhibition of the bisphosphatase are similar to the requirements for a substrate in the kinase reaction. LSorbose 6-phosphate (10,5= 0.05 mM) was almost as good an inhibitor of the bisphosphatase as was D-fructose 6-phosphate (Io,5 = 0.01 mM), while Dpsicose 6-phosphate (lo,5= 0.9 mM) and D-tagatose 6-phosphate (Io.5 > 2.5 mM) were much poorer inhibitors. Similarly, 2,5-anhydro-~-mannitol6-phosphate inhibited the bisphosphatase at concentrations = 0.5 mM) that were approximately the same as those effective in inhibiting the kinase. D-Ribose 5phosphate was a very poor inhibitor of the bisphosphatase (Io,5 2 10 mM), just as it was a poor inhibitor of the 6-phosphofructo-2-kinase(Ki= 10 mM). These results suggest that the functional determinants at the sugar-phosphate-binding site of the kinase and the bisphosphatase are similar if not identical.

C. SUBSTRATE INHIBITION Early kinetic studies on the fructose-2,6-bisphosphatasereaction indicated a K , in the range of 0.1-20 pA4 for Fru-2,6-P2 and a V,,, of 50-70 nmol/min/mg at 30". Fru-6-P was reported to be a potent product inhibitor and Pi and aglycerol-P were shown to be activators of the reaction over the range of 1-100 pA4 Fru-2,6-P2 (14, 16, 25, 31, 43, 49). Under these conditions both activators increased the apparent Kifor Fru-6-P inhibition. However, it has been shown that the true V,,, of the reaction can only be obtained in the presence of a Fru-6-P depleting system rather than by addition of Pi or a-glycerol-P (26).The response of the enzyme to Fru-2,6-P2 under these conditions is shown in Fig. 2. The dependence on substrate was hyperbolic below 100 nM in the absence of Pi and a-glycerol-P, the K,,, was 4 nM and the V,,, at 22°C was 12 nmol/mg/min at 20-50 nM Fru-2,6-P2. Substrate inhibition was observed above 100 nM Fru-2,6P,: The velocity obtained at 1 pM was only 70% of that obtained at 50 nM and at 10 pA4 the rate was only 14%. a-Glycerol-P or Pi strongly inhibited the hydrolytic rate observed below 50 nM of substrate and they increased the apparent K , over 20-fold to about 100 nM Fru-2,6-P2 (Fig. 2). At high substrate concentrations both effectors enhanced enzyme activity. Inhibition of hydrolysis by Pi and a-glycerol-P at low substrate concentrations is competitive and the Ki value obtained from Dixon plots is about 0.5 mM for both Pi and a-glycerol-P (26). This suggests that the apparent stimulation of hydrolysis by these effectors with enzyme assayed at high substrate reported in early studies was due to relief of substrate and product inhibition. The simplest explanation for the effects of Pi or a-glycerol-P to inhibit hydrolysis at sub-

16

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

a-glycerol- P

I

0

10

30

50

FRU -2,6 - P2, nM

0

1

I

500 FRU-2,6-P2, nM

1

lpoo

FIG. 2. Substrate concentration dependence of rat hepatic fructose-2,6-bisphosphatase.(A) Substrate concentration range 0-50 nM. (B) Substrate concentration range 0-1000 nM.

saturating substrate concentrations and to relieve substrate inhibition at saturating concentrations is that these ligands antagonize Fru-2,6-P2 binding. It was reported earlier that Pi actually enhanced affinity for Fru-2,6-P2 in the fructose-2,6-bisphosphatase reaction (5, 6 , 14). This apparent increase in affinity was probably due to relief of substrate inhibition seen when high concentration of substrate were employed in the absence of a Fru-6-P depleting system.

D. STUDIESON

THE

REACTIONMECHANISM

Evidence has accumulated that supports the notion that the fructose-2,6bisphosphatase reaction involves a two-step transfer mechanism that includes a phosphoenzyme intermediate ( 5 , 6 , 25). Such an intermediate has been isolated after incubation of the enzyme with [2-32P]Fru-2,6-P, and has been identified as 3-phosphohistidine ( I 7, SO). Work proving that this phosphoenzyme is an obligatory intermediate in the reaction (26) can be summarized as follows: 1 . Both the formation and breakdown of the phosphoenzyme in the absence and presence of Pi and a-glycerol-P were sufficiently fast for E-P to be a reaction intermediate. 2. At low substrate concentration, the steady-state level of phosphoenzyme

1.

17

FRU-6-P-FRU-I ,6-Pz SUBSTRATE CYCLE

3. 4.

5.

6.

correlated with hydrolytic rate at varying substrate and effector concentrations. The rate of phosphoenzyme breakdown increased as pH was lowered, corresponding to the increased hydrolytic rate at lower pH, both in the absence and presence of Pi or a-glycerol-P. The steady-state level of phosphoenzyme decreased as the pH was lowered, probably reflecting the increased rate of phosphoenzyme breakdown. Inhibition of E-P breakdown at very high substrate concentrations correlated with substrate inhibition. The net rate of the reaction could be approximated by the product of the fractional rate of phosphoenzyme breakdown and the amount of phosphoenzyme found in the steady state over a broad pH range, both in the absence and presence of Pi and a-glycerol-P.

Scheme I depicts the reaction mechanism for the fructose-2,6-bisphosphatase. The interpretation of the effects of phosphate and a-glycerol-P on fructose-2,6-bisphosphatase has been difficult since they have complicated effects on E-P, vis-A-vis net hydrolysis. The net effect of Pi and a-glycerol-P is to decrease the steady-state level of E-P by both inhibiting its formation and accelerating its breakdown. The decrease in steady-state E-P and hydrolysis by both Pi and aglycerol-P at low Fru-2,6-P2 concentrations (100 nM) substrate concentrations they also inhibit the rate of formation of E-P. Since net hydrolysis is enhanced by both Pi and a-glycerol-P at high substrate concentrations, acceleration of E-P breakdown must be the predominant effect. Inhibition of E-P breakdown and of Fru-2,6-P2 hydrolysis by Fru-2,6-P2 is seen at substrate concentrations in excess of 100 nM and this is largely overcome by both Pi and a-glycerol-P. It has been suggested that a-glycerol-P activates fructose-2,6-bisphosphatase at substrate concentrations above 1 by decreasing the interaction of the enzyme with Fru-6-P, due to its chemical resemblance to the C-4-C-6 portion of the sugar monophosphate (25). Phosphate may also antagonize the binding of the sugar

E. Fru-2,6-Pz Fru-2,6-Pz E-P .Fru-6-P

E

SCHEMEI

18

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

monophosphate since it also increases the apparent K j for Fru-6-P (5, 6, 25). Thus it would appear that the stimulation of hydrolysis by Pi and a-glycerol-P above 100 nM substrate is due to both relief of substrate inhibition by accelerating E-P breakdown and to relief of inhibition by Fru-6-P. The results of steady-state kinetics studies have resolved the dichotomy of the product inhibition pattern observed in earlier studies of the fructose-2,6bisphosphatase; namely, that one product of the reaction, Fru-6-P, was a noncompetitive inhibitor of the reaction while the other product, Pi, activated the enzyme. This was in contrast to findings obtained with the fructose-2,6bisphosphatase found in plants (18) and most hepatic fructose- 1,6-bisphosphatases which are inhibited by both Fru-6-P and Pi (13, 51, 52). Pi is a competitive inhibitor of net hydrolysis at low substrate concentrations but an activator at saturating concentrations (26). The product inhibition pattern of fructose-2,6bisphosphatase is consistent with either an ordered sequential mechanism, with Pi released last, or a ping-pong sequence with a phosphoenzyme intermediate. The results are also consistent with two rapid-equilibrium sequential processes (13, 51, 52). The weight of evidence supports the rapid equilibrium schemes for the fructose-l,6-bisphosphatase (13, 51) while the demonstration of a phosphoenzyme intermediate for the fructose-2,6-bisphosphatase strongly supports a ping-pong sequence in this case. Previous calculations of the catalytic efficiency of the fructose-2,6-bisphosphatase reaction ( 5 ) were underestimates since the affinity constant (K,) used in the calculation was in the micromolar range (5). Using a K , value of 4 nM and a turnover number (kcat) of 6 min, the catalytic efficiency of fructose-2,6bisphosphatase can be calculated to be 1.5 X lo9 Mlmin. The value of 6/min for k,,, is calculated from the hydrolytic rate obtained at 30°C. In comparison the catalytic efficiency for rabbit hepatic fructose- 1,6-bisphosphatase can be calculated to be 3 X lo9 Mlmin using values for k,,, (6 X 102/min) and K , (0.2 X M) from the work of Benkovic and deMaine (13). Though the turnover numbers (kcat) of these two bisphosphatases differ by lOO-fold, both enzymes operate at essentially the same catalytic efficiency because the tighter binding of Fru-2,6-P2 to the fructose-2,6-bisphosphatasecompensates for the lower turnover number of this enzyme.

VIII. Evidence for Two Catalytic Centers A.

EFFECTOF THIOL-GROUP MODIFICATION

A mixed-function oxidation system consisting of ascorbate-Fe3 completely inactivated the kinase as well as the Fru-6-P-Fru-2,6-P2 exchange reaction but had no effect on either the fructose-2,6-bisphosphataseor the ADP-ATP exchange reaction (7, 45, 53). That H,O, was involved in the modification was +

1.

19

FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

confirmed by using H,O, itself and by preventing the inactivation with catalase. Irreversible oxidative-destruction of a histidine residue has been reported to be the cause of inactivation of some of the enzymes tested with these systems (54). Since inactivation of the bifunctional enzyme was readily reversible by dithiothreitol, it appears that the modified residue in this case is probably a cysteine rather than histidine. Oxidation of the enzyme may involve the formation of either a disulfide bond or of a sulfenic acid (R-SOH). The oxidative-inactivation of a number of enzymes via formation of a sulfenyl derivative has been reported to be reversible by the addition of thiols (55-57). The inactivation of the kinase and sugar phosphate exchange and its reversal by thiols suggest that there are essential cysteine residues at or associated with the kinase active site. Alkylation of 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatasewith pmercuribenzoate caused a rapid stimulation of the kinase and an inhibition of the bisphosphatase whereas treatment with N-ethylmaleimide abolished kinase activity but had no effect on the bisphosphatase. Selective modification of residues involved in the kinase reaction was also seen with iodoacetamide which caused a 10-fold stimulation of the kinase V,,, without affecting the bisphosphatase. However, the stimulatory effect of carboxyamidomethylation was seen only when the kinase was assayed in the presence of inorganic phosphate. The iodoacetamide-treated enzyme had a 10- to 20-fold higher K , for fructose 6phosphate than the native enzyme and the Kifor fructose 2,6-bisphosphate was also increased. However, the adenine nucleotide site was not affected since there was no change in the K , for ATP, the Ki for ADP, or the adenine nucleotide exchange reaction. The residues modified by iodoacetamide were shown to be cysteines by the exclusive appearance of carboxymethylcysteine in protein hydrolysates. Activation was associated with alkylation of 2 cysteines per subunit, of the 12 that could be alkylated after denaturation-reduction. Iodoacetamideactivated kinase was also inhibited by ascorbate-Fe3 . There is an analogy between the effects of oxidation of sulfhydryl groups, formation of disulfide or sulfenyl derivatives, and their alkylation with iodoacetamide in that both treatments caused a decrease in the affinity of the kinase for Fru-6-P. In the case of oxidation by ascorbate-Fe3 , sugar phosphate affinity appears to be abolished, while alkylation causes a 20-fold reduction in affinity for the sugar phosphate and sugar diphosphate in addition to a marked increase of the maximal activity of the enzyme. The increase in kinase V,,, with alkylation may be a consequence of the decrease in affinity for Fru-2,6-P2. This would allow faster dissociation of nascent product and reduce product inhibition. The dissociation of the Fru-2,6-P2 from the enzyme may be the rate-limiting step of the kinase reaction. The differences in the response of the kinase and bisphosphatase to sulfhydryl modification suggest that there are separate and distinct sugar phosphate sites for the kinase and bisphosphatase. This conclusion is also supported by finding that alkylation of the enzyme by N-bromoacetylethanolamine phosphate results in +

+

20

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI TABLE IV

EFFECTOF IODOATETAMIDE TREATMENT O N SUGAR PHOSPHATE APFINITY OC. 6-PHOFRUCTO-2-KINASE-FRUCI.OSE-2.6-BlSPHOSPHATASF~

Kinase K,,, (d)

Bisphosphatase lo-5, (d)

Sugar phosphate

Native

IAM-treated

Native

o-Fructose 6-phosphate L-Sorbose 6-phosphate o-Psicose 6-phosphate o-Tagatose 6-phosphate 2.5-AM-6-P

0.035 0.175 7.4 15.0

I .o >2.0 N.D.1' N.D."

-

-

0.01 0.05 0.90 >2.50 0.50

IAM-treated 0.01

0.06 1.1

>2.50 0.70

"No phosphorylation was detected with sugar phosphate concentrations up to I 5 d

loss of 90% of the kinase activity but with no loss in bisphosphatase activity or change in inhibition by Fru-6-P (58). Additional evidence that the sugar phosphate binding sites of the 6-phosphofructo-2-kinase and fructose-2,6-bisphosphatase are different was obtained by studying the effect of iodoacetamide on the apparent affinity of sugar phosphate as substrate for the kinase and as product inhibitor of the bisphosphatase (32). As shown in Table IV, alkylation of the enzyme increased the K,, for D-fructose 6-phosphate as well as the K,, values for the three epimers for the kinase reaction. Little or no phosphorylation of Dpsicose 6-phosphate or D-tagatose 6-phosphate was detected at concentrations up to 12 mM. Thus, aklkylation decreased the affinity for all the epimers of Dfructose 6-phosphate in the kinase reaction. In contrast, the Io.5 values of the bisphosphatase for D-fructose 6-phosphate and its epimers were unaffected by iodoacetamide treatment. These results strongly suggest that there are discrete sugar phosphate sites. At the same time, these sites appear to have essentially identical structural requirements for sugar phosphate interaction suggesting that they have a high degree of homology. B.

EFFECTOF LIMITEDPROTEOLYSIS

Limited proteolysis of the enzyme with thermolysin yielded an enzyme core with a subunit molecular weight of 35,000-38,000 (53).This enzyme core had no kinase activity but had a 2-fold activated bisphosphatase activity whose sensitivity to the product inhibitor Fru-6-P was unchanged. The thermolysin-treated enzyme also did not catalyze the Fru-6-P-Fru-2,6-P2 exchange reaction but did catalyze the ADP-ATP exchange. These results suggest that ( a ) the enzyme's reactions may be catalyzed at two active sites; (b) there are at least two Fru-6-P binding sites; ( c )the Fru-6-P-Fru 2,6-P, exchange is catalyzed only at the kinase site; and (d)inactivation of the exchange and kinase reactions by thermolysin

1.

FRU-6-P-FRU-I ,6-Pz SUBSTRATE CYCLE

21

digestion is due to the loss of the Fru-6-P binding site of the kinase. Also consistent with these conclusions was the finding that limited proteolysis with trypsin yielded a cleavage product with a molecular weight of 50,000 which had no kinase activity but whose bisphosphatase was unaffected (59). The ADPATP exchange was lost upon trypsin treatment but the K jfor Fru-6-P of fructose-2,6-bisphosphatase was not altered. Partial protection against the trypsin proteolysis was provided by ATP, Fru-6-P, and Fru-2,6-P2.

C. EFFECTOF ADENINENUCLEOTIDEANALOGS Neither ATP nor ADP inhibited the fructose-2,6-bisphosphataseactivity suggesting that the two catalytic sites were distinct (16, 25). 8-Azido-ATP serves as a substrate for 6-phosphofructo-2-kinase with a K, of about 1 mM (59). Exposure of the enzyme-8-azido-ATP complex to light results in covalent incorporation (0.7 mol/mol of subunit) and 90% loss of kinase activity without loss of fructose-2,6-bisphosphatase.When the native and the first cleavage product of tryptic digestion were photoaffinity-labeled with [ ~ ~ ~ P ] 8 - a z i d o - A the T P radio, label occurred only in the native enzyme (59). Similarly, treatment of the enzyme with 5’-p-fluorosulfonylbenzoyladenosine resulted in inactivation of the kinase activity but had no effect on the bisphosphatase activity (42,53).

D. EFFECT OF HISTIDYLRESIDUEMODIFICATION If liver 6-phosphofructo-2-kinase-fructose-2,6-bisphosphataseis incubated with diethlylpyrocarbonate both the kinase and bisphosphatase are inactivated. However, there is a differential sensitivity of the two activities to this reagent, with the kinase much more sensitive to inactivation than the bisphosphatase (50, 53). While the results of the diethylpyrocarbonate experiments provide circumstantial evidence for involvement of histidine in both reactions, its reactivity with other amino acids necessitates further work to identify the residue(s) modified. Inactivation of the bisphosphatase is consistent with the demonstration of histidine at the active site. Furthermore, the difference in the response of the two activities to the reagent suggests different sites.

IX. Regulation of 6-Phosphofructo-2-KinaseFructose-2,6-Bisphophatase by Low-Molecular-Weight Effectors

The regulation of the 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase by effectors is considered in detail in various sections of this chapter. It is useful to summarize the major features as follows: 6-Phosphofructo-2-Kinase Activity

22

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

1. Product inhibition is observed with ADP (competitive with respect to ATP,

noncompetitive with respect to Fru-6-P) and Fru-2,6-P2 (noncompetitive with respect to both Fru-6-P and ADP) (5,16, 25). 2. Inorganic phosphate activates by increasing the affinity of the enzyme for Fru-6-P (6, 17,46). (See Section VI, C.) 3. a-Glycerol-P is a competitive inhibitor with respect to Fru-6-P (60). 4. It has been reported that AMP activates (46) and that PEP and citrate inhibit (46, 47)partially purified 6-phosphofructo-2-kinase,but none of these effects are observed with a homogeneous preparation of the enzyme (M. R. El-Maghrabi and S . J. Pilkis, unpublished results). Fructose-2,6-Bisphosphatase Activity 1. Substrate inhibition is seen at concentrations of Fru-2,6-P2 above 100 nM (26). 2. The product Fru-6-P is a potent noncompetitive inhibitor (2, 3, 5, 14, 17, 25, 49). 3. Inorganic phosphate and a-glycerol-P are competitive inhibitors at low Fru-2,6-P2 concentrations (26), but both effectors are activators at higher Fru-2,6-P2 concentrations where substrate inhibition is seen (5, 6, 14, 25, 26, 49). 4. It has been reported that GTP, and to a lesser extent ATP, activate partially purified fructose-2,6-bisphosphatase(47, 49), but this observation is not confirmed with a homogeneous preparation of the enzyme (M. R. ElMaghrabi and S. J. Pilkis, unpublished results).

X.

Regulation of 6-Phosphofructo-2-Kinase-Fructose-2,6Bisphosphatase by Phosphorylation-Dephosphorylation

The bifunctional enzyme has been shown to be an excellent in vitro substrate for the CAMP-dependent protein kinase (31), which is the only kinase that has been shown to catalyze significant phosphorylation of the enzyme. Uyeda and co-workers (61) claimed that the enzyme was a substrate for phosphorylase kinase but this has been shown not to be the case (29, 31). A single seryl residue per enzyme subunit is phosphorylated by the CAMP-dependent protein kinase with resulting reciprocal changes in the activities of the enzyme-a decrease in kinase activity and an increase in fructose-2,6-bisphosphataseactivity (Fig. 3). The phosphorylation-induced inactivation of the kinase activity is characterized by a shift in the Fru-6-P concentration curve to the right and by a small inhibitory effect on the V,,, of the enzyme (14, 17,22, 29, 49, 61-63). Phosphorylationinduced activation of the bisphosphatase activity is characterized by an increase in the V,,, but with no change in the affinity for Fru-2,6-P2 (14, 17,31, 49).

23

1. FRU-6-P-FRU- 1,6-P2 SUBSTRATE CYCLE

2

5

1

I

10

20

30

60

Time, min FIG.3. Effect of cyclic AMP-dependent protein kinase-catalyzed phosphorylation on rat liver 6phosphofructo-2-kinase-fructose-2,6-bisphosphatase.(A) The bifunctional enzyme was incubated with protein kinase and [y-32P]ATP-MgZ+and incorporation of 32P into the enzyme monitored. (B) At various times the activities of the enzyme were determined.

The apparent K i for Fru-6-P has been reported to be either unchanged (43) or increased (49) as a result of phosphorylation. The net result of CAMP-dependent kinase-catalyzed phosphorylation is that fructose-2,6-bisphosphatase activity predominates when the enzyme is assayed at submaximal concentrations of substrates leading to greatly decreased net synthesis of Fru-2,6-P2. Phosphorylation of 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase is also subject to “substrate-mediated” regulation (15). The initial rate of CAMPdependent protein kinase-catalyzed phosphorylation of the enzyme in vitro is inhibited by the addition of physiological concentrations of Fru-2,6-P2. No other effectors have been found to affect the rate of phosphorylation. Evidence that this “substrate-mediated’’ regulation of phosphorylation of the enzyme can occur in intact cells has been obtained (64). Although covalent modification has been shown to alter the kinetic parameters of a large number of enzymes (65), it has not been possible to identify the precise

24

PlLKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

step in a reaction pathway that is affected by phosphorylation. Such modification may alter an enzyme’s affinity for substrate or cofactor or it may affect maximal velocity, or both. In the case of fructose-2,6-bisphosphatase,a phosphoenzyme intermediate and the kinetic competence of this phosphoenzyme have been demonstrated (26). The rate of breakdown of the phosphoenzyme intermediate was shown to be the rate-limiting step in phosphoenzyme turnover (26). These discoveries have made it possible to identify what step or steps in the reaction pathway are influenced by phosphorylation. Phosphorylation had no effect on the K,, for Fru-2,6-P2 either in the presence of Pi or a-glycerol-P (31). Van Schaftingen et al. (49)also reported that phosphorylation affected the V,, of the bisphosphatase but had no effect on K,,,. At low substrate concentrations, the phosphorylation-induced activation of fructose-2,6-bisphosphataseis a result of increasing the rate at which E-P breaks down (66).However, it is still uncertain whether phosphorylation accelerates E-P breakdown per se (E-P H20 + E + Pi)or whether it enhances dissociation of Fru-6-P from the E-PeFru 6-P complex (E-P-Fru 6-P + Fru-6-P + E-P). Evidence in support of the E-PvFru-6-P dissociation being the step is the finding that phosphorylation decreased the ability of Fru-6-P to inhibit E-P breakdown (66). The effect of phosphorylation on fructose-2,6-bisphosphataseactivity measured at high substrate concentrations is due to decreased substrate inhibition, perhaps as a result of enhanced Fru-2,6-P2 dissociation. This is supported by the finding that under these conditions, phosphorylation decreases the rate of E-P formation (66). The finding that the bifunctional enzyme is an excellent substrate for CAMPdependent protein kinase in vitro and the lack of phosphorylation of the enzyme by any other protein kinases tested are consistent with a primary role of CAMPdependent protein kinase in regulation of the enzyme in liver (4-7). It is interesting to note that a number of protein kinases that do not possess an absolute substrate specificity, such as liver Ca2 -calmodulin-dependent glycogen synthase kinase and cGMP-dependent protein kinase, did not catalyze phosphorylation of the bifunctional enzyme. This suggests that the phosphorylation site sequence or some other structural factor confers a high degree of specificity. The amino acid sequence surrounding the phosphorylation site in 6-phosphofructo-2kinase-fructose-2,6-bisphosphatasehas been determined to be Val-Leu-GlnArg-Arg-Arg-Gly-Ser-Ser-Ile-Pro-Gln (31). Only the first seryl residue is phosphorylated by CAMP-dependent protein kinase. The amino acid sequence surrounding the phosphorylatable serine in the bifunctional enzyme provides a molecular basis for understanding why the enzyme is one of the best-known protein substrates for the CAMP-dependent protein kinase (Table V). In the bifunctional enzyme the phosphate-accepting serine is separated by one residue from three basic residues N-terminal to it whereas in pyruvate kinase there are only two basic residues separated by a single residue. In contrast, fructose- 1,6-bisphosphatase has only one basic residue N-terminal to

+

+

1.

25

FRU-6-P-FRU- 1,6-P~SUBSTRATE CYCLE TABLE V

AT AMINOACIDSEQUENCES

THE

PHOSPHOKYLATION SITESOF VAKIOUS GLUCONOEGENIC ENZYMES THEIRK, FORPKOIEIN KINASE~'

AND

Substrate 6-PF2-K-Fru-2,6-P2ase, rat liver Pyruvate kinase, rat liver Fructose- 1,6-bisphosphatase rat liver 6-Phosphofructo-I-kinase skeletal muscle rat liver

Sequenceb

Km (pM)

V (unitdmg)

Val-Leu-Gln-Arg-Arg-Arg-Gly-Ser(P)-Ser-lle-Pro-Gln 10

3.0

Arg-Arg-Ala-Ser(P)VaI-Ala-Glu-Leu

39

4.0

&gSer-&g-Pro-Ser(P)-Leu-Pro-Leu-Pro LYS

222

1 .O

His-Ile-Ser-Arg-Lys-Arg-Ser(P)-Gly-Glu-Ala ---

230 600

I .o 0.5

q h e amino acid sequence information for pyruvate kinase, fructose- I ,6-bisphosphatase, and 6-phosphofructo-2-kinase-fructose-2,6-bisphosphdtdsewas taken from Murray el a/. ( 3 I ) and for 6-phosphofructo- I -kinase from (138). "Basic residues are underlined.

the phosphorylatable serine, and there is no residue separating the serine from the three basic residues in 6-phosphofructo-1 -kinase. Both of these features have been shown to make synthetic peptides kinetically poorer substrates for CAMPdependent protein kinase (67-69). Thus it is tempting to speculate that the presence of three, rather than two, arginines in the phosphorylation site sequence make 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatasean even better substrate for the CAMP-dependent protein kinase than pyruvate kinase. Several other proteins contain three or more adjacent arginyl residues NH,-terminal to the site of phosphorylation. They include phosphatase inhibitor- 1 (70, 71), the neuronal phosphoprotein DARPP-32 (72), and protamines (73). Synthetic peptide studies support the speculation that the presence of three adjacent arginyl residues make for a better substrate for phosphorylation by CAMP-dependent protein kinase than do the presence of two basic residues (Table VI). The synthetic dodecapeptide 1 in Table VI has the same sequence as The CAMPthat found in 6-phosphofructo-2-kinase-fructose-2,6-bisphophatase. dependent protein kinase has similar kinetic constants for phosphorylation of the peptide as have been reported for the native bifunctional enzyme and both were phosphorylated on only the first serine (31). Peptide 1 is a better substrate for CAMP-dependent protein kinase than previously studied synthetic peptide substrates (67, 69, 74-83). Under the assay conditions used, it was a better substrate than Kemptide, the peptide modeled after the phosphorylation site sequence in Ltype pyruvate kinase (67). Peptide 2 , like the Kemptide, has two rather than three arginyl residues NH,-terminal to its phosphorylation site and the kinetic con-

26

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI TABLE VI OF

Peptide number 1

2 3 4

5 6

APPARENT KINETIC CONSTANTS OF CATALYTIC SUBUNIT CAMP-DEPENDENT PROTEIN KINASEFOR PHOSPHORYLATION OF SYNTHETIC PEPTIDES Apparent K,, Peptide sequenceo V-L-Q-R-R-R-G-S-S-I-P-Q V-L-Q-A-R-R-G-S-S-I-P-Q V-L-Q-R-R-R-_P-S-S-I-P-Q V-L-Q-R-R-R-G-_T-S-I-P-Q V-L-Q-_A-R-R-G-T-S-I-P-Q V-L-Q-R-R-R-P-1-S-I-P-Q

(W

3.8 11.3 12.5 39 I39 I 18

)

Vm,, ()Lmol/minlmg)

Ratio (V,,dK,,J

13.6 8.4 14.6 4.6 6.8 6.6

3.6 0.7 I .2 0.1 0.05 0.06

("When an amino acid residue of peptide 1 has been substituted by another amino acid, the latter is underlined.

stants for these two peptides are identical. Similar results indicating the importance of three or more arginyl residues for the CAMP-dependent protein kinase have been shown using peptides corresponding to the phosphorylation site in phosphatase inhibitor-1 (79, 80). These results strongly suggest that the primary sequence at the phosphorylation site in 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase is the major determinant that allows the enzyme to be readily phosphorylated by CAMP-dependent protein kinase. The finding that phosphorylation apparently decreases the affinity of the enzyme for Fru-6-P in the kinase reaction but increases the V,,, of the bisphosphatase raises the question of whether the apparent reciprocal changes in the two activities of the enzyme may be due to the increase in V,,,, for the bisphosphatase with no effect on the kinase reaction at all. This would explain the decreased kinase activity seen at low concentrations of Fru-6-P. Further studies are necessary to clarify this point and to elucidate how phosphorylation at a single regulatory seryl residue can affect both activities in a reciprocal manner. The dephosphorylation of liver 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase has also been studied (84). Fractionation of rat liver extracts by anion-exchange chromatography and gel filtration demonstrated that the only protein phosphatases acting on the enzyme were protein phosphatase-2A and -2C. Under the assay conditions used protein phosphatase-2A appeared to be the most powerful phosphatase acting on the enzyme. However, there is no known physiologically relevant regulators of this protein phosphatase. FranGois ef al. (85) claimed that 6-phosphofructo-2-kinase in yeast is affected differently than in rat liver by CAMP-dependent phosphorylation. When glucose is added to yeast in the stationary phase there is a transient increase in cyclic AMP and a persistent increase in Fru-2,6-P2 levels and in 6-phosphofructo-2-

FRU-6-P-FRU- 1,6-P2 SUBSTRATE CYCLE

1.

27

kinase activity. Addition of catalytic subunit of CAMP-dependent protein kinase from beef heart and ATP to a partially purified, but still crude, preparation of the yeast 6-phosphofructo-2-kinasecaused a 10-fold activation of the enzyme, which was characterized by a 4-fold increase in V,,, and a 2-fold decrease in K, for Fru-6-P (85). These results suggest that the glucose-induced elevation in Fru-2,6-P2 results, at least in part, from cyclic AMP-induced activation of the 6phosphofructo-2-kinase. However, these presumed effects of cyclic AMP-dependent phosphorylation will need to be confirmed with homogeneous preparations of the yeast 6-phosphofructo-2-kinase.No information is available on phosphorylation-induced changes on yeast fructose-2,6-bisphosphataseactivity.

XI. 6-Phosphofructo-1-Kinase: Possible Role of Phosphorylation in the Control of Enzyme Activity

A.

LIVER

6-Phosphofructo- 1-kinase has been purified from livers of a number of species (86-98). The rat liver enzyme consists of four apparently identical subunits with a molecular weight of 82,000 (90-92). Like that of heart (99) and muscle (ZOO), it tends to form aggregates with molecular weights of the order of several million (90-92, 101). This aggregation is an equilibrium process influenced by enzyme

concentration, the presence of allosteric effectors, the oxidation-reduction state of sulfhydryl groups, and temperature (8, 9, 91, 92, 102). The aggregation state of 6-phosphofructo-1-kinase may also influence its kinetic behavior. Reinhart and Lardy (92) observed that the rat liver enzyme gave nonlinear rates of activity when it was diluted whereas linear rates were obtained when high concentrations of enzyme were used. Evidence that various ligands, including ATP, ADP, and Fru-6-P, affect the quaternary structure of rat liver 6-phosphofructo- 1-kinase has been reported using enzyme labelled with the fluorescent probe pyrenebutyric acid (103). The liver enzyme exhibits homotropic cooperativity with regard to its substrate Fru-6-P (9, 86, 87, 90-92, 104, 105). Allosteric activators of the enzyme include AMP, ADP, and cyclic AMP, while ATP and citrate are allosteric inhibitors (9, 86, 90-92). The ATP inhibition of the enzyme decreased markedly as the pH increased from 6.5 to 8.0 while citrate potentiated the inhibitory effect of ATP (106, 107). The first suggestion that liver 6-phosphofructo- I -kinase may be regulated by a phosphorylation mechanism was the report that glucagon depressed activity of the enzyme within minutes after administration to rats (108, 109). Subsequently, the same observations were made in isolated liver systems and in hepatocytes (110-114). The inhibition was characterized in crude extracts by a twofold

28

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

increase in the So.5 for Fru-6-P, no change in the maximal activity of the enzyme, and an increased sensitivity of the enzyme to inhibition by ATP (110, 111, 113, 114). Glucagon increased 32P incorporation into the enzyme in hepatocytes (115) and in vivo (112) and cyclic AMP-dependent protein kinase was shown to catalyze the phosphorylation of purified rat liver 6-phosphofructo- 1 -kinase in vitro (91). Thus, it was originally postulated that glucagon caused inhibition of the enzyme by enhancing its phosphorylation (111, 112). However, it was noted that the increase in phosphorylation of the enzyme in hepatocytes induced by increasing concentrations of glucagon did not correlate well with the decrease in enzyme activity (115). Also, partial purification of the enzyme abolished the hormone effect (115) suggesting that changes in enzyme activity were due to changes in the level of an effector of the enzyme. This effector was subsequently identified as Fru-2,6-P2 [for review, see Refs. 1-6)]. The effect of Fru-2,6-P2 on the activity of rat liver 6-phosphofructo- 1-kinase is shown in Fig. 4. In the absence of any effectors, the enzyme exhibits a low affinity and a high degree of positive cooperativity toward its substrate, Fru-6-P (1-3, 6). Fru-2,6-P2 increases the affinity of the enzyme for Fru-6-P but has no effect on the maximum activity of the enzyme (1-3, 6, 23, 116, 117). The K , for Fru-2,6-P2 is about 0.05 pJ4 which makes this sugar diphosphate 50-100 times more effective than Fru-1,6-P2 (2, 23, 118) and 2500 times more effective than glucose 1,6-bisphosphate (2). Fru-2,6-P2 also overcomes the inhibition by high

3.

2.0 F 6 P , mM

4.0

6#0

8,0

ATP, mM

FIG.4. The effect of Fm-2,6-P2 on the kinetic properties of 6-phosphofmcto-1-kinase. (A) Fru-6or 30 pM Fm-2.6-Pz (A). P concentration dependent in the absence ( 0 )and presence of 150 pM (0) (B) ATP inhibition of 6-phosphofmcto-1-kinase in the absence (0)and presence of I a(0) and 5 (W) Fm-2,6-P2.

1.

FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

29

concentrations of ATP (2, 3 , 113,potentiates the activation by AMP (5, 6 , 116, 113, and acts synergistically with AMP to relieve ATP inhibition (117). It has also been reported to protect 6-phosphofructo- 1 -kinase against inactivation by heat (117), low pH (118), or 6-phosphofructo-1-kinasephosphatase (118). Fru-2,6-P2 also activates 6-phosphofructo- 1-kinase from rabbit muscle ( I I7), rat pancreatic islets (119), human erythrocytes (120), Phycomyces blukesleeonus spores (121), and swine kidney (122) in much the same way as it does the rat liver enzyme. Kityuma and Uyeda (123) studied the binding of Fru-2,6-P2 to muscle 6-phosphofructo- 1-kinase and found 1 mol bound/enzyme subunit and the binding exhibited negative cooperativity. They concluded that Fru-2,6-P2 binds to the enzyme at the same allosteric site as does Fru-1,6-P2. The sugar diphosphate also activates ascites tumor and platelet 6-phosphofructo- 1-kinase whereas Fru-1,6-P2 had no effect on these enzymes (124). Yeast 6-phosphofructo-1-kinase is also stimulated by Fru-2,6-P2 (125, 126). This enzyme, like the liver enzyme, exhibits cooperative kinetics with respect to fructose 6-phosphate, is inhibited by ATP, and activated by AMP. Fru-2,6-P2 has been found to increase the binding affinity of the enzyme for AMP (127). In some cell types, phosphate is transferred to fructose 6-phosphate from inorganic pyrophosphate instead of from ATP. Sabularse and Anderson (128) noted that the PP,-fructose 6-phosphate 1-phosphotransferase enzyme from rnung beans was almost completely dependent upon the presence of Fru-2,6-P2 for activity. The effect of 1 pJ4 Fru-2,6-P2 was to decrease the K,,, for Fru-6-P 67-fold and to increase the V,,, 15-fold. The combination of these two effects gave a 500-fold activation of the enzyme at 0.3 mM Fru-6-P. Van Schaftingen et al. (24) have isolated a similar enzyme from potatoes that appears to be 10 times more sensitive to activation by Fru-2,6-P2 than does the mung bean enzyme. The role of phosphorylation in regulating the activity of 6-phosphofructo- 1kinase remains uncertain. Furuya and Uyeda (129) isolated both a high- and a low-phosphate-containing form of rat liver 6-phosphofructo- 1-kinase and reported that the former form was more strongly inhibited by ATP than the latter form (129). It was postulated that the difference in sensitivity to ATP inhibition was due to the presence of Fru-2,6-P2 bound to the low-phosphate form of the enzyme and that phosphorylation affected the affinity of the enzyme for Fru-2,6P, (129). In contrast, Pilkis et ul. (91) reported that there was no change in the kinetic properties of a purified preparation of the enzyme upon phosphorylation in vitro by CAMP-dependent protein kinase. However, they did show that the enzyme was more sensitive to activation by Fru-2,6-P2 after limited proteolysis which removed the carboxyl terminal phosphorylation site (91) and they suggested that the low-phosphate form of the enzyme isolated by Furuya and Uyeda (129) may be a proteolytically modified form of the enzyme. Sakakibara and Uyeda (130) have purified the low- and high-phosphate-containing forms to homogeneity. A comparison of their allosteric properties showed that the high-

30

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

phosphate-containing enzyme was more sensitive to inhibition by ATP, had a higher for Fru-6-P, and was less sensitive to activation by AMP and Fru-2,6-P2 than the low-phosphate-containing form. Both forms could be phosphorylated suggesting that both had phosphorylation sites and were not proteolytically modified. However, the changes observed in the regulatory properties of the enzyme were small. It seems likely that phosphorylation may regulate 6-phosphofructo-1-kinase activity under certain conditions in virro. However, in the case of glucagon- or insulin-induced alterations in liver enzyme activity that were measured in crude extracts, partial purification of the enzyme results in complete disappearance of the hormone effect (6, 113). This suggests that changes in low-molecular-weight effectors are the most important regulating factors.

B.

SKELETAL MUSCLE

A number of laboratories have reported that 6-phosphofructo- 1-kinase from skeletal muscle contains covalently bound phosphate (131-136). The site of in vitro phosphorylation catalyzed by the CAMP-dependent protein kinase is near the carboxyl terminus of the enzyme subunit (137). This phosphorylation site has been sequenced and is His-Ile-Ser-Arg-Lys-Arg-Ser(P)-Gly-Glu-Ala-Thr-Val (138). However, phosphorylation by the CAMP-dependent protein kinase does not cause significant changes in the regulatory properties of muscle 6-phosphofructo-1-kinase (139, 140). For example, Kitajima et al. (140) reported that the phosphoenzyme is more sensitive to ATP inhibition than the dephosphorylated form, but the differences are very small. Similar results have been reported by Foe and Kemp (139). The changes observed do not appear to be physiologically relevant, particularly since hormones that raise cyclic AMP levels, such as epinephrine, enhance glycolysis in muscle and thus would not be expected to cause inhibition at the 6-phosphofructo- I-kinase step. This is an entirely different situation from the liver, where hormones that elevate cyclic AMP inhibit glycolysis and stimulate gluconeogenesis. Hofer et al. (141) have reported that the Ca2+- and phospholipid-dependent protein kinase C from rat brain catalyzes the phosphorylation of rabbit muscle 6phosphofructo- 1-kinase at the same site as the CAMP-dependent protein kinase and at one or more separate sites. Concomitant with this phosphorylation, there is activation of the enzyme which was characterized by a decrease in the K,,, for Fru-6-P. Since protein kinase C has been postulated to be involved in signal transduction from a,receptors, it is possible that this protein kinase may mediate some of these agents’ effects on glycolysis in skeletal muscle. Additional studies are needed to determine whether 6-phosphofructo-1-kinaseis a substrate for protein kinase C in vivo and whether such a regulatory mechanism is physiologically relevant.

1.

C.

31

FRU-6-P-FRU-I ,6-P* SUBSTRATE CYCLE

HEART

The regulation of 6-phosphofructo- 1-kinase in heart initially appeared quite different from that in skeletal muscle. There is a large literature on the adrenergic control of 6-phosphofructo-1-kinaseand glycolysis in heart, and the reader is referred to several excellent reviews (142-145), including one on covalent modification by phosphorylation-dephosphorylation (145). Clark and co-workers [reviewed in Ref. (142)] have summarized a large body of evidence on the regulation of heart 6-phosphofructo- 1-kinase by adrenergic agonists. They found that catecholamines acted by a predominantly a-adrenergic mechanism to activate 6-phosphofructo-1-kinase and glycolysis. P-Adrenergic agonists had similar effects, but only at high concentrations or in the presence of a-adrenergic blocking agents. There was no evidence for a role of phosphorylation or of changes in Fru-2,6-P2 levels in this activation of 6-phosphofructo- 1-kinase (146). Data were obtained suggesting that Ca2 was required for the expression of the activation; however, the mechanism whereby Ca2 effected these changes in 6-phosphofructo- 1-kinase activity remains unknown. A possible explanation for these effects has been provided by a report of Narabayashi et al. (21), who purified 6-phosphofructo-1-kinase from heart perfused with epinephrine and found that it contained 2-fold higher amounts of covalently bound phosphate than did the enzyme from control hearts. This phosphate appeared to be present at a site different from the CAMP-dependent protein kinase phosphorylation site. Purified 6-phosphofructo-1-kinase from epinephrine-treated hearts was less sensitive to ATP inhibition and its apparent K,,, for Fru-6-P and K , for Fru-2,6-P2 were 50% those of the enzyme from control hearts. These results strongly suggest that epinephrine-induced activation of heart 6-phosphofructo-1-kinase results from phosphorylation of the enzyme by an as yet unidentified protein kinase. It is possible that protein kinase C is responsible for this activation of the heart enzyme, but further work will be necessary to establish this mechanism. It was also shown that epinephrine caused a 2-fold increase in Fru-6-P and Fru-2,6-P2 levels in heart (21). Partially purified 6-phosphofructo-2-kinase from epinephrine-treated and control hearts had for Fru-6-P of 4 and 15 pl4. respectively. These results suggest that epinephrine may increase Fru-2,6-P2 levels in heart, at least in part, by a mechanism which involves a covalent modification of 6-phosphofructo-2-kinase. The identity of the presumptive protein kinase involved is unknown. Further experiments, particularly purification of the 6-phosphofructo-2-kinase to homogeneity and characterization of its phosphorylation in vitro by purified protein kinases, are clearly indicated. +

+

D. ASCARIS SUUM 6-Phosphofructo-1-kinase has been purified from the muscle of the nematode parasite Ascaris suum (147). The subunit molecular weight was found to be

32

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

95,000 and the native enzyme molecular weight was 398,000 suggesting that native enzyme is a tetramer. Cyclic AMP-dependent protein kinase catalyzed phosphorylation of this enzyme with a concomitant increase in the activity when the enzyme was assayed at low physiological concentrations of Fru-6-P but not when saturating Fru-6-P was employed (148). The CAMP-dependent protein kinase catalyzed incorporation of about 3 mol 32P/mol of enzyme but the enzyme already contained 3 mol P,/mol. The function of this endogenous phosphate was not investigated. Ascaris mum utilizes a predominately anaerobic carbohydrate metabolism for the production of energy. Glucose and glycogen serve as the sole energy source for the parasite and phosphorylation-induced activation of the 6phosphofructo-I-kinase would permit a coordinate regulation of the enzyme with glycogen metabolizing enzymes. However, the physiological relevance of the phosphorylation by the CAMP-dependent protein kinase is still uncertain and will require further study.

XII. Fructose-l,6-Bisphosphatase: Possible Role of Phosphorylation in Control of Enzyme Activity

A.

LIVER

Most studies on the regulatory and kinetic properties of hepatic fructose- 1,6bisphosphatase have been done with the rabbit and rat liver enzyme (149, 150). This cytosolic enzyme is subject to a multiplicity of controls [see Refs. (11-13, 149) for review] including allosteric inhibition by AMP (151-157) and substrate inhibition by Fru-I ,6-P, (152). Many of the early studies were done with enzyme that had a pH optimum of about 9. It was subsequently shown that this was not the native enzyme, but that it arose from proteolytic cleavage of a small peptide (M,-6000) from the N-terminus of the enzyme subunit during purification (158). The native enzyme had a pH optimum of less than 8, was more sensitive to AMP inhibition, and had a subunit molecular weight of 35,000 instead of 29,000; further evidence suggests that the subunit molecular weight may be even greater than 35,000 (159). Since many of the effectors of 6-phosphofructo- 1-kinase affect the activity of fructose-l,6-bisphosphatasein a reciprocal manner, it is not surprising that Fru-2,6-P, was found to be a potent competitive inhibitor, with a Kiof about 0.5 pill (160). Fructose- 1,6-bisphosphatase displays hyperbolic kinetics with regard to its substrate, Fru-l,6-P2 (157, 161-163). The inhibition by low concentrations of Fru-2,6-P2 also display hyperbolic kinetics with respect to substrate, indicative of competitive inhibition at the active site (160). Higher concentrations of Fru-2,6-P2 resulted in a sigmoidal response to increasing substrate concentrations (164, 165), which suggests that Fru-2,6-P2 may also interact with a site

1.

FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

33

other than the catalytic site. However, studies on the binding of Fru-2,6-P, to fructose-l,6-bisphosphataserevealed that only 1 mol Fru-2,6-P, bound per mol enzyme subunit (166). When the catalytic site of the enzyme was acetylated, Fru-2,6-P, binding was abolished, but when the active site was protected against acetylation by the presence of Fru-1,6-P,, Fru-2,6-P2 was able to bind to the enzyme (166). Fru-2,6-P2 binding exhibited negative cooperativity and was competitive with methyl a- and P-D-fructofuranoside-I ,6-P,, competitive substrate analogs of Fru-1,6-P,. Taken together, these results indicate that Fru-2,6P, binds to the catalytic site, and this conclusion has been confirmed by others (167-1 70) using various kinetic approaches. In contrast to most of these findings, FranGois et al. (171) have argued that Fru-2,6-P2 does not interact at all with the active site but instead binds to a specific allosteric site. The major points in favor of this view are ( a ) the sigmoidal substrate concentration curve in the presence of high concentration of Fru-2,6-P,; (b) potentiation of AMP inhibition by Fru-2,6-P,; and ( c )the similar response of Fru-2,6-P, and AMP inhibition to temperature. Corredia et al. ( I 72) have also reported that under certain conditions Fru-2,6-P, actually can activate fructose-1,6-bisphosphatase, an effect attributed to interaction at an allosteric site, but this effect has not been observed by others (160, 164, 165). In an attempt to resolve the question of where Fru-2,6-P2 binds, a number of groups have studied the mechanism whereby Fru-2,6-P2 potentiates the inhibition of fructose-l,6-bisphosphataseby AMP (166, 167, 173). Binding studies demonstrated that this effect was due to the ability of Fru-2,6-P, to enhance the affinity of the enzyme for AMP (166), and it seems reasonable to postulate that Fru-2,6-P, binding brings about a conformational change in the enzyme that facilitates AMP binding. Compatible with this hypothesis is the finding that Fru-2,6-P, and AMP both induce uv-difference spectra with saturable absorbance maxima at the same wavelengths (166). This suggests that Fru-2,6-P2 binding at the active site can induce a conformational change in the enzyme similar to that induced by AMP at the allosteric site. Studies using NMR and EPR show that the catalytic and AMP sites are in close proximity to one another (13, 173), and this may explain why similar conformational changes are brought about by Fru-2,6-P2 and by AMP. Recently 'H and 31PNMR have shown that the distances between the phospho group of Fru-6-P and enzyme-bound Mn2 and between the 6-phospho groups of Fru-2,6-P, or a-methyh-fructofuranoside-1,6-P2 and enzyme-bound Mn2+ were the same (173). The presence of Fru-2,6-P, caused the proton resonances of AMP to narrow, indicating that Fru-2,6-P2 affects the exchange between AMP and the enzyme. It was concluded that Fru-2,6-P, affected the interaction of AMP with fructose- 1,6-bisphosphatase by interacting with the active site. Meek and Nimmo (174) reported that Fru-2,6-P, protected fructose-I ,6bisphosphatase against partial inactivation by N-ethylmaleimide. The treated +

34

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

enzyme lost the sigmoidal component of the inhibition by Fru-2,6-P2 and the compound was then a simple competitive inhibitor. These workers suggested that Fru-2,6-P2 can bind to both the active site and to a separate allosteric site. This conclusion, however, is not consistent with the finding that only I mol of Fru-2,6-P2 binds per mole of enzyme subunit (166). An important question is whether an analogue of Fru-1,6-P, or Fru-2,6-P2 can bind to the active site of fructose- 1,6-bisphosphatase, exert competitive inhibition, and still potentiate allosteric inhibition at the AMP site. Maryanoff et al. (175) have synthesized a- and P-D-arabinose- I ,5-bisphosphate analogues of Fru-2,6-P2. Arabinose-l,5-P2 is a purely competitive inhibitor of both rat and rabbit liver fructose- 1,6-bisphosphatase and potentiates AMP inhibition in a manner similar to that seen with Fru-2,6-P2 (176). While the results with arabinose- 1,5-P, suggest that interaction with the catalytic site can modulate allosteric interactions, the opposite result has been obtained with 2S-anhydromannitol- 1,6-P, which is a potent competitive inhibitor of the enzyme but does not potentiate AMP inhibition (177). The sigmoidicity of the substrate concentration curve in the presence of Fru-2,6-P2 has been used as the main argument for a separate allosteric site for Fru-2,6-P2, but alternative explanations are possible. For example, Fru-2,6-P2 interaction with fructose- 1,6-bisphosphatase may represent an example of the “ligand exclusion” theory of inhibitor-induced sigmoidal behavior ( I 78, 179). In this case the velocity versus substrate curve is the usual hyperbola in the absence of inhibitor, while in the presence of inhibitor the curve becomes sigmoidal. In this essentially purely competitive situation, where binding of inhibitor to one site prevents the substrate binding to two identical sites, competition can occur because of distortion in substrate binding due to binding of the inhibitor, or mutual steric hinderance, or because the inhibitor binding site may overlap or utilize part of the substrate binding sites. The first suggestion that rat liver fructose- 1,6-bisphosphatase activity may be regulated by a phosphorylation mechanism came from the observations that injection of glucagon (108, 109, 180, 181) or CAMP (108, 109, 180) into rats increased the activity of the enzyme. Consistent with this idea was the observation that 32Pcould be incorporated into the rat liver enzyme in vivo (157) and the demonstration of hormone-stimulated 32P-incorporation into the enzyme in isolated hepatocytes (182). In addition, Riou et al. (157) reported that in vitro phosphorylation of the enzyme by the CAMP-dependent protein kinase resulted in a small increase in the V,,, when the enzyme was assayed in the absence of EDTA. Ekman and Dahlqvist-Edberg (183) confirmed this finding and also reported that phosphorylation decreased the K, for Fru- 1,6-P2. This group has extended this work to show that phosphorylation decreased inhibition of the enzyme by both AMP and Fru-2,6-P2 (184). McGrane et al. (185) have reported that several forms of the enzyme can be detected by isoelectric focussing and

1.

FRU-6-P-FRU-I ,6-P2 SUBSTRATE CYCLE

35

raised the possibility that the different forms of the enzyme may be regulated differently by phosphorylation-dephosphorylation. Despite these observations, the role of phosphorylation in the hormonal regulation of fructose- 1,6-bisphosphatase is uncertain. Even though glucagon stimulated 32P incorporation into the enzyme in isolated hepatocytes no glucagoninduced activity change was observed (182). Furthermore, the concentration of glucagon needed for half-maximal stimulation of 32P-incorporation (1 nM) was more than three times that needed for half-maximal stimulation of gluconeogenesis (0.3nM). Also, there have been no reports of effects of hormones on fructose- 1,6-bisphosphatase activity in hepatocyte extracts or after partial purification of the enzyme. These results suggest that fructose- 1,6-bisphosphatase activity, like that of 6-phosphofructo- 1-kinase, is regulated primarily by hormone-induced changes in the level of Fru-2,6-P2. The in vifro phosphorylation of the rat liver enzyme by the cyclic AMPdependent protein kinase has been well characterized, however. Four moles of phosphate are incorporated per mole of enzyme or 1 mol of phosphate/mol of subunit (157). Fru-1,6-P2, Fru-2,6-P2, nor AMP have any effect on the initial rate of phosphorylation of the enzyme. Fructose-l,6-bisphosphatasewas not as good a substrate for the cyclic AMP-dependent protein kinase as pyruvate kinase or 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase(Table V). The K , for fructose-l,6-bisphosphatasewas 6-fold greater (222 pM) than that for pyruvate kinase (39 pM), while the maximal rate of phosphorylation was about one-third that for pyruvate kinase and for the bifunctional enzyme. These results can be explained by the fact that pyruvate kinase and the bifunctional enzyme contain two and three arginine residues, respectively, on the NH,-terminal side of the phosphorylated serine whereas fructose- 1,6-bisphosphatase contains only one (see Section X). The sequence around the phosphorylated serine in rat liver fructose- 1,6-bisphosphatase has been reported to be either Ser-Arg-Pro-Ser(P)Leu-Pro-Leu-Pro (186) or Ser-Arg-Tyr-Ser(P)-Leu-Pro-Leu-Pro (187). Rittennhouse et al. (188) confirmed the sequence obtained by Pilkis ef al. (186) and identified a second CAMP-dependent phosphorylation site, Arg-Ala-Arg-GluSer(P)-Pro, at the carboxyl terminal region of the subunit. However, they could not detect an effect of phosphorylation on the activity of the enzyme. Both phosphorylation sites of rat liver fructose- 1,6-bisphosphatase are located near the carboxyl terminus of the enzyme (159, 186-188). Hosey and Marcus (144) have noted that among fructose-l,6-bisphosphatases from livers of a number of mammalian species, only the rat enzyme contained a carboxyl-terminal phosphorylation site. Work by El-Dony and MacGregor (189) has suggested that limited proteolysis was not responsible for the absence of the phosphorylation site on the rabbit liver enzyme since immunoprecipitation of in v i m translational products yielded a rat liver enzyme that was larger than the rabbit liver form. The finding of a carboxyl-terminal phosphorylation site only in the rat liver

36

PlLKlS, CLAUS, KOUNTZ, AND EL-MAGHRABI

enzyme casts some doubt on a universal role of hormonal modulation of phosphorylation in the regulation of mammalian liver fructose- 1,6-bisphosphatase activity.

B.

YEAST

There is a growing amount of evidence that implicates phosphorylation-dephosphorylation in the regulation of fructose- 1,6-bisphosphatase from Saccharomyces cerevisiae. This enzyme has been purified to homogeneity from Baker's yeast and consists of a dimer with a subunit molecular weight of 57,000 (190). The specific activity of the yeast enzyme is 46 units/mg which is similar to that of the rat liver enzyme, and it is inhibited by AMP, Fru-2,6-P,, and its substrate Fru-1,6-P,. The inhibition by AMP is noncompetitive and does not exhibit cooperative behavior (176). Addition of glucose to Saccharomyces cerevisiae grown on a gluconeogenic carbon source caused an increase in cAMP and inactivation and proteolytic degradation of fructose-l,6-bisphosphatase(192, 193). Within 1-3 min after glucose addition about 60% of fructose- 1,6-bisphosphatase activity was lost and a concommitant phosphorylation of the enzyme occurred (194, 1 9 3 , suggesting that phosphorylation was responsible for the inactivation. This was supported by the finding that the purified enzyme is phosphorylated in vitro by cAMP dependent protein kinase and that the phosphorylation lowered enzyme activity by 50% when measured with a saturating substrate concentration; I mol of phosphate was incorporated per mol of enzyme or 0.5 mol/mol of subunit and the rate of phosphorylation was greatly stimulated by Fru-2,6-P,. Holzer (196) found that the catabolic inactivation of fructose- 1,6-bisphosphatase occurs as a two-step process. The first step was a rapid, reversible inactivation of the enzyme presumably mediated by CAMP-dependent phosphorylation. The second step is believed to involve proteolytic degradation since the antigenic properties of the enzyme changed. Different results have been obtained with fructose- 1,6-bisphosphatase from Kluyveromyces fragilis (197). This enzyme has an apparent molecular weight of 155,000 and is composed of 4 subunits of 35,000 daltons. The enzyme is also phosphorylated by a CAMP-dependent protein kinase purified from yeast but the rate and extent of phosphorylation is greatly dependent on the presence of the inhibitors AMP and Fru-2,6-P,. Phosphorylation had no effect on enzymic activity assayed with saturating substrate concentrations. However, changes in kinetic parameters such as K,,, for Fru- 1,6-P, or apparent K j for AMP or Fru-2,6P, were not investigated. These workers suggested that the rapid regulation of fructose- 1,6-bisphosphatase seen in this yeast following glucose addition is controlled primarily by changes in levels of low-molecular-weight effectors.

37

I . FRU-6-P-FRU- I ,6-P2 SUBSTRATE CYCLE

XIII. Role of 6-Phosphofructo-2-Kinase-Fructose-2,6Bisphosphatase in the Hormonal Control of Hepatic Gluconeogenesis and Glycolysis

The bifunctional enzyme is an excellent substrate in vitro for the CAMPdependent protein kinase, suggesting that phosphorylation of the enzyme also occurs in vivo. Glucagon, which affects hepatic metabolism by activation of the cyclic AMP-dependent protein kinase and the subsequent increase in phosphorylation of specific enzyme proteins, when added to isolated hepatocytes caused reciprocal changes in 6-phosphofructo-2-kinase and fructose-2,6bisphosphatase activities (1, 4-6, 14, 22, 29, 41, 46, 49, 61, 198). This effect was characterized by a decrease in affinity for fructose 6-phosphate in the kinase reaction and by increase in both V,,, and affinity for Fru-2,6-P2 in the bisphosphatase reaction (4-6, 14,22,49, 199). These hormone-induced changes were similar to those observed when the purified enzyme was phosphorylated in vitro by the cyclic AMP-dependent protein kinase. Direct evidence for phosphorylation of the enzyme in cells came from Garrison and Wagner (30) who measured 32Pincorporation into proteins in intact hepatocytes and then identified 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase after separation by twodimensional gel electrophoresis. The addition of 10 nM glucagon to the hepatocytes enhanced phosphorylation of 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase by 15-fold. Epinephrine addition to hepatocytes has also been reported to inhibit 6-phosphofructo-2-kinase activity and to increase fructose-2,6-bisphosphatase activity via P-adrenergic receptor-mediated changes in cAMP (6, 29, 200). Insulin counteracts the effects of both glucagon and epinephrine and affects the activities of 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase in a manner that is also consistent with the effects of the hormone on cAMP levels (5, 29, 200, 201). Epinephrine has also been reported to cause an inhibition of 6-phosphofructo2kinase activity and an increase in fructose-2,6-bisphosphatase activity by an aadrenergic mechanism that involves Ca2 -induced activation of phosphorylase kinase rather than changes in cAMP and CAMP-dependent protein kinase (61). However, several lines of evidence indicate that an a-adrenergic mechanism is not involved in regulation of this enzyme in liver. First, Hue et al. (202) reported that the a-adrenergic agonist phenylephrine had no effect on the enzyme. Second, Garrison and Wagner (30) reported that vasopressin and angiotensin, which act by a Ca2+-linked, CAMP-independent mechanism, had no effect. Addition of phorbol esters or calcium ionophore to hepatocytes also had no effect on phosphorylation of the bifunctional enzyme. Third, purified phosphorylase kinase was not able to phosphorylate purified 6-phosphofructo-2-kinase-fructnse-2,6-bisphosphatase (29). It may be concluded that this enzyme is affected +

38

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

only by cyclic AMP-linked hormones and that Ca2 -linked phosphorylation by phosphorylase kinase, protein kinase C, or Ca2 -calmodulin-dependent protein kinase is not involved. Consistent with this conclusion is the inability to demonstrate significant phosphorylation of the enzyme in vitro by any protein kinase other than the CAMP-dependent variety. Regulation of 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase by phosphorylation plays an important role in the regulation of glycol ysis and gluconeogenesis in mammalian liver. A summary of these effects is shown in Fig. 5A. Hormones that stimulate cyclic AMP production cause phosphorylation of the bifunctional enzyme. This results in a decrease in Fru-2,6-P2 levels due to inhibition of the kinase reaction and activation of the bisphosphatase reaction. The decrease in Fru-2,6-P2 in turn leads to decreased allosteric activation of 6phosphofructo-1-kinase and decreased inhibition of fructose- 1,6-bisphosphatase. These changes in enzyme activity may be further amplified by concomitant phosphorylation of 6-phosphofructo- 1-kinase and fructose- 1,6-bisphosphatase. The decrease in 6-phosphofructo- I-kinase activity and the activation of fructose-1,6-bisphoshatase cause a reduction in the level of Fru-1,6-P, which is a potent allosteric activator of pyruvate kinase. Inhibition of this enzyme occurs not only by a decrease in Fru-l,6-P2 levels but also by CAMP-dependent phosphorylation of a specific seryl residue (see L. Engstrom et al. Chapter 2, this volume). Inhibition of pyruvate kinase plays a major role in the stimulation of gluconeogenesis and the inhibition of glycolysis by hormones that elevate cyclic AMP levels. Inhibition of pyruvate kinase by hormones that act by cyclic AMPindependent mechanisms also occurs, and this appears to be mediated by Ca2 calmodulin-dependent protein kinase catalyzed phosphorylation of a specific threonyl residue in addition to the seryl residue (203). In contrast, the ability of insulin to counteract hormones that elevate cyclic AMP results in dephosphorylation of 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase and pyruvate kinase (Fig. 5B). Insulin also causes a reversal of CAMP-independent protein kinase-mediated phosphorylation of pyruvate kinase (6),but the mechanism of this effect is unknown (200). Activation of the kinase reaction and inhibition of the bisphosphatase reaction results in an increase in Fru-2,6-P,. This elevation, in turn, activates 6-phosphofructo-I-kinaseand inhibits fructose-l,6 bisphosphatase and thus leads to an increase in Fru- 1,6-P, levels. The increase in the level of this allosteric activator, along with the concomitant dephosphorylation of pyruvate kinase, significantly increases the activity of pyruvate kinase and thus promotes glycolysis and inhibits gluconeogenesis. Regulation of the Fru-6-P-Fru- 1,6-P2 substrate cycle by Fru-2,6-P,, besides controlling glycolysis and gluconeogenesis, may be significant during the early phase of glycogen deposition from glucose in starved animals. When a starved rat, which has low levels of hepatic Fru-2,6-P,, is given a glucose load, glycogen +

+

+

1.

FRU-6-P-FRU-1,6-P2 SUBSTRATE CYCLE

39

LAC

+LAC

FIG.5 . Regulation of the hepatic glycolytic-gluconegenic pathway by phosphorylation reactions. (A) Elevation of cyclic AMP levels leads to phosphorylation of pyruvate kinase, 6-phosphofructo-lkinase, fructose-I ,6-bisphosphatase (in rat liver). and 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase on seryl residues (as indicated by *). Pyruvate kinase can also be phosphorylated by CaZ+, calmodulin-dependent protein kinase on a threonyl residue (as indicated +). These phosphorylations result in inhibition of glycolytic enzyme activities (pyuruvate kinase, 6-phosphofructo-2-kinase, and, at least indirectly via the decrease in Fru-2,6-P2 levels, 6-phosphofructo-1 -kinase) and to activation of enzymes favoring gluconeogenesis (fructose-2,6-bisphosphatase and fructose-I ,6-bisphosphatase). The final result is enhanced lactate to glucose flux. (B) In states where cyclic AMP is low (e.g., with high insulin to glucagon ratios, the phosphorylations in A are all reversed leading to enhanced glycolysis).

40

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

is deposited in the liver but with little change in the level of Fru-2,6-P2. From studies with isolated liver systems, glucose loading would be expected to elevate hepatic Fru-2,6-P2 and to enhance glycolytic flux. However, studies have indicated that the majority of glycogen synthesis occurs by an indirect route whereby glucose is first metabolized to 3-carbon precursors in the periphery or in the liver itself. These precursors then traverse the gluconeogenic pathway before being converted to glycogen (204, 205). In this case Fru-2,6-P2 levels would be expected to remain low in order to promote gluconeogenic flux but the mechanism responsible for the continued low levels is unknown (205). In summary, the steady-state level of Fru-2,6-P2 is controlled by the activity of the multifunctional catalyst 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase. It appears reasonable to postulate that regulation of this unique bifunctional enzyme and of pyruvate kinase by covalent modification represent the most significant sites of the acute action of glucagon, insulin, and P-adrenergic agonists in the glycolytic and gluconeogenic pathway.

XIV.

Summary and Overview

Only in a few instances has phosphorylation-dephosphorylation been shown to regulate an enzyme involved in the interconversion between Fru 6-P and Fru 1,6-P, in a physiologically relevant way. The best example is regulation of (6, 7). The phoshepatic 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase phorylation of this enzyme by the cyclic AMP-dependent protein kinase with resulting changes in the enzyme activities has been well characterized in vitro using purified preparations. Similar reciprocal changes in the enzyme activities have been demonstrated both in vivo and in isolated liver systems in response to elevated levels of cyclic AMP. Concomitant with these changes were the predicted modulations of glycolytic and gluconeogenic flux in intact cells (see Figs. 5A and 5B).There is also preliminary evidence that 6-phosphofructo-2-kinase in yeast (85) and heart (21) are regulated by phosphorylation. However, in neither case has the 6-phosphofructo-2-kinase been purified to homogeneity and its regulatory properties and in vitro phosphorylation studied in detail. These early results suggest that with regard to regulation by phosphorylation the enzymes in yeast and heart are different from that found in liver. In general, we know very little about the properties of 6-phosphofructo-2-kinase and fructose-2,6-bisphosphatase in extrahepatic tissues. It has yet to be clearly shown whether or not the enzyme is bifunctional in extrahepatic tissues and/or whether other enzyme forms are present. While mammalian 6-phosphofructo- 1-kinase from heart and skeletal muscle have been shown to contain covalently bound phosphate in vivo (131-134) and to be substrates for the cyclic AMP-dependent protein kinase in vitro (137-138),

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no convincing changes in the regulatory properties of the enzyme as a result of such phosphorylation have been demonstrated. It has been reported that the muscle enzyme from Ascaris mum is activated by phosphorylation catalyzed by the cyclic AMP-dependent protein kinase (148). The report that both phosphorylation by protein kinase C (141) and epinephrine addition to heart (21) cause activation of 6-phosphofructo- 1-kinase is intriguing and deserves further study. Even less certain is the effect of phosphorylation on mammalian liver fructose 1,6-bisphosphatase since only the rat liver enzyme appears to have a carboxyl terminus phosphorylation site (159). Interestingly, in Saccharomyces cerevisiae cyclic AMP-dependent phosphorylation of fructose-I ,6-bisphosphatase appears to play a role in catabolite repression leading to a decrease in enzyme activity and acting as a signal for proteolytic degradation (196). However, in Kluyveromyces fragilis no effects of phosphorylation on the fructose- 1,6-bisphosphatase have been observed (197). It is certain that in future years further investigation will continue to provide new information on the role of phosphorylation in modulating activities of the enzymes at this key switch point for glycolysis and gluconeogenesis.

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58. Sakakibara, R., Kitajima, S., Hartrnan, F. C., and Uyeda, K. (1984). JBC 259, 14023. 59. Sakakibara, R., Kitajima, S., and Uyeda, K. (1984). JBC 259, 8366. 60. Claus, T. H., Schlumpf, J. R., El-Maghrabi, M. R., and Pilkis, S. J. (1982). JBC 257,7541. 61. Furuya, E., Yokoyama, M., and Uyeda, K. (1982). PNAS 79, 325. 62. Van Schaftingen, E., Davies, D. R., and Hers, H.-G. (1981). BBRC 103, 362. 63. Yokoyama, M., Furuya, E., and Uyeda, K. (1982). BBRC 105, 264. 64. Bartrons, V. E. R., Van Schaftingen, E., and Hers, H . G . BJ 222, 511 1. 65. Cohen, P. (1982). Nature (London) 296, 613. 66. Stewart, H. B., El-Maghrabi, M. R., and Pilkis, S. J. (1986). JBC 261, 8793. 67. Kemp, B. E., Graves, D. J., Benjamini, E., and Krebs, E. G.(1977). JBC 252, 4888. 68. Glass, D. B., and Krebs, E. G. (1980). Annu. Rev. Pharmacol. Toxicol. 20, 363. 69. Glass, D. B., and Krebs, E. G. (1979). JBC 254, 9728. 70. Cohen, P., Rylatt, D. B., and Nimmo, G.A. (1977). FEBS Left. 76, 182. 71. Aitken, A., Bilham, T., and Cohen, P. (1982). EJB 126, 235. 72. Hemmings, H. C., Williams, K. R., Konigsberg, W. H., and Greengard, P. (1984). JBC 259, 14486. 73. Shenolikar, S., and Cohen, P. (1978). FEES Letf. 86, 92. 74. Glass, D. B., and Krebs, E. G.(1982). JBC 257, 1196. 75. Daile, P., Camegie, R. R., and Young, J. D. (1975). Narure (London) 25, 416. Ragnarsson, U.,Humble, E., Berglund, L., and Engstrom, L. (1976). BBRC 76. Zetterqvist, 0.. 70, 696. 77. Pomerantz, A. H., Allfrey, V. G.,Merrifield, R. B., and Johnson, E. M. (1977). PNAS 74, 4261. 78. Feramisco, J. R., Kemp, B. E., and Krebs, E. G. (1979). JBC 254, 6987. 79. Kemp, B. E., Rae, J. D., Minasian, E., and Leach, S. I . (1979). Pept., Srrucr. Biol. Funct., Proc. Am. Pepr. Symp., 6th. 1979 p. 169. 80. Chessa, G.,Borin, G.,Marchiori, F., Meggio, F., Brunati, A. M., and Pinna, L. A. (1983). EJB 135, 609. 81. Meggio, F., Fessa, G.,Borin, G.,Pinna, L. A., and Marchiori, F. (1981). BBA 662, 94. 82. Zetterqvist, O., and Ragnarsson, U. (1982). FEES Letr. 139, 287. 83. Glass, D. B., and May, J. M. (1984). Collagen Relat. Res.: Clin. Exp. 4, 63. 84. Pelleh, S., Cohen, P., Fisher, M. J., Pogson, C., El-Maghrabi, M. R., and Pilkis, S. J. (1984). EJB 145, 39. 85. Francois, J., Van Schaftingen, E., and Hers, H.-G. (1985). EJB 145, 187. 86. Massey, T. H., and Deal, W. C., Jr. (1973). JBC 248, 56. 87. Massey, T. H., and Deal, W. C., Jr. (1975). “Methods in Enzymology,” Vol. 42, Part C, p. 99. 88. Dunaway, G. A., Jr., and Weber, G. (1974). ABB 162, 620. 89. Kasten, T. P., Naqui, D., Kruep, D., and Dunaway, G.A. (1983). BBRC 111, 462. 90. Brand, I. A., and Soling, H.-D. (1974). JBC 249, 7824. 91. Pilkis, S. J., El-Maghrabi, M. R., and Claus, T.H. (1982). ABB 215, 379. 92. Reinhart, G.D., and Lardy, H.A. (1980). Biochemistry 19, 1491. 93. Brock, D. J. H. (1969). BJ 113, 235. 94. Kono, N., and Uyeda, K. (1971). BBRC 42, 1095. 95. Kono, N.,and Uyeda, K. (1973). JBC 248, 8592. 96. Ramaiah, A., and Tejwani, G. A. (1970). BBRC 39. 1149. 97. Kemp, R. G. (1971). JBC 246, 245. 98. Kemp, R. G. (1975). “Methods in Enzymology,” Vol. 42, Part C, p. 67. 99. Mansour, T. E. (1965). JBC 240, 2165. 100. Paetkau, V., Younathan, E. S., and Lardy, H. A. (1968). JMB 33, 721.

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PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI Trujillo, J. L., and Deal, W. C., Jr. (1977). Biochemistry 16, 3098. Bloxharn, D. P., and Lardy, H. A. (1973). “The Enzymes,” 3rd ed., Vol. 8, p. 229. Reinhart, G. D. (1983). JBC 258, 10827. Claus, T. H., and Pilkis, S. J. (1981). In “Biochemical Actions of Hormones” (G. Litwack, ed.), Vol. 8, p. 209. Academic Press, New York. Tsai, M. Y., and Kernp, R. JBC 248, 785. Passonneau, J. V., and Lowry, 0. H. (1964). Adv. Enzyme Regul. 2, 265. Underwood, A. H., and Newsholme, E. A. (1965). BJ 95, 868. Taunton, 0. D., Stifel, F. B., Greene, H. L., and Herman, R. H. (1972). BBRC 48, 1663. Taunton, 0. D., Stifel, F. B., Greene, H. L., and Herman, R. H. (1974). JBC 249, 7228. Pilkis, S. J., Schlumpf, J. R., Pilkis, J., and Claus, T. H. (1979). BBRC 88, 960. Castano, J. G., Nieto, A., and Feliu, J. E. (1979). JBC 254, 5576. Kagirnoto, T., and Uyeda, K. (1979). JBC 254, 5584. Claus, T. H., Schlumpf, J. R., El-Maghrabi, M. R., Pilkis, J., and Pilkis, S. J. (1980). PNAS 77, 6501. Nieto, A , , and Castano, J. G. (1980). BJ 186, 953. Claus, T. H., Schlumpf, J. R., El-Maghrabi, M. R.,Pilkis, J., and Pilkis, S. J. (1980). PNAS 77, 6501. Van Schaftingen, E., Jett, M.-F., Hue, L., and Hers, H.-G. (1981). PNAS 78, 3483. Uyeda, K., Furuya, E., and Luby, L. (1981). JBC 256, 8394. Soling, H.-D., Kuduz, J., and Brand, I. A. (1981). FEBS Lett. 130, 390. Malaise, W. J., Malaise-Lagae, F., Sener, A,, Van Schaftingen, E., and Hers, H.-G. (1981). FEES Lett. 125, 217. Heylen, A., Van Schaftingen, E., and Hers, H.-G. (1982). FEBS Lett. 143, 141. Van Loere, A., Van Schaftingen, E., and Hers, H.-G. (1983). PNAS 80, 6601. Muniyappa, K., Leibach, F. H., and Mendicino, J. (1983). Life Sci. 32, 271. Kityuma, S . , and Uyeda, K. (1983). JBC 258, 7352. Bosca, L., Aragon, J. J., and Sols, A. (1982). BBRC 106, 486. Bartrons, R., Van Schaftingen, E., Vissers, S., and Hers, H.-G. (1982). FEBS Lett. 143, 137. Nissler, K., Otto, A,, Schellenberger, W., and Hofmann, E. (1983). BBRC 111, 294. Kessler, R., Nissler, K., Schellenberger, W., and Hofrnann, E. (1982). BBRC 107, 506. Sabularse, D. C., and Anderson, R. L. (1981). BBRC 103, 848. Furuya, E., and Uyeda, K. (1980). JBC 255, 11656. Sakakibara, R., and Uyeda, K. (1983). JBC 258, 8656. Hofer, H. W . , and Furst, M. (1976). FEBS Lett. 62, 118. Hussey, C. R., Liddle, P. E., Ardron, D., and Kellet, G. L. (1977). EJB 80, 497. Riquelme, P. T., Fox, R. W., and Kemp, R. G. (1978). BBRC 81, 864. Uyeda, K., Miyatake, A., Luby, L. J., and Richards. E. G. (1978). JBC 253, 8319. Hofer, H. W . , and Sorensen-Ziganke, B. (1979). FEBS Lett. 90, 199. Krystek, E., and Hofer, H. W. (1981). BBRC 99, 1138. Riquelme, P. T., Hosey, M. M., Marcus, F., and Kemp, R. G. (1978). BBRC 85, 1480. Kernp, R. G., Foe, L. G., Latshaw, S. P., Poorman, R. A,, and Heinrikson, R. I. (1981).JBC 256, 7282. Foe, L. G., and Kernp, R. G. (1982). JBC 257, 6368. Kitajima, S . , Sakakibara, R., and Uyeda, K. (1983). JBC 258, 13292. Hofer, H. W . , Schlatter, S., and Graefe, M. (1985). BBRC 129, 892. Clark, M. G., and Patten, G. S. (1984). Curr. Top. Cell. Regul. 23, 127. Hofmann, E. (1976). Rev. Physiol. Biochem. Pharmacol. 7, 1. Mansour, T. E., Choate, G., and Weng, L. (1979). In “Modulation of Protein Function” (D. E. Atkinson and C. F. Fox, eds.), Vol. 13, p. 1. Academic Press, New York.

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145. Soling, H.-D., and Brand, I. A. (1981). Curr. Top. Cell. Regul. 20, 107. 146. Clark, M. G., Filsell, 0. H., and Patten, G. S. (1982). JBC 257, 271. 147. Starling, J. A., Allen, B. L., Kaeini, M. R., Payne, D. M., Blytt, H. J., and Hofer, H. W. (1982). JBC 257, 3795. 148. Hofer, H. W., Alley, B. L., Kaeini, M. R., and Harris, D. G. (1982). JBC 257, 3807. 149. Horecker, B. L., Melloni, E., and Pontremoli, S. (1975). Adv. Enzymol. 42, 193. 150. Pilkis, S. J., Park, C. R., and Claus, T. H. (1978). Vitam. Horm. (N.Y.) 36, 383. 151. Taketa, K.,and Pogell, B. M. (1963). BBRC 12, 229. 152. Taketa, K., and Pogell, B. M. (1965). JBC 240, 651. 153. Underwood, A. H., and Newsholme, E. A. (1965). BJ 95, 767. 154. Datta, A. G., Abrams, B., Sasaki, T., van den Berg, J. W. O., Pontremoli, S . , and Horecker, B. L. (1974). ABB 165, 641. 155. Nimmo, H. G., and Tipton, K. F. (1975). BJ 145, 323. 156. Tejwani, G. A., Pedrosa, F. O., Pontremoli, S., and Horecker, B. L. (1976). ABB 17, 253. 157. Riou, J. P., Claus, T. H., Flockhart, D. A,, Corbin, J. D., and Pilkis, S. J. (1977). PNAS 74, 4615. 158. Traniello, S., Pontremoli, S., Tashima, Y., and Horecker, B. L. (1971). ABB 146, 161. 159. Hosey, M. M., and Marcus, F. (1981). PNAS 78, 91. 160. Pilkis, S. J., El-Maghrabi, M. R., Pilkis, J., and Claus, T. H. (1981). JBC 256, 3619. 161. Pontremoli, S., Grazi, E., and Accorsi, A. (1968). Biochemistry 7, 3628. 162. Samgadharan, M. G., Watanabe, A., and Pogell, B. M. (1969). Biochemistry 8, 1411. 163. Tejwani, G. A. (1983). Adv. Enzymol. Refat. Areas Mol. Biol. 54, 121. 164. Pilkis, S. J., El-Maghrabi, M. R., McGrane, M., and Pilkis, J. (1981). JBC 256, 11489. 165. Van Schaftingen, E., and Hers, H.-G. (1981). PNAS 78, 2861. 166. McGrane, M.M., El-Maghrabi, M. R., and Pilkis, S. J. (1983). JBC 258, 10445. 167. Gottsschalk, M. E., Chatterjee, T., Edelstein, I., and Marcus, F. (1982). JBC 257, 8016. 168. Pontremoli, S . , Melloni, E., Michetti, F., Salamino, F., Sparatore, B., and Horecker, B. L. (1982). ABB 218, 609. 169. Ganson, N. J., and Fromm, H. J. (1982). BBRC 108, 233. 170. Marcus, F., Edelstein, I., and Rittenhouse, J. (1984). BBRC 119, 1103. 171. FranGois, J., Van Schaftingen, E., and Hers, H.-G. (1983). EJB 134, 269. 172. Corredia, C., Bosca, L., and Sols, D. L. (1984). FEES Lett. 167, 199. 173. Ganson, N. J., and Fromm, H. (1985). JBC 260, 2837. 174. Meek, D. W., and Nimmo, H. G. (1983). FEES Lett. 160, 106. 175. Maryanoff, B. E., Reitz, D. B., Tutwiler, G. F., Benkovic, S. J., Benkovic, P. A., and Pilkis, S. J. (1984). JACS 106, 7851. 176. Pilkis, S. J., McGrane, M. M., Kountz, P. D., El-Maghrabi, M. R., Pilkis, J., Maryanoff, B. E., Reitz, A. B., and Benkovic, S. J. (1986). BBRC 138, 159. 177. Riquelme, P. T., Wamette-Hammond, M. E., Kneer, N., and Lardy, H. A. (1984). JBC 259, 5115. 178. Marcus, F., Edelstein, I., and Rittenhouse, J. (1984). BBRC 119, 1103. 179. Fisher, H. F., Gates, R. E., and Cross, D. G. (1970). Nature (London) 228, 247. 180. Chatterjee, T., and Datta, A. G. (1978). BBRC 84, 950. 181. Morikofer-Zwez, S., Stoecklin, F. B., and Walter, P. (1981). BBRC 101, 104. 182. Claus, T. H., Schlumpf, J. R., El-Maghrabi, M. R., McGrane, M., and Pilkis, S. J. (1981). BBRC 100, 716. 183. Ekman, P., and Dahlqvist-Edberg, U. (1981). BEA 662, 265. 184. Ekdahl, K. N., and Ekman, P. (1984). FEES Lett. 167, 203. 185. McGrane, M., El-Maghrabi, M. R., and Pilkis, S. J. (1983). JBC 258, 10445.

46

PILKIS, CLAUS, KOUNTZ, AND EL-MAGHRABI

186. Pilkis, S. J., El-Maghrabi, M. R., Claus, T. H., Tager, H. S., Steiner, D. E., Keim, P., and Heinrikson, R. (1980). JBC 255, 2770. 187. Humble, E., Dahlqvist-Edberg, U., Ekman, P., Netzel, R., Ragnarsson, U., and Engstriim, L. (1979). BBRC 90, 1064. 188. Rittenhouse, J., Chatterjee, T., Marcus, F., Reardon, I., and Heinrikson, R. (1983). JBC 258, 7648. 189. El-Dony, H. A., and MacGregor, J. S. (1982). BBRC 107, 1384. 190. Nada, T., Hoffschulte, H., and Holzer, H. (1984). JBC 259, 7191. 191. Gancedo, C., Salas, M. L., Giner, A., and Sols, A. (1965). BBRC 20, 15. 192. Holzer, H. (1976). TIES 1, 178. 193. Funayama, S., Malano, J., and Gancedo, C. (1979). AEB 197, 170. 194. Muller, D., and Holzer, H. (1981). BBRC 103, 926. 195. Mazon, M. J., Gancedo, J. M., and Gancedo, C. (1982). JBC 257, 1128. 196. Holzer, H. (1983). In “Enzyme Regulation by Reversible Phosphorylation-Further Advances” (P. Cohen, ed.) pp. 143-154. Elsevier, Amsterdam. 197. Toyoda, K.,and Sy, J. (1984). JBC 259, 8718. 198. Richards, C. S., Yokoyoma, M., Furuja, E., and Uyeda, K. (1982). BBRC 104, 1073. 199. Yokoyoma, M., Furuya, E., and Uyeda, K. (1982). BBRC 105, 204. 200. Pilkis, S. J., El-Maghrabi, M. R., and Claus, T. H. (1986). In “Symposium on the Mechanism of Action of Insulin,’’ p. 305. Elsevier, Amsterdam. 201. Richards, C. S., and Uyeda, K. (1980). BBRC 97, 1535. 202. Hue, L., Blackmore, P., and Exton, J. H. (1981). JBC 256, 8900. 203. Schworer, C., El-Maghrabi, M. R., Pilkis, S. J., and Soderling, T. R. (1986). JBC 260,13018. 204. Katz, J., and McGany, J. D. (1984). J . Clin. Invest. 74, 1901. 205. Pilkis, S. J., Regen, D. M., Claus, T. H., and Cherrington, A. D. (1985). BioEssuys 2, 273.

Pyruvate Kinase L. ENGSTROM P. EKMAN E. HUMBLE 0. ZETTERQVIST Institute of Medical and Physiological Chemistry University of Uppsala Uppsala, Sweden

I. Introduction .............. 11. Influence of Phosphorylation on the Kinetic Properties of Liver Pyruvate Kinase ................................ 111. Influence of Phosphorylation o Kinase to Proteolytic Enzymes ................. IV. The Reaction of Cyclic AMP-D Pyruvate Kinase as Substrate ...................................... V. Dephosphorylation of Liver Pyruvate Kinase with Phosphoprotein Phosphatases ..................... Intact Cells

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

A. Kidney Enzyme, Type L

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

55

59

65

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

D. Pyruvate Kinase in Chicken Liver References ....................................

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

70 72

I. Introduction Pyruvate kinase catalyzes the final reaction in glycolysis in which pyruvate and ATP are formed from phosphoenolpyruvate (PEP) and ADP. During 47 THE ENZYMES,Vol. XVIII Copyright 8 1987 by Academic Press, Inc. All rights of reproduction in any form reserved.

48

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0,ZETTERQVIST

gluconeogenesis PEP is synthesized from pyruvate via oxaloacetate in two reactions catalyzed by pyruvate carboxylase and PEP carboxykinase, respectively. In this way one of the three substrate cycles of the glycolytic and gluconeogenetic pathways is formed. The activity of the glycolytic and gluconeogenetic pathways in gluconeogenetic tissues (i.e., mainly in the liver and kidney) ( I ) ,seems to be regulated preferentially by control of the enzyme reactions of the substrate cycles, especially the fructose 6-phosphate-fructose I ,6-diphosphate (FDP) and the pyruvate-PEP cycles (2). This is brought about by hormones and diet, both of which affect the concentrations of enzymes, substrates, and effectors. In addition, hormones and metabolites may influence the regulatory phosphorylation of enzymes. Pyruvate carboxylase is essentially an intramitochondrial enzyme, whereas pyruvate kinase and most of the PEP carboxykinase activity are generally present in the cytosol (1, 2). Therefore, regulation of the pyruvate-PEP cycle may also include effects on transport of pyruvate to the mitochondria, conversion of oxaloacetate to malate or aspartate within the mitochondria, their transport to the cytosolic compartment, and reconversion back to oxaloacetate. Four different isozymes of pyruvate kinase are present in mammalian tissues (Table I) (3-5).They are all tetrameric molecules of similar size. The M , and M, types are very similar with regard to their amino acid sequence and may be coded by the same gene (6).The same relationships exist between the L and R types (7). However, there seem to be separate messenger RNA molecules for each isozyme (8). In glycolytic tissues, such as muscle and brain, there is no obvious need for regulation of the pyruvate kinase reaction. The M, enzyme that is present in these tissues is the least sophisticated with regard to its regulation ( 3 ) and is apparently not subject to allosteric control. However, in gluconeogenetic organs there is a pronounced need to regulate the pyruvate kinase activity. Hepatocytes contain only the L type of the enzyme (9, 10), which is also present as a minor component in the kidney (11, 12) and the small intestine (11, 13). The nonparenchymal liver cells contain the M, enzyme (9, 10). The L and R isozymes are the most complicated types with regard to regulation. The concentration of the L isozyme in the liver is increased by insulin and a carbohydrate-rich diet and is decreased by fasting and in diabetes (14-16). In rats the activity of the enzyme is higher in the perivenous zone than in the periportal zone of the liver lobuli (17). Evidence has been obtained that indicates that the concentration of liver pyruvate kinase is determined preferentially by its rate of synthesis, and that this in turn is regulated by the rate of synthesis of the messenger RNA of the enzyme ( 5 , 8, 18). It has been claimed that insulin, corticosteroids and carbohydrate are important for maintenance of the pyruvate kinase in hepatocytes in tissue cultures (19, 20). Fructose is more efficient than glucose in inducing the synthesis of

TABLE I PROPERTIES OF

ISOZYMES OF

PYRUVATE KtNASE

IN

RAT TISSUESu

Characteristic Tissue distribution

Liver, kidney, small intestine

Erythrocytes

Muscle, brain

Subunit molecular weightc Kinetics with regard to PEP Activation by FDP Inhibition by ATP Inhibition by alanine

58,500 Sigmoidal Yes Yes Yes

62,000 Sigmoidal Yes Yes Yes

59,000

OData are from Ref. (3) except for the R isozyme and molecular weights. bData are from Ref. ( 4 ) and refer to the human enzyme. (‘Data are from Ref. (5).

Hyperbolic No Yes No

Fetal tissues, and most adult tissues (e.g. kidney and fat cells) 60,000 Sigmoidal Yes Yes Yes

50

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND O.ZE7TERQVIST

pyruvate kinase in diabetic rats (5). A new steady-state concentration of pyruvate kinase in the tissues is not reached until after several days, since the rate of degradation of the enzyme is fairly slow, with a t , , , of about 45-75 h (21, 22). Pyruvate kinase type L exhibits sigmoidal kinetics with regard to its substrate PEP. It is allosterically activated by FDP and inhibited by ATP and certain amino acids such as alanine and phenylalanine (15).In the presence of FDP or at a low pH, the enzyme exhibits Michaelis-Menten kinetics (23).At physiological concentrations of PEP, alanine, and ATP, liver pyruvate kinase is almost completely inhibited in vitro in the absence of FDP. Therefore, FDP has been regarded as the most important factor for regulation of the enzyme in vivo (24, 25). The kinetic and physicochemical properties of the human R isozyme are similar to those of the L isozyme (26). The M, type of pyruvate kinase that occurs in fetal and most adult tissues is an intermediate type with respect to its kinetic properties and is allosterically activated by FDP (27), whereas the M, type is insensitive to this compound ( 3 ) . The maximal activity of liver pyruvate kinase type L is high compared with the maximal rate of gluconeogenesis in the liver (1). This enzyme therefore has to be inhibited during gluconeogenesis in order to avoid wasteful substrate cycling between PEP and pyruvate. Many years ago it was found that glucagon enhanced the gluconeogenetic activity in the liver and that the concentration ratio of PEP to pyruvate was concomitantly increased, indicating stimulation of the formation of PEP from pyruvate (28, 29). Since the maximal rate of gluconeogenesis from dihydroxyacetone is higher than that from lactate or pyruvate ( 2 ) ,the rate-limiting step in gluconeogenesis in the latter case seems to be somewhere between pyruvate and PEP. Glucagon acts via cyclic 3',5'-AMP (CAMP), whose only known effect in mammals is to stimulate CAMP-dependent protein kinase (30).It therefore seems conceivable that one or more of the enzymes of the PEP-pyruvate cycle may be phosphorylated by this protein kinase. An indication of such phosphorylation was observed by Herman and collaborators, who found that liver pyruvate kinase was inhibited in rats after an intravenous injection of glucagon (31). This was also the case with phosphofructokinase, whereas fructose- 1,6-diphosphatase was activated. These effects of glucagon were counteracted by insulin. In order to test the hypothesis that liver pyruvate kinase is regulated by a CAMP-dependent phosphorylation, the rat and pig enzymes were highly purified and tested as substrates for CAMP-dependent protein kinase. They were found to be phosphorylated on serine residues, with concomitant inhibition of the enzyme activity, when assayed at a low PEP concentration (32, 33). Maximally 1 mol of phosphate is incorporated per subunit of the enzyme. A lower degree of maximal phosphorylation has also been reported (34). One dominating [32P]phosphopeptide is obtained from 32P-labeled enzyme (35, 36). It contains two arginine residues N-terminal to the phosphate-accepting serine residue.

51

2. PYRUVATE KINASE

The phosphorylation of one single serine residue in each subunit of the enzyme indicates that the reaction is specific. This is further supported by the fact that neither pig kidney pyruvate kinase type M, nor pig or rabbit muscle enzyme type M, are phosphorylated by CAMP-dependent protein kinase (37, 38). In this chapter the regulatory phosphorylation of pyruvate kinase type L by CAMP-dependent protein kinase is reviewed. Most of the results discussed were obtained with the mammalian liver enzyme. Work performed on the erythrocyte isozyme, as well as on the chicken liver pyruvate kinase type M,, is described. The role of phosphorylation of liver pyruvate kinase in the regulation of glycolysis and gluconeogenesis is briefly discussed. Reviews on the phosphorylation of liver pyruvate kinase have been previously published (39-41),

11.

Influence of Phosphorylation on the Kinetic Properties of liver Pyruvate Kinase

The dependence of the activities of unphosphorylated and phosphorylated rat liver pyruvate kinase on the concentration of the substrate PEP is illustrated in Fig. 1 . Under the in vitro conditions specified ( 4 2 ) , phosphorylation increases

7 I

.-

-~

5

2.0

E

I-

I //

--04

0

I

I

05

10

I

15

r-

PEP (mM) FIG. I , The dependence of unphosphorylated (Aand A) and phosphorylated (0 and 0 )rat liver pyruvate kinase activity on the PEP concentration. Open and filled symbols represent enzyme activity in the absence and presence of FDP, respectively. The concentration of FDP, when present, was 5 pM. From Ekman et ul. (42).

52

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST

P E P (mM) FIG. 2. The effect of phosphate and sulfate ions on the PEP dependence of unphosphorylated 50 mM irnidazole-HCI buffer, pH 7.5; (A),10 mM potassium pyruvate kinase from pig liver: substitution of 10 mM potassium phosphate added; (a),10 mM potassium sulfate added; (I), phosphate buffer (pH 7.5) for the standard imidazole-HCI buffer. From Ljungstrom et a/. (43).

(o),

Phosphorylation (mol phosphate lmol enzyme)

Fic. 3. The inactivation of pig liver pyruvate kinase, measured at 0.2 mM PEP, as a function of phosphate incorporation. From Ljungstrom et al. (43).

53

2. PYRUVATE KINASE

I

O'

515

6$0

I

I

1

I

6.5

7 .O

7.5

8O .

P FIG.4. The influence of pH on the activity of unphosphorylated (0 and 0 )and phosphorylated (A and A) pig liver pyruvate kinase. Open symbols represent 0.2 mM PEP, filled symbols 5 mM PEP. From Ljungstrom et a/. (43).

the apparent K , for this substrate from 0.3 to 0.8 mM. V,,, is not influenced by the phosphorylation. The ratios between the apparent K,,, values of the two enzyme forms reported by other authors for the rat, pig, and human enzyme are similar, but the absolute K,,, values vary between 0.3 and 1.6 mM PEP for unphosphorylated pyruvate kinase and between 0.8 and 2.5 mM PEP for the phospho-form of the enzyme (42-46). This can be explained by differences in the assay conditions. For instance, the buffer employed has a pronounced effect both on the apparent K, and on V,,, of the unphosphorylated pig liver enzyme (Fig. 2) (43). The concentration of ADP used by different investigators has varied from 1 to 2.5 mM. ADP does not have any effect on the apparent K,, for PEP, but the V,, obtained differs, since ADP in a concentration higher than 1 mM slightly inhibits the activity of pyruvate kinase (42-44). Potassium and magnesium ions are needed for pyruvate kinase to be active. For both ions a free concentration of above 20 mM is required for maximal enzyme activity, although higher concentrations of potassium are inhibitory at suboptimal PEP concentrations (42, 43). Thus, a direct comparison of kinetic constants from different laboratories is difficult unless the assay conditions are the same. During the phosphorylation of purified pyruvate kinase with CAMP-dependent protein kinase, the activity of the enzyme measured at a suboptimal concentration of PEP decreases roughly in parallel with the incorporation of phosphate (Fig. 3 ) (34, 43, 47, 48). The difference in the apparent K , values for PEP between the two enzyme forms is only found at pH values higher than 6.5, at which the enzyme exhibits

54

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST TABLE I1 THE INFLUENCE

OF

PHOSPHORYLATION ON S O M E KINETIC PARAMETERS OF PYRUVATE KINASETYPEL"

Substrates and effectors

Unphosphorylated pyruvate kinase

Phosphorylated pyruvate kinase

0.3mM

0.8mM

0.25 mM

0.25 mM

1.0 mM

0.5 mM

0.70 mM

0.35 mM

0.06 pM

0.13 WM

0.4 pM

1.4 p,M

3.0 pM

5.0 pM

PEP (apparent K,,,) ADP (apparent K,,,) ATP (50% inhibition at 0.5 mM PEP) L-Alanine (50% inhibition at 0.5 mM PEP) FDP (50% activation at 0.2 mM PEP) FDP (50% activation in the presence of I .5 mM ATP, 0.5 mM alanine and 0.2 mM PEP) FDP" (50% bound)

"Data are from Ref. ( 4 2 ) except for values for FDP binding "Data are from Ref. (51).

sigmoidal kinetics (Fig. 4). The difference is most pronounced around pH 8 (42, 43). The physiological importance of this finding is not known. ATP and alanine in the physiological concentration range (24) are potent inhibitors of both unphosphorylated and phospkylated liver pyruvate kinase, the phosphorylated enzyme being somewhat more sensitive to both inhibitors (Table 11) (42-45, 49). FDP activates both forms of the enzyme, to give similar although not identical hyperbolic activity curves (Figs. 1 and 5 ; also Table 11). In these experiments the apparent K,,, for PEP decreased to 0.04 mM. The concentration of FDP which gave half-maximal activation was 0.06 pA4 for the unphosphorylated form of the enzyme and 0.13 I.1.M for the phosphorylated form ( 4 2 ) .It has been reported that 4 mol of FDP binds to the tetrameric structure of both these enzyme forms (34, 50). The concentration of FDP needed for half-maximal binding to phosphorylated pyruvate kinase is twice that for the unphosphorylated form (Table 11) (51). In the presence of substrates, ATP and alanine, as well as of phosphate, magnesium, and potassium ions, at concentrations in the physiological range and at pH 7.4 (24), both the unphosphorylated and the phosphorylated forms of the enzyme are totally inhibited in the absence of FDP (Fig. 5). This inhibition is

55

2. PYRUVATE KINASE n v-

I

.-C

E

I

FDP( y M I FIG. 5. The activity of unphosphorylated (A and A) and phosphorylated (0and 0 ) rat liver pyruvate kinase, measured at 0.2 mM PEP, as a function of FDP concentration in the presence (filled symbols) and absence (open symbols) of ATP and alanine. The concentrations of ATP and alanine, when added, were 1.5 mM and 0.5 mM, respectively. From Ekman el al. ( 4 2 ) .

counteracted by FDP. Again, this activator has a more pronounced effect on unphosphorylated than on phosphorylated pyruvate kinase. The FDP concentration range used in the experiments of Fig. 5 could very well be in the physiological range of free FDP, as it has been demonstrated that FDP binds with high affinity to certain proteins in the cytosol (25, 52) and that its concentration decreases upon glucagon treatment of hepatocytes (53).The amount of FDP is also prone to decrease in the liver during fasting, when the rate of glycolysis approaches zero. The activator would therefore have little influence, if any, on the activity of pyruvate kinase during gluconeogenesis, which would satisfy the need for low pyruvate kinase activity under these circumstances.

111.

Influence of Phosphorylation on the Sensitivity of liver Pyruvate Kinase to Proteolytic Enzymes

The phosphorylated or phosphorylatable site of pyruvate kinase type L can be easily removed by proteolytic enzymes in vitro without any change in V,,, (5457). With bound phosphate, pyruvate kinase becomes more sensitive to proteolytic attack. For instance, there is a need for a ten times higher concentration

56

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST

of subtilisin to remove the phosphorylatable site of unphosphorylated pyruvate kinase than is necessary to split off the phosphorylated site of this enzyme (Fig. 6). This modification of the enzyme was found to give it an even higher apparent K , for the substrate PEP than phosphorylation of the enzyme. This value increased from 0.8 to 1.8 mM PEP, while V,,, remained unchanged (Fig. 7). In the presence of FDP the difference was abolished, the sigmoidality disappeared and the apparent K , for PEP decreased to about 0.05 mM (Fig. 7 ) for both the phosphorylated and the proteolytically modified form of the enzyme (55, 56). A Ca2 -activated protease from rat liver and erythrocytes also removes the phosphorylated site of pyruvate kinase at a concentration where no proteolytic activity is seen with unphosphorylated pyruvate kinase as substrate. The modified enzyme has a similar kinetic behavior to that of the subtilisin-treated enzyme (Fig. 7) (57). After removal of the phosphorylated site by the mild proteolytic modification, the molecular weight of the subunit of pyruvate kinase was not significantly reduced, as judged from polyacrylamide gel electrophoresis in sodium dodecyl sulfate under reducing conditions. This result means that peptide(s) removed amount to less than about M, 2000 (54, 56), and it also implies that the phosphorylated site is located in one end of the subunit polypeptide chain. Simon et al. obtained evidence that this site i s located in the C-terminal part of the chain (58). However, recent sequence data (59, 60) show homologies between the N+

t 30

15 TIME

L5

60

Iminl

FIG. 6. The time course of release, by subtilisin, of 32P-labeled phosphopeptides from phosphorylated pyruvate kinase (EP) and phosphate-accepting sites from unphosphorylated pyruvate kinase (E). (a),E without subtilisin; (O), EP without subtilisin; (A), E with 0.20 pglml of subtilisin; (W), E with 2.0 pg/ml of subtilisin; (A),EP with 0.20 pg/ml of subtilisin. From Bergstrom et al. (55).

2 . PYRUVATE KINASE

57

1

2 PEP

5

3

(mM)

FIG.7. The activity of phosphorylated and proteolytically modified rat liver pyruvate kinase as a no FDP present; (A), 20 p M FDP present. Open symbols, function of PEP concentration. (O), phosphorylated pyruvate kinase; filled symbols, phosphorylated pyruvate kinase treated with subtilisin; crossed symbols, phosphorylated pyruvate kinase treated with Ca2 -activated protease. Data from Bergstrom et al. (55) and Ekman and Eriksson (57). +

terminal region of M,-type pyruvate kinase and the C-terminal part (residues 2033 of the sequence shown in Table 111) of a phosphopeptide obtained after cleavage of the phosphorylated liver pyruvate kinase by cyanogen bromide (61). This rather seems to be compatible with the location of the phosphorylated site in the N-terminal region (59, 60). Several reports describing purified pyruvate kinase that incorporates less than 4 mol phosphate per mol tetrameric enzyme have appeared in the literature. Whether these pyruvate kinase batches contain enzyme that has been partially degraded by proteolysis during purification, or enzyme that has been processed in the cell for degradation or for alteration of its catalytic function is not clear (22, 34, 62-66). Slightly modified forms of pyruvate kinase cannot easily be detected through a change in their molecular weights or by an assay with saturating PEP concentrations. Nor do polyclonal, monospecific antibodies to pyruvate kinase type L seem to discriminate between native and modified enzyme. Methods employing such antibodies therefore do not allow the purification of the intact enzyme alone if modified forms are present (67). However, various forms of the liver enzyme can be separated, for example by chromatofocusing (66). Thus, proteolytically modified pyruvate kinase has been shown to exist in vivo, since even rapid purification of pyruvate kinase in the

58

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND O.ZETTERQVIST TABLE 111 AMINOACIDSEQUENCE OF PHOSPHORYLATED PEPTIDESISOLATED FROM LIVERPYRUVATE KINASEPHOSPHORYLATED B Y CAMPDEPENDENTPROTEIN KINASE Amino acid sequence" Leu-Arg-Arg-Ala-~-Leu

Source

Ref.

Pig

(35)

Pig

(61)

Rat

(36)

Rat

(61)

5

I

Glu-Gly-Pro-Ala-GI y-Tyr15

10

-Leu- Arg-Arg-Ala-Ser(P)-Leu-Ala-Gln-Leu-Thr2s

20

-Gln-Glu-Leu-Gly-Thr- Ala-Phe-Phe-Gln-Arg-Gln30

-Gln-Leu-Pro-(Ala, Ala, Homoserine)

Asx-Thr-Lys-Gly-Pro-Glx-Ile-Glx-Thr-Gly-Val-Leu-Arg-Arg-Ala-Ser-VaI-Ala-Glx-Leu I

5

Glu-Gly-Pro-Ala-Gly-Tyr15

I0

-Leu-Arg-Arg-Ala-Ser(P)-Val- Ala-Gln-Leu-Thr20

2.5

-Gln-Clu-Leu-Gly-Thr- Ala-Phe-Phe-Gln-Gln-Gln-

-Gln-(Leu, Pro, Ala, Ala, Homoserine)

OUnderlined residues indicate phosphorylated amino acids

presence of inhibitors of proteolytic enzymes gives rise to a pyruvate kinase fraction with kinetic and other properties that are almost identical to the form that has been proteolytically modified in vitro (66). In control experiments no [32P]phosphopeptides were released from added 32P-labeled pyruvate kinase during purification of the different enzyme forms. The modified enzyme must therefore exist in the intact liver before homogenization. The modified form amounts to about 15% of the pyruvate kinase in livers from fasted rats and to about 10 and 5% of that in normally fed animals and animals fed on a high-carbohydrate diet, respectively (Fig. 8) (66). In vitro, phosphorylated pyruvate kinase is more easily modified than the unphosphorylated enzyme. Somewhat unexpectedly, however, no correlation was found between the amounts of the phospho-form and the modified form in these experiments. The relative amount of phosphorylated pyruvate kinase was about equal in the livers from the three dietary groups, that is, between 50 and 60% (66). However, stress has a great influence on the metabolic state of the liver (68) and may have increased the phosphorylation of the enzyme when the animals were killed. Thus, the relative amount of phosphorylated pyruvate kinase in vivo in these experiments was somewhat uncertain and therefore no firm conclusion can be drawn regarding the role of phosphorylation in the modification of the pyruvate kinase in vivo.

59

2. PYRUVATE KINASE

u Elution volume

FIG.8. Chromatofocusing of partially purified L-type pyruvate kinase from rat liver. (O),enzyme activity at 5 mM PEP: (A), pH. The peak at pH 5.3 corresponds to a form similar to a pyruvate kinase proteolytically modified in vitro, the peak at pH 5.2 to unphosphorylated enzyme, and the peak at pH 5.0 to phosphorylated pyruvate kinase. (A) Starved rat, 45 units; (B)normally fed rats, 110 units; (C) fructose-fed rats, 300 units. Twenty milligrams of protein corresponding to about 3 g of liver were applied to 6 ml Polybuffer Exchanger columns. From Nilsson Ekdahl and Ekman (66).

IV. The Reaction of Cyclic AMP-Dependent Protein Kinase with liver Pyruvate Kinase as Substrate

Phosphorylation of liver pyruvate kinase alters the kinetic behavior of the enzyme, including the influence of allosteric effectors on the enzyme, as described in Section 11. This is interpreted as a change of the enzyme’s conformation. Conversely, the induction of an “inhibited” conformation of the unphosphorylated enzyme by negative effectors facilitates the phosphorylation, as demonstrated in several experiments. At a low pH, at which the enzyme displays Michaelis-Menten kinetics, the rate of phosphorylation is low, and an increase in pH results in an increased rate of phosphorylation (48, 69-71). The positive allosteric effector, FDP, has only minor influence on the rate of phosphorylation at lower pH values, but at higher values it inhibits the phosphorylation. Negative effectors, such as alanine and phenylalanine, have the reverse effect (Table IV). In experiments on pig-liver enzyme it was found that the influence of allosteric effectors on the rate of

60

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST TABLE IV INFLUENCE OF

PH ON THE EFFECTSOF FDP A N D ALANINE ON PHOSPHORYLATION OF PYRUVATE KINASF?‘

THE RATE

OF

Rate of phosphorylation (pmolimin) PH

Control

6

3.99 5.09 10.12

I 8

+

FDP

3.94 (- 1.3) 3.74 (-26.5) 6.62 (-34.6)

+ Alanine 5.41 (+35.6) 6.34 (+24.6) 10.21 ( + 0.9)

“Dephosphorylated pyruvate kinase (30 pg) was incubated at 30°C with 0.3 mM [y-”P]ATP, 5 mM MgC12, and 50 mM Tris-HCI at the specified pH and where indicated with 100 pA4 FDP or with I mM alanine and the reaction started by the addition of CAMP-dependent protein kinase. The percentage change from control values is given in parentheses. From El-Maghrabi et a / . (48).

phosphorylation was not due to effects on the protein kinase, since the rate of representing phosphorylation of the heptapeptide Leu-Arg-Arg-Ala-Ser-Val-Ala, the phosphorylatable site of rat liver pyruvate kinase, was not changed (69). The precise actions of allosteric effectors are dependent, however, on the degree of phosphorylation of the pyruvate kinase. When precautions were taken to dephosphorylate the enzyme by incubation with CAMP-dependent protein kinase and high concentrations of MgADP, alanine only slightly stimulated the rephosphorylation at pH 7.4, while the substrate PEP and, particularly, the positive allosteric effector FDP, decreased the rate of phosphorylation significantly (Fig. 9) (48). Although by itself alanine had only a minor effect under these conditions, it was able to relieve the inhibition caused by PEP or FDP (48). From these experiments the general concept emerged that effectors that increase the activity of liver pyruvate kinase prevent its CAMP-dependent phosphorylation, and vice versa. What structural properties make liver pyruvate kinase a substrate of CAMPdependent protein kinase? Initially it was thought that the common feature of the various substrates was some property of the three-dimensional structure (38, 72). However, in one type of substrate, histones, the site of phosphorylation appeared to be located in a more flexible part of the molecule (73). Attempts to reveal whether this part has an ordered structure did not provide any evidence of specific secondarv structure. This raised the possibility that the structural requirements of CAMP-dependent protein kinase for phosphorylation of liver pyruvate kinase are fulfilled by a small part of the polypeptide chain. Support for this hypothesis was obtained by Humble et al. who showed that both alkali-inactivated liver pyruvate kinase and a cyanogen bromide fragment of the enzyme,

61

2. PYRUVATE KINASE

I 0

w

k

a

D:

0

n

D: 0 U

f a ro N

-0 E a

‘Ll~M

0

5

10

FDP

20

30

MINUTES

FIG.9. The effects of FDP, PEP, and alanine on the rate of phosphorylation of pyruvate kinase. Dephosphorylated pyruvate kinase (30 pg) was incubated with 0 . 3 mM [y-”P]ATP, 5 mM MgCI2, with 1 mM PEP and 50 mM Tris-HCI, pH 7.4, at 30°C with no additions (e),with I ph4 FDP (O), (A),or with 1 mM alanine (A) and the reaction was started by the addition of cyclic AMP-dependent protein kinase. From El-Maghrabi er ul. (48).

were more rapidly phosphorylated than the native enzyme (37). Thus, a small part of the pyruvate kinase polypeptide chain seemed to fulfill the structural requirements for phosphorylation. The sequence of the phosphorylated site of liver pyruvate kinase has been determined both for the pig (35, 61) and the rat (36, 61) enzyme (Table 111). Two sequences of the phosphorylated site of rat liver pyruvate kinase, differing in a region not essential for phosphorylation, have been found (Table 111). The reason for this difference is not clear, but can hardly be explained by error in determination, since both sequences were compatible with the corresponding amino acid analyses (36, 61). Whether the difference is due, for instance, to subunit heterogeneity with respect to the phosphorylatable site is not known at present. The amino acid sequence data shown in Table 111 became the basis for extensive investigations to elucidate the structural requirements of CAMP-dependent protein kinase. By the use of synthetic peptides of various lengths, representing the phosphorylated site and variations of this sequence, the shortest sequence that could be phosphorylated at a significant rate was found to be Arg-Arg-Ala-SerVal (74). In addition, it was shown that both arginine residues were essential for

62

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST

a significant rate of phosphorylation. Similar results were reported by Kemp et al. (75). The implications of these data for the general understanding of the action of CAMP-dependent protein kinase are discussed elsewhere in this series. One fact that strongly supports the idea that the structural determinants for phosphorylation of liver pyruvate kinase largely reside in the primary structure of the phosphorylatable site, is that the apparent K , are of the same order for the peptides and the native enzyme. Thus, for the peptide Leu-Arg-Arg-Ala-SerVal-Ala the K , value is of the order of 0.01 mM (74), as compared to 0.02 mM for the native enzyme (71). As described in Section 111, mild proteolytic treatment removes the phosphorylated site of pyruvate kinase more readily than it removes the corresponding, unphosphorylated site. This suggests that the role of phosphorylation is to attenuate the interaction of a part of this site with the remainder of the enzyme, in order to elicit the change in kinetic properties. This part may also be responsible for mediating the cooperativity of FDP binding to the pyruvate kinase (50).

V. Dephosphorylation of liver Pyruvate Kinase with Phosphoprotein Phosphatases

Regulation of liver pyruvate kinase by means of reversible phosphorylation requires the presence of protein phosphatase activity in the same cell compartment. Evidence for such phosphatase activity was first obtained by the demonstration in v i m that phosphorylated liver pyruvate kinase was dephosphorylated by a partially purified histone phosphatase of rat liver cell sap (76). The dephosphorylation of liver pyruvate kinase is not, however, a prominent property of all protein phosphatases in the cell sap. In the extensive investigation of protein phosphatases by Ingebritsen et al. (77-79, 82, 83), most of the phosphatase activity involved in glycogen metabolism, glycolysis and gluconeogenesis, fatty acid synthesis, cholesterol synthesis, and protein synthesis was found to be accounted for by four types of enzymes, termed protein phosphatases-1, -2A, -2B, and -2C. The dephosphorylation of pyruvate kinase is mainly accounted for by protein phosphatase-2A, provided the liver extracts are highly diluted, (80-82). This type of phosphatase can be further resolved into three enzymes, namely 2A,, 2A, and 2A,, with the apparent M, values of 210,000, 210,000, and 150,000, respectively (78). As judged from their elution on DEAE-cellulose and their molecular weights, the two latter enzymes are probably identical to the two protein phosphatases of rat liver cell sap that were previously shown to be active on liver pyruvate kinase (40, 79, 84). In concentrated extracts of rat liver, protein phosphatase-2C, a highly Mg2 dependent enzyme, showed an activity towards pyruvate kinase that was equal to +

2. PYRUVATE KINASE

63

that of phosphatase-2A in the same extracts (82). It was concluded that phosphatase-2C was identical to the pyruvate kinase phosphatase studied by Jett et al. (8.5). The relative importance of phosphatases-2A and -2C in the dephosphorylation of liver pyruvate kinase in vivo is thus an open question at present. However, the identification of a particular phosphatase active on pyruvate kinase in vivo may be of value in the continued attempts to elucidate the mechanism of action of insulin. Not only the phosphorylation, but probably also the dephosphorylation is influenced by the conformation of the pyruvate kinase, at least when the enzyme is studied in Sephadex G-25-filtrated, high-speed supernatant of isolated hepatocytes (86).The dephosphorylation, measured as the activation of pyruvate kinase and dependent on divalent cations, such as Mg2+ and Mn2+, was inhibited in the presence of the substrate PEP (0.5 mM) or of the positive effector FDP (0.05 mM). The effects of these compounds were antagonized by 1-10 mM alanine (86). It is noteworthy that the effectors apparently have the same effect on the rates of phosphorylation and dephosphorylation (cf Section 1V). It may appear paradoxical that effectors aimed at activation of the pyruvate kinase counteract the activation to be achieved by dephosphorylation. This is compatible with the view, however, that the activators induce a conformation of pyruvate kinase that makes the phosphorylatable or phosphorylated serine residue less accessible to either type of converting enzyme (86). If the rate of dephosphorylation is influenced by the availability of the phosphorylated site, conformational changes of the pyruvate kinase would influence the effect of any protein phosphatase that is active on the enzyme. This would explain why the dephosphorylation of pyruvate kinase was inhibited by 1 mM PEP, in the presence of 2.5 mM MgCI,, even when a low-molecular-weight protein phosphatase of rat liver was used (87). However, experiments by Pelech et al. (80), where the dephosphorylation was measured as the release of 32Pi, showed that the rate of dephosphorylation of pyruvate kinase was not significantly changed by the presence of FDP or alanine. The inhibition of dephosphorylation obtained by high concentrations of PEP was seen also with other phosphorylated enzymes. These authors therefore concluded that no unequivocal evidence for a role of substrates or allosteric effectors in regulating the dephosphorylation of the glycolytic or gluconeogenic enzymes has so far been obtained (80). Low-molecular-weight protein phosphatase can be prepared by procedures that include an ethanol precipitation step, as introduced by Brandt et al. (88).By such a procedure, a catalytic subunit with an M,of -35,000 can be obtained both from protein phosphatase-1 and from the various phosphatases-2A (83). The preparation used in the studies of the dephosphorylation of liver pyruvate kinase (87)has been considered to be a mixture of catalytic subunits from phosphatase- 1 and phosphatase-2A (77, 79). However, chromatography of this phosphatase on

64

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST TABLE V DEPHOSPHORYLATION Ol- PHOSPHOPEPTIDES REPRES~NTINC THE PHOSPHORYLATEU SITEOF RAT LIVERPYRUVATE KINASEB Y A M, 32,000 PROTtlN PHOSPHATASE OF RAT LIVER" Substrate

K,,, (d) Relative V,,,,,

Leu-Arg-Arg-Ala-Ser( P)-Val-Ala-Gln-Leu Leu-Arg-Arg-Ala-Ser( P)-Val-Ala

0.06 0.50 0.37 0.08 b 0.03 0.03

Arg-Ala-Ser(P)-Val- Ala

Ala-Ser( P)-Val- Ala Ser(P)-Val-Ala Phosphoprotamine Phosphopyruvate kinase

4.2 1.1 1.o 0.7 b 1 .o I .o

~~

"Reactions were run in duplicate, and the mean rates were used to calculate the slopes and intercepts of Lineweaver-Burk diagrams by the method of least squares. Data from Titanji et (I/. ( 9 / ) . bThe rate of dephosphorylation of this compound at 20 pM was negligible, that is, less than 2% of the rate of dephosphorylation at an equimolar concentration of ["PI phosphoprotamine.

histone-agarose gave an apparently homogeneous preparation that was not detectably inhibited by phosphatase inhibitor- 1 or inhibitor-2 (89). This would indicate that the enzyme was derived mainly from phosphatase-2A, since the inhibitors are active only on phosphatase-1 (83). One important aspect of the dephosphorylation of pyruvate kinase is that of the structural requirements of the phosphatase. For phosphorylation of the enzyme by CAMP-dependent protein kinase, the structural requirements reside mainly in the amino acid sequence of the phosphorylatable site (see Section IV). It is therefore of interest to investigate whether a similar principle exists with respect to the dephosphorylation. Thus, phosphopeptides representing various lengths of the phosphorylated site of rat liver pyruvate kinase were assayed with a M , 32,000-protein phosphatase of rat liver, and in most cases were found to be dephosphorylated (90, 91). Contrary to the case of CAMP-dependent protein kinase, the M , 32,000-phosphatase does not seem to require basic residues just N-terminal to the phosphorylated serine. When the phosphopeptide contained two amino acids on the C-terminal side of the phosphoserine, the K,,, value was in fact lower when the basic residues were removed (Table V). The minimum sequence required for a significant rate of dephosphorylation was thus AlaSer(P)-Val-Ala. However, extension of the peptide in the C-terminal direction, without removal of the basic residues, makes it an even better substrate, with kinetic parameters similar to those of native, phosphorylated pyruvate kinase (Table V). Whereas this may indicate that essential structural determinants for

2. PYRUVATE KINASE

65

the dephosphorylation of pyruvate kinase reside in the primary structure of the phosphorylated site, comparison of the amino acid sequences on the C-terminal side of the phosphorylated serine of a number of phosphoproteins that are substrates, for example of phosphatase-2A, has not revealed any common structural features (77). The role of extension of the phosphopeptides in the C-terminal direction may therefore be essentially a matter of masking a free carboxyl group near the arginine residues. In conclusion, it appears that provided the phosphorylated site of liver pyruvate kinase is accessible to the protein phosphatase, it can be rapidly dephosphorylated. Restriction to dephosphorylation may therefore rather depend on factors that regulate the conformation of pyruvate kinase and/or protein phosphatase.

VI. Acute Hormonal Regulation of liver Pyruvate Kinase in Vivo and in Intact Cells

A prerequisite for attribution of a physiological role to the phosphorylation of liver pymvate kinase detected in vitro should be that the enzyme is phosphorylated in intact cells. Taunton et al. (31) demonstrated in 1972 that the activity of pyruvate kinase was decreased in extracts of livers from rats injected with glucagon and was increased after insulin injection. In their assay system a high concentration of PEP was used, but this must apparently have still been unsaturating, since they observed an inhibition of the activity of the pyruvate kinase. Several groups have obtained a similar effect on the enzyme activity, but only with suboptimal concentrations of this substrate (34, 44, 45, 49, 92-96). The changes in activity that occur upon hormone treatment or by varying the diet have been observed in different systems such as whole animals (70, 93), perfused livers (44), and hepatocytes (45, 49, 92). When livers are perfused or hepatocytes incubated for about 30 min before the hormonal treatment, the pyruvate kinase becomes activated and is no longer influenced by insulin unless the liver or cells are first treated with glucagon (44, 49). The apparent K,,, for PEP, as measured in extracts of control and insulin-treated hepatocytes, is similar to those of the purified, unphosphorylated pyruvate kinase studied in vitro. The apparent K,,, value for glucagon-treated systems has been found to be increased and similar to that of the phosphorylated pyruvate kinase studied in vitro (see Section 11) (44, 45, 49, 92, 93). Extracts of control and insulin-treated cells react like purified unphosphorylated pyruvate kinase with respect to inhibition by ATP and alanine, and extracts of cells treated with glucagon contain pyruvate kinase that reacts like phosphorylated pyruvate kinase (44, 45, 49, 92).

66

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0,ZETTERQVIST

These results imply that the influence of glucagon on pyruvate kinase of hepatocytes is due to glucagon-induced phosphorylation of the enzyme. Evidence of this was first obtained by Ljungstrom and Ekman (94)in experiments on rat liver slices. The inactivation of pyruvate kinase measured at unsaturating concentrations of PEP was accompanied by an increase in 32P-labelingof pyruvate kinase isolated from liver slices by an immunosorbent, after incubation with 32Piand glucagon (Fig. 10). It was also demonstrated by phosphopeptide mapping that the [32P]pho~phopeptide~ obtained from the phosphorylated pyruvate kinase of the slices were identical to those from the enzyme phosphorylated in v i m . This indicates that CAMP-dependent protein kinase is also responsible for the glucagon-induced phosphorylation in vivo. Similar evidence has been obtained in other laboratories when hepatocytes or perfused rat livers have been used (34, 95, 96). As seen in Fig. 10, the incorporation of phosphate had reached a maximal value when the cAMP concentration was still rising. This is consistent with the demonstration that cAMP exerts its effects at concentrations far below the maximal (97, 98). The fact that pyruvate kinase purified from rat liver contains various amounts of covalently bound phosphate, also indicates that phosphorylation of pyruvate kinase is involved in the regulation of the enzyme in vivo (34, 48, 66). It has been shown that not only glucagon but also epinephrine, norepinephrine, and phenylephrine lower the activity of pyruvate kinase (49, 99101). The time-dependent inactivation by epinephrine is concomitant with an increase in the phosphate content of the enzyme (99, ZOO). Digestion of this I

I

2:

2

0

y

,

0

2

1

5 Incubation time

I

l o

10

Iminl

Fig. 10. The effect of 10-7 M glucagon on pyruvate kinase phosphorylation and the cAMP concentration in rat liver slices. Samples were analyzed for phosphorylation of pyruvate kinase, the ratio of enzyme activity at 0.5 to that at 5.0 mM PEP, and the cAMP concentration. From Ljungstrom and Ekman ( 9 4 ) .

67

2. PYRUVATE KINASE

0.6 >,

c ..-c>

0.4

c

0

.-c0 m

a

0.2

t

t

insulin

glucagon

10

20 Time (rnin)

Fig. 1I . The effect of incubation of hepatocytes with glucagon and insulin on the activity ratio of pyruvate kinase ( ~ 0 .mM 5 p~p/vs.o PEP): (O), control; (O), 2 X 10- 12Mglucagon added at 5 min; (A), 2 X 10-l2M glucagon added at 5 min and 2 X 10- l o M insulin at 15 min. The amounts of phosphate incorporated at 25 min were 0.40, 0.71, and 0.45 mol/mol subunit in the control, the glucagon-treated and the glucagon-plus-insulin-treatedsamples, respectively (P. Ekman and L. Engstrom, unpublished data).

pyruvate kinase by trypsin yields the same phosphopeptide pattern as tryptic digestion of pyruvate kinase from glucagon-treated hepatocytes (99). The epinephrine-induced phosphorylation seems to be mediated by both a- and P-receptors, although to a varying degree, depending on the species used and the age of the animal (100-105). The P-receptor-mediated effects are apparently exerted via CAMP, while those mediated by a-receptors seem to be CAMP-independent. Other hormones that influence the activity and phosphorylation of pyruvate kinase are vasopressin and angiotensin I1 (104, 106, 107).The molecular basis of the action of these hormones is not quite understood, but some observations have indicated the involvement of calcium ions ( 3 ) .It has been claimed that glucocorticoids have a permissive effect on the phosphorylation of pyruvate kinase by glucagon, since the phosphorylation and inhibition of the enzyme activity are impaired considerably in hepatocytes from adrenalectomized rats (108). Nor does glucagon injected in vivo into such rats inhibit the pyruvate kinase activity assayed at a low PEP concentration. The mechanism of this permissive effect is not known. It may be hypothesized that the reactivation of pyruvate kinase that is observed when hepatocytes are incubated with insulin after pretreatment with glucagon, is

68

L. ENGSTROM, P. EKMAN, E. HUMBLE, A N D 0.ZETTERQVIST

paralleled by a decrease in the phosphorylation of the enzyme. We found that the incorporation of 32P per mol of subunit decreased significantly upon insulin treatment (Fig. 11). This indicates that the acute activation of the pyruvate kinase by insulin is due to the dephosphorylation of the enzyme. The mechanism of this action of insulin is, however, unknown.

VII. Phosphorylation of Other Pyruvate Kinases A. KIDNEYENZYME,TYPEL Renal cortex is a tissue with a gluconeogenetic capacity, and rat kidney is reported to contain, as a minor component of its pyruvate kinase activity, the Ltype isozyme (11,12). Like its counterpart in the liver, kidney pyruvate kinase, type L, is phosphorylated on serine residues upon incubation with ATP and CAMP-dependent protein kinase. The phosphorylation is accompanied by an increase in the apparent K, for PEP. This effect on the kinetic properties of pyruvate kinase is reversed by the action of phosphoprotein phosphatase (12). Thus, the results of in v i m experiments indicate that in the kidney, also, pyruvate kinase activity might be regulated via cAMP during gluconeogenesis. Furthermore, it has been demonstrated that pyruvate kinase activity in the rat renal cortex decreases after injection of glucagon or cAMP into the portal vein (109).

B. INTESTINAL PYRUVATE KINASE Pyruvate kinase present in the rat small intestine can be separated by electrophoresis into five forms (I], 13, ]lo), representing type L, type M, and hybrids (13, 110). It seems reasonable to assume that also the intestinal L isozyme and, possibly, hybrids containing the L subunit can be phosphorylated by CAMP-dependent protein kinase. In support of this idea, it has been found that the concentration of cAMP and the activity of fructose- 1,6-bisphosphatase increase, whereas the activity of pyruvate kinase decreases, when rabbit jejunal mucosa, maintained in organ culture, is exposed to cholera toxin (111).

C.

ERYTHROCYTE ENZYME

Human erythrocyte pyruvate kinase seems to be a homotetramer which coexists with proteolytically modified enzyme (4,112, 113). During aging of the red cell, the proportion of enzyme molecules with somewhat reduced molecular weights increases (112). Both the parent molecule and slightly degraded forms incorporate phosphate when partially purified preparations containing endoge-

2. PYRUVATE KINASE

69

nous protein kinase are incubated with ATP and cAMP (4, 114). A maximal incorporation of 1 mol phosphate per mol subunit has been reported. The specific inhibitor of CAMP-dependent protein kinase is capable of inhibiting all pyruvate kinase phosphorylation in these preparations (4). Evidence of in vivo phosphorylation of erythrocyte pyruvate kinase has been obtained in experiments on rats, from which radioactively labeled pyruvate kinase was isolated from red cells following intravenous injection of 32Pi(115). It has also been demonstrated that 32P-labeledpyruvate kinase can be isolated from human and rat erythrocytes after incubation of the cells with 32Piand cAMP (116-118). Omission of the cyclic nucleotide results in a severalfold lower degree of phosphorylation (I16, I 17). Phosphorylation of purified pyruvate kinase from human erythrocytes has been reported to result in the same kinetic modifications as for the liver enzyme [i.e., an increased K , value for PEP in the absence of saturating concentrations of FDP (4, 119), and increased inhibition by ATP and alanine ( 4 ) ] .The enzyme can be dephosphorylated and reactivated by incubation with phosphoprotein phosphatase (115, 119, 120). The physiological significance of CAMP-dependent phosphorylation of pyruvate kinase in mature erythrocytes is not clear, since gluconeogenesis does not occur in these cells. However, the pyruvate kinase reaction is reported to be important in establishing the steady-state level of 2,3-diphosphoglycerate. This level was found to be inversely related to the amount of pyruvate kinase (121). An increase in the level of 2,3-diphosphoglycerate has been observed in human red cells incubated with catecholamines, prostaglandin E,, or cAMP together with phosphodiesterase inhibitors (122). Westhead et al. reported that the concentration of 2,3-diphosphoglycerate increases rapidly in erythrocytes stimulated with cAMP (120). Inactivation of pyruvate kinase by phosphorylation would thus serve to promote the release of oxygen to the tissues, as discussed in more detail by Westhead and co-workers (120). Marie et al., however, found that the concentration of 2,3-diphosphoglycerate did not change in human erythrocytes during incubation with cAMP (4). It has been suggested that 2,3-diphosphoglycerate might be involved in the regulation of red cell pyruvate kinase since it inhibits phosphorylation of the enzyme in vitro (I1 7 , 119) and since pyruvate kinase is more heavily phosphorylated in red cells depleted of both ATP and 2,3-diphosphoglycerate than in those depleted of ATP alone (117). Another matter of discussion is what physiological events might lead to activation of the protein kinase in the erythrocytes. Catecholamine-stimulated protein kinase activity has been demonstrated in human erythrocyte membranes (12.9, although several investigators have reported that in mature human red cells adenylate cyclase is only slightly activated by catecholamines [see Refs. (124,

70

L. ENGSTROM, P. EKMAN, E. HUMBLE, AND 0.ZETTERQVIST

125), and references therein]. However, cAMP from the outside penetrates the erythrocyte membrane (117, 126, 127). The possibility that cAMP released from epinephrine-stimulated endothelial cells might activate erythrocyte protein kinase is discussed by Westhead et al. (120). Red cell pyruvate kinase has also been reported to be phosphorylated by a calcium- and calmodulin-dependent mechanism in a cell-free system and phosphorylated pyruvate kinase has been isolated from erythrocytes incubated with 32Pi and calcium (128). The effect of this phosphorylation on pyruvate kinase activity and the possible physiological significance of the reaction remain to be elucidated.

D.

LIVER PYRUVATE KINASEIN CHICKEN

Chicken hepatocytes respond to glucagon and dibutyryl-CAMP by a decrease in glucose utilization and an increase in gluconeogenesis from lactate and dihydroxyacetone, although the pyruvate kinase activity appears to be unchanged (129).

Chicken liver contains predominantly the M, isozyme of pyruvate kinase, but also a certain amount of the L type (130, 131). The amino acid composition of chicken liver pyruvate kinase of type M, differs from that of M, in other species and from that of M, in the chicken (132). Chicken liver pyruvate kinase of types M, and L is phosphorylated and inactivated in vitro by a CAMP-independent protein kinase purified from the same tissue (132, 133). The protein kinase can utilize both ATP and GTP and in addition to chicken liver pyruvate kinase, it also phosphorylates phosvitin and casein, but not histones or phosphorylase b (133). When phosphorylated to 1 mol phosphate per mol subunit, M, pyruvate kinase is completely inactivated, even when tested at a high concentration of PEP in the absence or presence of FDP (133). The phosphate-accepting amino acid is a serine residue in an acid environment (132). Chicken liver also contains a pyruvate kinase-reactivating phosphoprotein phosphatase. Based upon the effects of FDP and alanine on the inactivating and reactivating processes, and upon the apparent molecular weight of phosphorylated pyruvate kinase, a model has been suggested in which the protein kinase preferably phosphorylates the less active dimeric form of pyruvate kinase, whereas the tetrameric form is a better substrate for the phosphatase (134). Pyruvate kinase of type M,, phosphorylated on serine, has been isolated from chicken hepatocytes after their incubation with 32Pi. Chicken embryo cells contain M, pyruvate kinase. Transformation of these cells by the Rous sarcoma virus leads to a lower affinity of the pyruvate kinase for PEP, high stimulation by FDP, and rapid inactivation by ATP (136). Furthermore, a protein kinase associated with the gene product of Rous sarcoma virus catalyzes the phosphorylation and inactivation of purified M, pyruvate kinase

2. PYRUVATE KINASE

71

from chicken liver. In this case tyrosine residues seem to be the acceptors of phosphate (136, 137).

VIII. Concluding Remarks From the results reported it is fairly well ascertained that the phosphorylation of mammalian liver pyruvate kinase is of the regulatory type as defined by Nimmo and Cohen (138) and by Krebs and Beavo (139).Thus, in virro the enzyme is stoichiometrically phosphorylated by CAMP-dependent protein kinase at an adequate rate with a physiologically meaningful change of the activity of the former enzyme. The phosphorylation of the pyruvate kinase and the changes of its activity are reversed by phosphoprotein phosphatase. In vivo and in isolated hepatocytes the enzyme is subject to the same functional changes in the presence of glucagon or cAMP as the purified enzyme when it is incubated with CAMP-dependent protein kinase and ATP. The same serine residue in the pyruvate kinase seems to be phosphorylated in both cases. The degree of phosphorylation appears to correlate fairly well with the changes in the kinetic properties of the enzyme in vitro, and with the hormonal response and cAMP level in intact cells. The phosphorylation of liver pyruvate kinase seems to be one important point-but not the only one-for rapid hormonal regulation of glycolysis and gluconeogenesis (2, 3 ) . Evidences indicating such a role of pyruvate kinase have been obtained by several groups when correlating the influence of hormones or cAMP on the flux through the pyruvate kinase reaction or through the whole glycolytic and gluconeogenic pathways with their effects on the pyruvate kinase. In 1976 Hers and collaborators found that inactivation of pyruvate kinase is related to stimulation of gluconeogenesis (49). They added different concentrations of glucagon to isolated hepatocytes from fed rats. The activity of pyruvate kinase decreased roughly in parallel with an increase in the rate of gluconeogenesis. When insulin was added together with glucagon, the effect of glucagon on the pyruvate kinase activity and on the rate of gluconeogenesis were counteracted to approximately the same extent. Studies on the flux through the pyruvate kinase step in isolated hepatocytes by Rognstad and Katz have shown that glucagon inhibits this flux during stimulation of gluconeogenesis in cells from fed rats but hardly at all in cells from fasted rats (140). Treatment of cells from fed rats with epinephrine had virtually no effect on the flux through the pyruvate kinase reaction. Thus, the mechanism of the stimulation of gluconeogenesis by epinephrine is different from that of glucagon. The question whether or not phosphorylation of pyruvate kinase increases the rate of degradation of the enzyme in vivo has not yet been clarified. The possibility that phosphorylation not only influences the activity of the enzyme but also effects its turnover rate needs further investigation.

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L. ENGSTROM, P. EKMAN, E. HUMBLE, AND OZETTERQVIST

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Ekman, P., Dahlqvist, U., Humble, E., and Engstrom, L. (1976). BBA 429, 374. Ljungstrom, 0.. Berglund, L., and Engstrom, L. (1976). EJB 68, 497. Blair, J. B., Cimbala, M. A., Foster, J. L., and Morgan, R. A. (1976). JBC 251, 3756. Riou, J . P., Claus, T. H., and Pilkis, S. J. (1976). BBRC 73, 591. van den Berg, G.B., van Berkel, T. J. C., and Koster, J. F. (1978). BBRC 82, 859. Titanji, V. P. K., Zetteqvist, O., and Engstrom, L. (1976). BBA 422, 98. El-Maghrabi, M. R., Haston, W. S., Flockhart, D. A,, Claus, T. H., and Pilkis, S. J. (1980). JBC 255, 668. 49. Feliu, J. E., Hue, L., and Hers, H.-G. (1976). PNAS 73, 2762. 50. El-Maghrabi, M. R., Claus, T. H., McGrane, M. M., and Pilkis, S. J. (1982).JBC 257, 233. 51. Blair, J . B., and Walker, R. G.(1984). ABB 232, 202. 52. Sols, A., and Marco, R. (1970). Curr. Top. Cell. Regul. 2, 227. 53. Pilkis, S. J . , Riou, J. P., and Claus, T. H. (1976). JBC 251, 7841. 54. Bergstrom, G.,Ekman, P., Dahlqvist, U., Humble, E., and Engstrom, L. (1975). FEBS Lett. 56, 288. 55. Bergstrom, G.,Ekman, P., Humble, E., and Engstrom, L. (1978). BBA 532, 259. 56. Nakai, N . , Fujii, Y., Kobashi, K., and Hase, J. (1983). BBRC 110, 682. 57. Ekman, P., and Eriksson, I. (1980). Acta Chem. Scand., BM, 419. 58. Simon, M.-P., Marie, J., Bertrand, O., and Kahn, A. (1982). BBA 709, 1. 59. Hoar, C. G.,Nicoll, G. W., Schiltz, E., Schmitt, W., Bloxham, D. P., Byford, M. F., Dunbar, B., and Fothergill, L. A. (1984). FEBS Lert. 171, 293. 60. Lonberg, N., and Gilbert, W. (1983). PNAS 80, 3661. 61. Humble, E. (1980). BBA 626, 179. 62. Kohl, E. A., and Cottam, G.L. (1976). ABB 176, 671. 63. van Berkel, T. J. C., Kruijt, J. K., van den Berg, G.B., and Koster, J. F. (1978).EJB 92,553. 64. Hall, E. R., McCully, V., and Cottam, G.L. (1979). ABB 195, 315. 65. Poole, G.P., and Bloxham, D. P. (1982). BJ 204, 89. 66. Nilsson Ekdahl, K., and Ekman, P. (1984). J. Biochem. (Tokyo) 95, 917. 67. Dahlqvist-Edberg, U. (1978). FEBS Lert. 88, 139. 68. Faupel, R. P., Seitz, H. J., Tarnowski, W., Thiemann, V., and Weiss, C. (1972). ABB 148, 509. 69. Berglund, L., Ljungstrom. O., and Engstrom, L. (1977). JBC 252, 613. 70. Feliu, J . E., Hue, L., and Hers, H.-G. (1977). EJB 81, 609. 71. El-Maghrabi, M. R., Claus, T. H., and Pilkis, S. J. (1983). “Methods in Enzymology,” Vol. 99, p. 212. 72. Langan, T. A. (1973). Adv. Cyclic Nucleoride Res. 3, 99. 73. Lewis, P. N., and Bradbury, E. M. (1974). BBA 336, 153. 74. Zetterqvist, 0.. Ragnarsson, U., Humble, E., Berglund, L., and Engstrom, L. (1976). BBRC 70, 696. 75. Kemp, B. E., Graves, D. J., Benjamini, E., and Krebs, E. G.(1977). JBC 252, 4888. 76. Titanji, V. P. K., Zetterqvist, O., and Engstrom, L. (1976). BBA 422, 98. 77. Ingebritsen, T. S . , and Cohen, P. (1983). EJB 132, 255. 78. Ingebritsen, T. S., Foulkes, J. G.,and Cohen, P. (1983). EJB 132, 263. 79. Ingebritsen, T. S . , Blair, J., Guy, P., Witters, L., and Hardie, D. G.(1983). EJB 132, 275. 80. Pelech, S . , Cohen, P., Fischer, M. J., Pogson, C. I., El-Maghrabi, M. R., and Pilkis, S. J. (1984). EJB 145, 39. 81. Alemany, S., Tung, H. Y. L., Shenolikar, S., Pilkis, S. J., and Cohen, P. (1984). EJB 145, 5 1 . 82. Ingebritsen, T. S . , Stewart, A. A., and Cohen, P. (1983). EJB 132, 297. 83. Ingebritsen, T. S . , and Cohen, P. (1983). Science 221, 331.

42. 43. 44. 45. 46. 47. 48.

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L. ENGSTROM, P. EKMAN, E. HUMBLE, AND OZETTERQVIST

84. Titanji, V. P. K. (1978). UpsalaJ. Med. Sci. 83, 129. 85. Jett, M. F., Hue, L., and Hers, H. G. (1981). FEES Lett. 132, 183. 86. Mojena, M., and Feliu, J. E. (1983). Mol. Cell. Biochem. 51, 103. 87. Titanji, V. P. K. (1977). BBA 481, 140. 88. Brandt, H., Capulong, Z. L., and Lee, E. Y. C. (1975). JBC 250, 8038. 89. Titanji, V. P. K., Zetterqvist, O., and Engstrom, L. (1980). FEES Lett. 111, 209. 90. Titanji, V. P. K., Zetterqvist, 0.. and Ragnarsson, U. (1977). FEES Lett. 78, 86. 91. Titanji, V. P. K., Ragnarsson, U., Humble, E., and Zetterqvist, 0. (1980). JBC 255, 11339. 92. van Berkel, T. J. C., Kruijt, J. K., Koster, J. F., and Hulsmann, W. C. (1976). BBRC 72,917. 93. Kohl, E. A., and Cottam, G. L. (1977). BBA 484, 49. 94. Ljungstrom, O., and Ekman, P. (1977). BBRC 78, 1147. 95. Riou, J. P., Claus, T. H., and Pilkis, S. J. (1978). JBC 253, 656. 96. Ishibashi, H., and Cottam, G. L. (1978). JBC 253, 8767. 97. Okajima, F., and Ui, M. (1976). ABB 175, 549. 98. Park, C. R., and Exton, J. H. (1972). In “Glucagon: Molecular Physiology, Clinical and Therapeutic Implications” (P. J. Lefebre and R. H. Unger, eds.), p. 77. Pergamon, Oxford. 99. Nagano, M., Ishibashi, H., McCully, V., and Cottam, G. L. (1980). ABB 203, 271. 100. Steiner, K. E., Chan, T. M., Claus, T. H., Exton, J. H., and Pilkis, S. J. (1980). BBA 632, 366. 101. Blair, J. B., James, M. E., and Foster, J. L. (1979). JBC 254, 7585. 102. Kemp, B. E., and Clark, M. G. (1978). JBC 253, 5147. 103. Yorek, M. A., Blair, J. B., and Ray, P. D. (1982). BBA 717, 143. 104. Chan, T. M., and Exton, J. H. (1978). JBC 253, 6393. 105. Blair, J. B., James, M. E., and Foster, J. L. (1979). JBC 254, 7579. 106. Garrison, J . C., Borland, M. K., Florio, V. A., and Twible, D. A. (1979). JBC 254, 7147. 107. Garrison, J. C., and Wagner, J. D. (1982). JBC 257, 13135. 108. Postle, A. D., and Bloxham, D. P. (1982). EJB 124, 103. 109. Taunton, 0. D., Stifel, F. B., Greene, H. L., and Herman, R. H. (1974). JBC 249, 7228. 110. Saheki, S., Harada, K., Sanno, Y., and Tanaka, T. (1978). BBA 526, 116. 111. Sherr, H. P., Stifel, F. B.. and Herman, R. H. (1978). Gastroenterology 75, 71 1. 112. Kahn, A., Marie, J . , Garreau, H., and Sprengers, E. D. (1978). BBA 523, 59. 113. Sprengers, E. D., and Staal, G. E. J. (1979). BBA 570, 259. 114. Marie, J., and Kahn, A. (1980). BBRC 94, 1387. 115. Fujii, S., Nakashima, K., and Kaneko, T. (1981). Biomed. Res. 2, 316. 116. Marie, J., Tichonicky, L., Dreyfus, J.-C., and Kahn, A. (1979). BBRC 87, 862. 117. Fujii, S., Nakashima, K., and Kaneko, T. (1980). Biomed. Res. 1, 230. 118. Dahlqvist-Edberg, U . , and Ekman, P. (1981). BBA 660, 96. 119. Kiener, P. A., Massaras, C. V., and Westhead, E. W. (1979). BBRC 91, 50. 120. Westhead, E. W., Kiener, P. A,, Carroll, D., and Gikner, J. (1984). Curr. Top. Cell. Regul. 24,21. 121. Rose, I. A. (1971). Exp. Eye Res. 11, 264. 122. Badwey, J. A., and Westhead, E. W. (1978). In “The Red Cell” (G. J. Brewer, ed.), p. 299. Alan R. Liss, Inc., New York. 123. Tsukamoto, T., and Sonenberg, M. (1979). J. Clin. Invesr. 64, 534. 124. Sager, G. (1982). Biochem. Pharmacol. 31, 99. 125. Nakagawa, M., Willner, J., Cem, C., and Reydel, P. (1984). BBA 770, 122. 126. Thomas, E. L., King, L. E., Jr., and Momson, M. (1979). ABB 196, 459. 127. Tsukamoto, T., Suyama, K., Germann, P., and Sonenberg, M. (1980). Biochemistry 19,918. 128. Nakashima, K., Fujii, S., Kaku, K., and Kaneko, T. (1982). BBRC 104, 285. 129. Ochs, R. S., and Hams, R. A. (1978). ABB 1% 193.

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130. Strandholm, J . J . , Cardenas, J . M., and Dyson, R. D. (1975). Biochemistry 14, 2242. 131. Eigenbrodt, E., and Schoner, W. (1977). Hoppe-Seyler’s Z. Physiol. Chern. 358, 1033. 132. Brunn, H . , Eigenbrodt, E., and Schoner, W. (1979). Hoppe-Seyler’s 2. Physiol. Chem. 360, 1357. 133. Eigenbrodt, E., Abdel-Fattah Mostafa, M., and Schoner, W. (1977). Hoppe-Seyler’s Z. Physiol. Chem. 358, 1047. 134. Eigenbrodt, E., and Schoner, W. (1977). Hoppe-Seyler’s Z . Physiol. Chern. 358, 1057. 135. Fister, P., Eigenbrodt, E., Presek, P., Reinacher, M., and Schoner, W. (1983). EERC 115, 409. 136. Presek, P., Glossman, H., Eigenbrodt, E., Schoner, W., Riibsamen, H., Friis, R. R., and Bauer, H. (1980). Cancer Res. 40, 1733. 137. Glossman, H., Presek, P., and Eigenbrodt, E. (1981). Mol. Cell. Endocrinol. 23, 49. 138. Nimmo, H. G., and Cohen, P. (1977). Adv. Cyclic Nucleotide Res. 8, 145. 139. Krebs, E. G., and Beavo, J. A. (1979). Annu. Rev. Eiochern. 48, 923. 140. Rognstad, R., and Katz, J. (1977). JEC 252, 1831.

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Pyruvate Dehydrogenase LESTER J. REED* STEPHEN J. YEAMANT *Department of Chemistry The University of Texas at Austin Austin, Texas 78712 fDepartment of Biochemistry The University of Newcastle upon Tyne Newcastle upon Tyne, NEI 7RU United Kingdom

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Mammalian Pyruvate Dehydrogenase Complex A. Subunit Composition and Structure ..............................

B. Phosphorylation Sites . . . . . . . 111. Pyruvate Dehydrogenase Kinase . . A. Isolation and Physicochemical ......................... B. Regulatory Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Pyruvate Dehydrogenase Phosphatase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Isolation and Physicochemical Properties ......................... B. Regulatory Properties V. Regulation of Mammalian Pyruvate Dehydrogenase Complex . . . . . . . . . . . VI. Comparison of Properties of Mitochondria1 a-Ketoacid Dehydrogenase Kinases and Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . References

1.

77 79 79 81

82 82 83 84 84 84 86 92 93

Introduction

Enzyme systems that catalyze a lipoic acid-mediated oxidative decarboxylation of a-ketoacids have been isolated from microbial and eukaryotic cells as functional units with molecular weights in the millions. Three types of complex77 THE ENZYMES, Vol. XVIll Copyright Q 1987 by Academic Press. Inc. All rights of reproduction in any form reserved.

78

LESTER J . REED AND STEPHEN J . YEAMAN NAD+

OH I [RC = TPP]

0 II

[RC-SLipSH]

I CoASH

0 II RCCO,H

0

II + COASH + NAD+ -+ RC-SCOA + co, + N A D H + H+

FIG. I . Reaction sequence in pyruvate oxidation (R = CH3). The following abbreviations are used: TPP, thiamin diphosphate; Lipsz and Lip(SH)z, lipoyl moiety and its reduced form; CoASH, coenzyme A; FAD, flavin adenine dinucleotide; NAD and NADH, nicotinamide adenine dinucleotide and its reduced form; E l , pyruvate dehydrogenase; Ez, dihydrolipoamide acetyltransferase; E-,, dihydrolipoamide dehydrogenase. +

es have been obtained: one specific for pyruvate, a second specific for a-ketoglutarate, and a third specific for the branched-chain a-ketoacids (a-ketoisovaleric, a-ketoisocaproic and a-keto-p-methylvaleric acids). Each complex is composed of multiple copies of three major components: a substrate-specific dehydrogenase (E,); a dihydrolipoamide acyltransferase (E,) specific for each type of complex; and dihydrolipoamide dehydrogenase (E3), a flavoprotein that is a common component of the three types of multienzyme complexes. These three enzymes, acting in sequence, catalyze the reactions shown in Fig. 1 ( 1 , 2). El catalyzes both the decarboxylation of the a-ketoacid (reaction 1) and the subsequent reductive acylation of the lipoyl moiety (reaction 2), which is covalently bound to E,. E, catalyzes the transacylation step (reaction 3), and E, catalyzes the reoxidation of the dihydrolipoyl moiety with NAD as the ultimate electron acceptor (reactions 4 and 5 ) . The pyruvate dehydrogenase complexes from mammalian and avian tissues and Neurospora crussu and the mammalian branched-chain a-ketoacid dehydrogenase complex also contain small amounts of two specific regulatory enzymes, pyruvate dehydrogenase kinase and phosphatase and branched-chain a-ketoacid dehydrogenase kinase and phosphatase, respectively, which modulate the activity of El by phosphorylation and dephosphorylation [Refs. (3-7); also see Chapter 4, this volume]. There is no +

3. PYRUVATE DEHYDROGENASE

79

evidence that the pyruvate dehydrogenase complex or the branched-chain aketoacid dehydrogenase complex in prokaryotic cells or the a-ketoglutarate dehydrogenase complexes in eukaryotic or prokaryotic cells undergo phosphorylation and dephosphorylation. This chapter discusses some aspects of the structural organization of the mammalian pyruvate dehydrogenase complex and regulation of its activity by a phosphorylation-dephosphorylation cycle.

II. Mammalian Pyruvate Dehydrogenase Complex A.

SUBUNIT COMPOSITION AND STRUCTURE

Each of the three types of a-ketoacid dehydrogenase complexes is organized about a core, consisting of the oligomeric E,, to which multiple copies of E, and E, are bound by noncovalent bonds. Two polyhedral forms of E, have been observed in the electron microscope, the cube and the dodecahedron (Fig. 2) (2); both designs are based on cubic point group symmetry. The former design is exhibited by the E, components of the pyruvate dehydrogenase and a-ketoglutarate dehydrogenase complexes of Escherichia coli and the mammalian a-ketoglutarate dehydrogenase and branched-chain a-ketoacid dehydrogenase com-

a

FIG.2. Interpretive models of the quaternary structure of dihydrolipoamide acyltransferases. (A) Model of those acyltransferases consisting of 24 subunits arranged in groups of 3 about the 8 vertices of a cube. (B) Model of the 24-subunit acyltransferases illustrating the proposed domain and subunit structure. Each of the 24 acyltransferase subunits is represented by one sphere and its attached ellipsoid. The spheres represent the assemblage of compact domains (inner core), and the ellipsoids represent the extended lipoyl domains. (C) Model of those acyltransferases consisting of 60 subunits arranged in groups of 3 about the 20 vertices of a pentagonal dodecahedron. (D) Model of the 60subunit acyltransferases illustrating the proposed domain and subunit structure. The figure is viewed down a 2-fold axis of symmetry.

80

LESTER J . REED AND STEPHEN J . YEAMAN

plexes. These E, components consist of 24 identical subunits arranged with octahedral (432) symmetry. On the other hand, the E, components of the pyruvate dehydrogenase complexes from mammalian and avian tissues, fungi, and Bacillus stearothermophilus have the appearance of a pentagonal dodecahedron in the electron microscope and consist of 60 subunits apparently arranged with icosahedral (532) symmetry. The E, subunit of the E . coli and Azotobacter vinelandii pyruvate dehydrogenase complexes contains two and possibly three covalently bound lipoyl moieties, but all other dihydrolipoamide acyltransferase subunits examined contain only one lipoyl moiety. The lipoyl moiety is bound in amide linkage to the €-amino group of a lysine residue. It should be noted that the E, components have a rather large cavity in their structure (2). The physiological significance, if any, of this cavity has yet to be determined. Another interesting feature of the structure, revealed by limited proteolysis and electron microscopy, is that the E, subunit consists of two different domains: a compact domain and a flexible, extended domain (Fig. 2) (8-11). The compact domain contains the acyltransferase active site, and the assemblage of these domains constitutes the “inner core” of E,, conferring the cube-like or pentagonal dodecahedron-like appearance in the electron microscope. The extended domain, which is readily released from the inner core by limited proteolysis, contains the covalently bound lipoyl moiety or moieties (lipoyl domain). Proton nuclear magnetic resonance spectroscopy has provided evidence that the lipoyl domain is attached to the inner core by a highly mobile segment of the polypeptide chain (12, 13). This unique architectural feature is thought to facilitate interaction of the lipoyl moiety with successive active sites on the complex, that is, a multiple random coupling mechanism (14). The pyruvate dehydrogenase complexes isolated from bovine kidney and of about 7,000,000 and 8,500,000, respecheart have molecular weights (M,) tively. The component enzymes of the two complexes are very similar, if not identical (15). El has a M, of about 154,000 and possesses the subunit composition a,P, (Table I). The M, of the two subunits are about 41,000 and 36,000, respectively. The core enzyme (E,) has a M , of about 3,100,000 and consists of 60 apparently identical polypeptide chains of M, about 52,000. Each E, chain contains one covalently bound lipoyl moiety. The isolated E, is a homodimer of M, about 110,000 and contains two molecules of FAD. The bovine kidney pyruvate dehydrogenase complex contains about 20 E, tetramers (a2P2)and about 6 E, dimers, whereas the heart complex contains about 30 E, tetramers and 6 E, dimers. The kidney complex can bind about 10 additional E, tetramers, but neither complex can bind additional E, dimers. The dissociation constant ( K J of the E,-E, subcomplex is about 13 nM, and the Kd of the E,-E, subcomplex is about 3 nM (16).The E, tetramers appear to be located on the 30 edges, and the E, dimers in the 12 faces of the E, pentagonal dodecahedron. The kinase is tightly bound to E, and copurifies with the pyruvate dehydrogenase complex.

81

3. PYRUVATE DEHYDROGENASE

TABLE I SUBUNIT COMPOSITION OF BOVINE HEARTPYRUVATE DEHYDROCENASE COMPLEX Subunits Enzyme

Mr

Native Complex El EIa EIP E2 E3 Kinase Phosphatase

8,500,000 154,000

No.

Mr

4 2 2

41,000

3,100,000 110,000

60

- 100,000

2 1 1

150,000

I 1

36,000 52,000 55,000 48,000 45,000

Subunits per molecule of complex

60 60 60 12

97,000 50,000

The amount of endogenous kinase in the bovine kidney and heart complexes is small, about three molecules per molecule of kidney complex and less in the heart complex. The phosphatase also binds to E,, and this attachment requires the presence of Ca2+ ions (17). There appear to be about five molecules of phosphatase per molecule of complex in bovine kidney mitochondria.

B. PHOSPHORYLATION SITES Phosphorylation and concomitant inactivation of pyruvate dehydrogenase (E,) occurs on three serine residues in the a subunit (M,= 41,000) (18, 19). Tryptic digestion of 32P-labeled pyruvate dehydrogenase from bovine kidney and heart yielded three phosphopeptides, a monophosphorylated (site- 1) and a diphosphorylated (site- 1 and -2) tetradecapeptide, and a monophosphorylated nonapeptide (site-3) (Fig. 3). Although Yeaman et al. (18) concluded that the tryptic tetradecapeptide contained asparagine at residue eight, later studies indicate that this residue is aspartic acid (L. R. Stepp, T. R. Mullinax, and L. J. Reed, unpublished data). The revised sequence, containing an acid-labile Asp-Pro bond, is in agreement with the sequence found for the tryptic tetradecapeptide obtained from pig heart pyruvate dehydrogenase (19).

Tyr-His

Site I -

- Giy- H is -Ser(P)-Met

-Ser-Asp-Pro-

Site 2 Gly-Val -Ser (PI- Tyr -Ar g

Site 3 Tyr - GI y - Met - GIy - T hr S e r (PI Va I G lu Arg

-

- - -

FIG.3. Phosphorylation sites on pyruvate dehydrogenase.

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LESTER J . REED AND STEPHEN J . YEAMAN

Phosphorylation at site-1 proceeds markedly faster than at site-2 and -3, and phosphorylation at site-1 correlates closely with inactivation of E, . These findings indicate that phosphorylation site-2 and -3 do not play a physiological role in the inactivation of pyruvate dehydrogenase. Randle and co-workers (20, 2 1 ) reported that phosphorylation at site-2 and -3, in addition to site-1, on pig heart pyruvate dehydrogenase markedly inhibited the rate of its reactivation by pyruvate dehydrogenase phosphatase, and they proposed that multisite phosphorylation of pyruvate dehydrogenase may play a regulatory role. These results are at variance with those of Teague et al. (22), who observed that the presence of phosphoryl groups at site-2 and -3 on bovine kidney pyruvate dehydrogenase did not significantly affect the rate of reactivation of the enzyme by pyruvate dehydrogenase phosphatase. Phosphorylation of pyruvate dehydrogenase results in essentially total loss of its activity ( 3 ) . No allosteric activator of the phosphorylated enzyme has been reported. Phosphorylation of pyruvate dehydrogenase inhibits the decarboxylation of pyruvate (reaction 1 , Fig. 1) (23) and may also inhibit reductive acetylation of the lipoyl moiety (reaction 2, Fig. 1) (24). It appears that the E , a subunit catalyzes reaction 1 and that the p subunit catalyzes reaction 2 (23, 25).

111.

Pyruvate Dehydrogenase Kinase

A. ISOLATION AND PHYSICOCHEMICAL PROPERTIES Pyruvate dehydrogenase kinase has been purified about 2,700-fold to apparent homogeneity from extracts of bovine kidney mitochondria (26, 27). Kidney mitochondria contain at least four times as much pyruvate dehydrogenase kinase activity as heart mitochondria and are the preferred source for isolation of the kinase. Nevertheless, the amount of kinase present is small, and only 2-4 mg are recovered from about 12 kg of kidney cortex. The kinase is tightly bound to the E, core of the pyruvate dehydrogenase complex and copurifies with the complex. The complex is then resolved at pH 9.0 in the presence of 1'M NaCl to obtain an E,-kinase subcomplex. To separate E, and the kinase, the E,-kinase subcomplex is pretreated with dithiothreitol at pH 9.0, and then p-hydroxymercuriphenyl sulfonate is added. E, precipitates, and the kinase remains in solution. The pretreatment with dithiothreitol at alkaline pH is essential for subsequent resolution of the E,-kinase subcomplex in the presence of mercurial and presumably involves reduction of a disulfide bond or bonds. Highly purified preparations of the kinase show a doublet on sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (27). The two subunits have M,s of about 48,000 and 45,000, respectively. This finding, together with a sedimentation coefficient (sz0,J of 5.5 S, indicate that the kinase has the subunit

83

3. PYRUVATE DEHYDROGENASE

composition [email protected] the kinase comprises less than 5% by weight of the kidney pyruvate dehydrogenase complex, and because of interference by trace amounts of impurities, including products of limited proteolysis, it is difficult to detect with certainty the kinase doublet on SDS-polyacrylamide gels of the pyruvate dehydrogenase complex. The kinase doublet can be detected on SDSpolyacrylamide gels of the E,-kinase subcomplex, provided limited proteolysis is minimal. The turnover number (kcat) of pyruvate dehydrogenase kinase is about 32 min -

B. REGULATORYPROPERTIES Limited proteolysis with chymotrypsin selectively modified the kinase a subunit ( M , = 48,000) and was accompanied by loss of kinase activity. Limited tryptic digestion selectively modified the @ subunit ( M , = 45,000) without affecting kinase activity. These observations, together with the results of peptide mapping, indicate that the two subunits are distinctly different proteins. Kinase activity resides in the a subunit. The function of the p subunit remains to be established. An attractive possibility is that it functions as a regulatory subunit. Pyruvate dehydrogenase kinase requires Mg2 or Mn2 (apparent K, = 0.02 mM for both cations) (4). Kinase activity is stimulated by acetyl-CoA and by NADH, products of pyruvate oxidation, provided K or NH, ions are present (28,29);and kinase activity is inhibited by ADP and by pyruvate ( 4 , 30). ADP is competitive with ATP, and this inhibition apparentlv requires the presence of monovalent cation (31). The coenzyme, thiamin diphosphate, inhibits kinase activity, apparently as a result of binding at the catalytic site of pyruvate dehydrogenase and thereby altering the conformation about phosphorylation site- 1 so that the serine hydroxyl group is less accessible to the kinase (23).Treatment of highly purified preparations of the pyruvate dehydrogenase complex from bovine kidney and heart with an excess of N-ethylmaleimide resulted in a timedependent loss of endogenous kinase activity, but had little effect on the ability of the preparations to oxidize pyruvate (32). This inhibition was not reversed by dithiothreitol. Endogenous kinase activity was also inhibited by certain disulfides. This inhibition was reversed by dithiothreitol. 5,5’-Dithiobis(2-nitrobenzoic acid) was the most potent inhibitor, showing significant inhibition at 1 pM. It appears that pyruvate dehydrogenase kinase contains a thiol group (or groups) that is involved in maintaining a conformation of the enzyme that facilitates phosphorylation of its protein substrate. Modulation of kinase activity by thiol-disulfide exchange may be an important physiological mechanism. Pyruvate dehydrogenase kinase appears to be specific for pyruvate dehydrogenase. It exhibits little activity, if any, toward rabbit skeletal muscle phosphorylase b, glycogen synthase a , histones, or casein (4, 27). It has been suggested that the stimulatory effects of acetyl-CoA and NADH on +

+

+

+

84

LESTER J. REED AND STEPHEN J. YEAMAN

kinase activity are mediated through reduction and acetylation of the lipoyl moieties covalently bound to E, (33, 34). This suggestion is at variance with the findings of Reed et af. (35) with highly purified pyruvate dehydrogenase kinase and dephosphotetradecapeptide substrate. The rate of phosphorylation of the peptide substrate was stimulated by acetyl-CoA and NADH and inhibited by ADP and pyruvate. These results indicate that these effectors act directly on the kinase. Because pyruvate dehydrogenase kinase is tightly bound to E, and there are only about two kinase molecules per core of the bovine heart pyruvate dehydrogenase complex, it is not clear how these few kinase molecules can rapidly and completely inactivate a full complement of 30 E, tetramers (a2P2)attached to the E, core. Brandt and Roche (36) have made the interesting suggestion that the El molecules migrate on the surface of E, to the fixed kinase subunits.

IV. Pyruvate Dehydrogenase Phosphatase A.

ISOLATION AND F'HYSICOCHEMICAL PROPERTIES

Pyruvate dehydrogenase phosphatase has been purified to apparent homogeneity from bovine heart and kidney mitochondria (37, 38). Heart mitochondria contain at least three times as much phosphatase as kidney mitochondria and are the preferred source for isolation of the phosphatase. A key step in the purification procedure is affinity chromatography on E, coupled to Sepharose 4B. In the presence of Ca2 , the phosphatase binds to E, (1 7) and is subsequently released in the presence of ethylene glycol bis(P-aminoethyl ether)-N,N,N' ,"-tetraacetate (EGTA). The phosphatase has a s,~,,, of about 7.4 S and an M, of about 150,000 as determined by sedimentation equilibrium and gel-permeation chromatography. The phosphatase consists of two subunits with M,s of about 97,000 and 50,000 as estimated by SDS-polyacrylamide gel electrophoresis. Phosphatase activity resides in the M, = 50,000 subunit, which is sensitive to proteolysis (37).The phosphatase contains approximately 1 mol of FAD per mole of 150,000-protein. FAD is apparently associated with the M, = 97,000 subunit. The function of this subunit remains to be established. The k,,, of pyruvate dehydrogenase phosphatase with phosphorylated pyruvate dehydrogenase is about 300 min- I . It should be noted that the k,,, of the bovine pyruvate dehydrogenase phosphatase is about ten times the k,,, of pyruvate dehydrogenase kinase. The possible physiological significance of this difference needs to be evaluated. +

B.

REGULATORY PROPERTIES

Pyruvate dehydrogenase phosphatase requires Mg2 (apparent K , = 2 mM) or Mn2+ (apparent K , = 0.5 mM), when acting on both its physiological +

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3. PYRUVATE DEHYDROGENASE

substrate (phosphorylated E,) and phosphopeptide substrates (4, 39). At saturating Mg2+ concentration (about 10 mM), phosphatase activity toward its protein substrate is stimulated about 10-fold by micromolar concentrations of Ca2 , provided E, is present (17, 40, 41). However, phosphatase activity toward phosphopeptide substrates is not affected by Ca2 , whether or not E, is present (39). These observations indicate that Ca2+ is not directly involved in phosphatase catalysis. In the presence of Ca2+, the phosphatase binds to E,, and its apparent K,,, for phosphorylated E, is decreased about 20-fold, to 2.9 ph4 (17). 45Ca2 -binding studies have shown that the uncomplexed phosphatase binds one Ca2+ per molecule of M, = 150,000with a dissociation constant (K,) of about 8 pA4 (37). When both the phosphatase and E, are present, two equivalent and independent Ca2+-binding sites are detected with a Kd value of about 5 ph4. In the presence of 0.2 M KCl, which produces virtually complete inhibition of phosphatase activity, the enzyme binds only one Ca2 per molecule even in the presence of E,. These results are interpreted to indicate that pyruvate dehydrogenase phosphatase possesses an “intrinsic” Ca2+-binding site and that a second Ca2+-binding site is produced when both the phosphatase and E, are present. The second site is apparently altered by increasing the ionic strength, with a concomitant decrease in phosphatase activity. Localization of the second Ca2 binding site remains to be established. An attractive possibility is that this second site is at the interface between the phosphatase and E,, with Ca2+ acting as a bridging ligand for specific attachment of the phosphatase to E,. Alternatively, the second Ca2 -binding site may be on either the phosphatase or E,, produced by a conformational change in either enzyme when both are present. Favorable topographical positioning of the phosphatase and phosphorylated E, on E, apparently facilitates the Mg2 -dependent dephosphorylation. Preliminary studies on the binding stoichiometry of the phosphatase to E, indicate that there may be as few as 5 or 6 binding sites for the phosphatase on the 60-subunit E, (42). Because pyruvate dehydrogenase phosphatase and the pyruvate dehydrogenase complex are located in the mitochondria1 matrix, changes in free CaZ+ concentrations in the matrix could play an important role in regulation of phosphatase activity and hence pyruvate dehydrogenase complex activity (see Section V). At saturating concentrations of Mg2+ (10 mM) and Ca2+ (0.1 mM), the polyamines spermine, spermidine, and putrescine stimulated the activity of highly purified pyruvate dehydrogenase phosphatase 1.5- to 3-fold (43). Spermine was the most active of the polyamines. At a physiological concentration of Mg2+ (about 1 mM) (44, 45) and saturating Ca2+ concentration, the stimulation by 0.5 mM spermine was 4- to 5-fold; and at 0.3 mM M g 2 + , the stimulation was 20- to 30-fold. In the absence of Mg2+ or Ca2+, spermine had no effect. Thus spermine can spare but not completely replace Mg2 . Pyruvate dehydrogenase phosphatase exhibited slight activity with phosphorylated branched-chain aketoacid dehydrogenase complex (i.e., 0.5-1 .O% of the activity observed with +

+

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phosphorylated pyruvate dehydrogenase complex) (46).With this alternate substrate, the effect of spermine on pyruvate dehydrogenase phosphatase activity was similar to that observed with phosphorylated pyruvate dehydrogenase complex. In contrast with pyruvate dehydrogenase phosphatase, branched-chain aketoacid dehydrogenase phosphatase activity toward phosphorylated branchedchain a-ketoacid dehydrogenase complex was not affected by spermine. Branched-chain a-ketoacid dehydrogenase phosphatase exhibited about 10% of maximal activity with phosphorylated pyruvate dehydrogenase complex as substrate, but this activity was not affected by spermine. These results suggest that polyamines act, at least in part, directly on pyruvate dehydrogenase phosphatase. The stimulatory effect of polyamines on pyruvate dehydrogenase phosphatase activity may be relevant to the insulin stimulation of pyruvate dehydrogenase complex activity in adipose tissue (see Section V). Pyruvate dehydrogenase phosphatase activity is inhibited by NADH, and this inhibition is reversed by NAD+ (28). The phosphatase is inactive toward p nitrophenyl phosphate (37). It exhibits slight activity toward phosphorylase a from rabbit skeletal-muscle and phosphorylated branched-chain a-ketoacid dehydrogenase complex (Le., about 10% and 0.5- I%, respectively, of the activity observed with phosphorylated pyruvate dehydrogenase complex). Pyruvate dehydrogenase phosphatase activity is not inhibited by protein phosphatase inhibitor-1 or -2, and the activity is not affected by addition of highly purified calmodulin from porcine brain. It should be noted that the broad-specificity protein phosphatase (M,= 35,000) from rabbit liver cytosol shows significant activity in dephosphorylating and reactivating phosphorylated pyrvvate dehydrogenase complex from bovine kidney (35).

V. Regulation of Mammalian Pyruvate Dehydrogenase Complex

The pyruvate dehydrogenase system is well designed for fine regulation of its activity. Interconversion of the active and inactive phosphorylated forms of pyruvate dehydrogenase is a dynamic process that leads rapidly to the establishment of steady states, in which the fraction of phosphorylated E, can be varied progressively over a wide range by changing the concentration or molar ratios of effectors that regulate activities of the kinase and the phosphatase (28, 31). Thus, the steady-state activity of the purified pyruvate dehydrogenase system is affected markedly by varying the concentration of Mg2+ or Ca2+ and thereby changing the activity of the phosphatase. On the other hand, at optimum Mg2+ and Ca2 concentrations, the steady-state activity is affected markedly by varying the concentration of K + at a fixed ADP/ATP molar ratio or by varying the ADP/ATP ratio at a fixed concentration of K , and thereby changing the ac+

+

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active

a--

P hosphat ase

Kinase--

NADH

-

FIG. 4. Schematic representation of the covalent modification of pyruvate dehydrogenase and its control by effectors.

tivity of the kinase. The steady-state activity of the complex is also sensitive to the acetyl-CoAICoA and to the NADH/NAD molar ratios. Pyruvate dehydrogenase and its two converter enzymes, kinase and phosphatase, comprise a monocyclic interconvertible enzyme cascade (47). Fig. 4 summarizes the control of kinase and phosphatase activities by effectors, observed with the purified pyruvate dehydrogenase system. The acute regulation of pyruvate dehydrogenase in response to hormonal and other influences is mediated by two regulatory mechanisms-namely, end-product inhibition by acetyl-CoA and NADH and reversible covalent phosphorylation. Longer-term regulation may also involve changes in the total amount of enzyme present in the cell. The extent of phosphorylation of the complex can be estimated by measuring the activity of the enzyme in initial fresh extracts and then after treatment of the extracts with preparations of pyruvate dehydrogenase phosphatase. Under appropriate conditions this latter treatment fully dephosphorylates and activates the phosphorylated form of the enzyme and allows determination of the total activity. Activity state is defined as the initial activity (i.e., that of the dephosphorylated form) expressed as a fraction of the total activity. Furthermore, the activity in intact mitochondria, isolated cells, and perfused organs can be measured by several methods, the most popular of which is by quantifying the release of [14C]C0, from added [l-'4C]pyruvate. Comparison of the flux through the enzyme with the activity state can also give an indication of the extent of end-product inhibition operating in the cell. Possibly because of the availability of such methods, relatively few studies have been made using incubation of cells, tissues, etc. with [32P]Pito quantitate directly the phosphate content of the E,a subunit of the complex. In eukaryotic cells the pyruvate, a-ketoglutarate, and branched-chain a-ketoacid dehydrogenase complexes are located in mitochondria, within the inner +

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membrane-matrix compartment. Because of this localization of pyruvate dehydrogenase phosphatase and its sensitivity to Ca2+ ions, its activity can be regulated by changes in free Ca2+ concentration in the mitochondrial matrix. Using Ca2 -EGTA buffers with mitochondrial extracts and uncoupled mitochondria, Denton and McCormack (48) estimated that half-maximum activity of pyruvate dehydrogenase phosphatase (and of two other intramitochondrial Ca2 -sensitive enzymes, a-ketoglutarate dehydrogenase and NAD -linked isocitrate dehydrogenase) was obtained at a calculated free Ca2+ concentration of about 1 pM. Furthermore, these workers and Hansford (49) showed, using coupled mitochondria, that in the presence of physiological concentrations of Mg2 and Na+ ions the activity state of pyruvate dehydrogenase could be varied bv changes in extramitochondrial Ca2 in the concentration range 0.1- 1 pM. However, Williamson and co-workers (50),using a null point titration method, estimated that the concentrations of free Ca2 in the matrix of rat liver and heart mitochondria are about 9.7 and 5.7 @, respectively. These latter results would indicate that pyruvate dehydrogenase phosphatase is saturated with Ca2 over the physiological range if the Ao.5 value is 1 pM or less. It seems possible that this latter for the phosphatase (37). In view value is low, in view of Kd values of 5-8 of these discrepancies, a clear role for Ca2+ in the regulation of pyruvate dehydrogenase phosphatase has yet to be established (51). The activity state of the pyruvate dehydrogenase complex in fed rats varies between tissues, ranging from 0.2 to 0.7 (52). In catabolic states such as diabetes and starvation there is a marked decrease in the activity state of the complex in heart, liver, and kidney (53, 54). This partly results from increased oxidation of fatty acids and ketone bodies, causing increased intramitochondrial ratios of NADH/NAD and acetyl-CoA/CoA, which in turn stimulate pyruvate dehydrogenase kinase (28). inhibitors of fatty acid oxidation have been shown to reverse effects of starvation and diabetes on the complex (55). However, there is evidence for an additional mechanism whereby the complex in heart is inhibited during starvation and diabetes. Randle and co-workers (56, 57) have shown that pyruvate dehydrogenase kinase activity in heart mitochondria from diabetic rats is approximately 3-fold higher than in mitochondria from control animals. This increase in kinase activity cannot be accounted for by changes in the ratios of its allosteric effectors. Instead, it has been suggested that under these conditions there is increased synthesis of either a protein activator of the kinase or the kinase molecule itself, because the increase in kinase activity is blocked by inhibitors of cytoplasmic protein synthesis (58). Identification of the putative activator protein will obviously be an important advance. Decreased activity of pyruvate dehydrogenase phosphatase is also found in starvation or diabetes, but this effect apparently results from a change in the ability of the complex to act as substrate for the phosphatase as opposed to an effect on the phosphatase itself (59). One possible explanation of this observation is that increased occupancy of the second and +

+

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third phosphorylation sites resulting from the increased kinase activity (60, 61) causes inhibition of pyruvate dehydrogenase phosphatase (20). In contrast to other tissues, the activity state of pyruvate dehydrogenase in brain is essentially unaffected by starvation or diabetes, reflecting the key role of pyruvate oxidation in this tissue (6). The two best-studied acute effects of hormones on the activity of pyruvate dehydrogenase are those of positive inotropic agents such as adrenaline on the enzyme in heart and the effect of insulin on the enzyme in several tissues, but particularly in adipose tissue. Studies with perfused rat hearts have shown that adrenaline and other positive inotropic agents increase the initial activity of the pyruvate dehydrogenase complex about 4-fold (62, 63). This effect can be blocked by prior perfusion with the dye Ruthenium Red, which blocks mitochondrial Ca2+ ion uptake. These results suggest that the adrenaline effect may be due to increased transport of Ca2 ions into mitochondria and consequent stimulation of pyruvate dehydrogenase phosphatase (64). Studies with isolated heart mitochondria are consistent with this possibility (65). Positive inotropic agents increase the cytoplasmic concentration of Ca2 ions (66), and this presumably then leads to the increased levels of Ca2+ ions within mitochondria. Furthermore, Crompton el al. (67) have shown that perfusion of rat heart with adrenaline results in an increased total Ca2 content in mitochondria isolated from the perfused tissue. The increased activity state of the enzyme in skeletal muscle during and after exercise may also be due to increased activity of pyruvate dehydrogenase phosphatase resulting from increased levels of free Ca2 ions within mitochondria (68, 69). Similarly, stimulation of pyruvate dehydrogenase activity in liver by hormones such as vasopressin, angiotensin, and adrenaline (a-adrenergic action), which act via formation of inositol 1,4,5-trisphosphate and mobilization of cytoplasmic Ca2 (70), may be due to resultant increases in the intramitochondrial levels of free Ca2 . The most extensive studies on the hormonal control of pyruvate dehydrogenase are those on stimulation of its activity in fat cells by insulin. Physiological concentrations of insulin increase the activity state of the enzyme from 0.2-0.3 to 0.5-0.7. This increase is accompanied by net dephosphorylation of the E,a subunit, all three sites being dephosphorylated to approximately the same extent (71). This observation is in apparent conflict with the suggestion of Randle and co-workers (59), from work on diabetic rats, that the function of the second and third phosphorylation sites on pyruvate dehydrogenase is to inhibit dephosphorylation of the first site by pyruvate dehydrogenase phosphatase and hence to lock the enzyme into an inactive form. The effect of insulin on pyruvate dehydrogenase persists during isolation and incubation of mitochondria from treated fat pads (72, 73). Inhibition of the pyruvate dehydrogenase kinase by its known allosteric effectors can apparently be discounted because no changes in the intramitochondrial ratios of ATP/ADP, +

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NADH/NAD , and acetyl-CoA/CoA were detected as a result of insulin treatment (73).The effect of insulin is therefore presumably mediated via an increase in the activity of pyruvate dehydrogenase phosphatase. Elucidation of the mechanism by which insulin exerts its acute effects on key target enzymes, including pyruvate dehydrogenase, remains one of the major outstanding questions in the area of metabolic regulation. Considerable effort has been directed towards identification of an intracellular second messenger, generated at the plasma membrane in response to insulin, which leads to alterations in the phosphorylation state of target enzymes, but agreement has not yet been reached as to the identity of that putative second messenger. The situation concerning the effect of insulin on pyruvate dehydrogenase is even more complex, because any messenger generated must transmit its signal across the inner mitochondrial membrane. For many years a rise in the intramitochondrial concentration of free Ca2+ ions has been considered as a possible means by which insulin stimulates pyruvate dehydrogenase phosphatase and hence increases pyruvate dehydrogenase activity in adipose tissue (74). However, evidence from Marshall et al. (75) indicates that a rise in intramitochondrial Ca2+ ion concentration is not involved. Essentially, these workers found that the stimulatory effect on pyruvate dehydrogenase of extracellular Ca2+ ions can be blocked by Ruthenium Red, but that this compound does not block the effect of insulin on pyruvate dehydrogenase. Furthermore, the activities of the Ca2 -sensitive NAD 4socitrate dehydrogenase and a-ketoglutarate dehydrogenase are not increased in mitochondria from insulin-treated fat pads, and the increased activity of pyruvate dehydrogenase is retained when these mitochondria are subsequently depleted of CaZ . However, evidence has been presented that in adipose tissue insulin may exert effects on some of the enzymes of polyphosphoinositide metabolism (76-78). For example, insulin increases phospholipase C (the enzyme responsible for inositol 1,4,5-trisphosphate production) activity 2- to 3-fold in fat cells (78). Addition of phospholipase C to adipose tissue segments or adipocytes can mimic some effects of insulin, including stimulation of pyruvate dehydrogenase activity and effects on phospholipid metabolism, but not the insulin-like stimulation of glycogen synthetase activity (77). Furthermore, micromolar concentrations of exogenous inositol trisphosphate result in a several-fold increase in pyruvate dehydrogenase activity in permeabilized adipocytes (79). However, it has not been established that the effects of insulin on phospholipid metabolism are causally linked to its mechanism of action, especially as some of the cytoplasmic effects of insulin such as its potent antilipolytic action are not mimicked by other hormones that act via inositol trisphosphate (e.g., the a-adrenergic action of adrenaline) (80, 81). Another postulated second messenger for insulin is hydrogen peroxide. Evidence in support of this suggestion includes the observations that (a) insulin +

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stimulates an NADPH oxidase in adipocytes, both in intact cells and in isolated plasma membranes, leading to increased production of hydrogen peroxide (82), and (b) that added hydrogen peroxide can mimic some of insulin’s actions, including stimulation of pyruvate dehydrogenase activity (83). Addition of low levels of hydrogen peroxide to mitochondria isolated from adipocytes also causes stimulation of pyruvate dehydrogenase activity (84). The major weakness in the argument that hydrogen peroxide (or a related peroxide) may mediate insulin’s action is that no direct effect of peroxide has been demonstrated on any of the target enzymes or on the kinases and phosphatases that regulate the phosphorylation state of these enzymes. Evidence has accumulated indicating that a peptide or glycopeptide mediator (M,of 1000-2000) is released from plasma membranes by proteolytic action in response to insulin treatment, and that this mediator can mimic some of the acute effects of insulin on target enzymes such as glycogen synthetase and pyruvate dehydrogenase. A peptide mediator of insulin action has been reported in insulintreated muscle, where it can inhibit cyclic AMP-dependent protein kinase and stimulate phosphoprotein phosphatase(s), leading to increased activity of glycogen synthetase (85, 86). Incubation of plasma membranes from adipocytes (87, 88) and liver (89) with insulin leads to production of a mediator that can cause dephosphorylation and activation of pyruvate dehydrogenase in isolated mitochondria, apparently via stimulation of pyruvate dehydrogenase phosphatase (90). However, the observed changes in the activity state of pyruvate dehydrogenase are relatively small, and it has been pointed out that the mitochondria used in these studies were probably damaged or broken (73). It should also be noted that the assays were carried out in the presence of a Mg2+ concentration (50 @ that I is ) suboptimal for pyruvate dehydrogenase phosphatase (90). In view of the marked stimulation of pyruvate dehydrogenase phosphatase activity in v i m by polyamines at physiological Mg2 concentration (about 1 mM), it has been suggested that the putative insulin mediator may be polybasic in character (43)* It has also been suggested that more than one mediator is produced, a stimulator of pyruvate dehydrogenase activity being produced in response to low levels of insulin and an inhibitor of pyruvate dehydrogenase activity being released in the presence of higher levels of insulin (91).The stimulator and inhibitor can apparently be separated by high-voltage electrophoresis (92)or by utilizing differences in solubility in ethanol (93). Mediator from muscle that stimulates glycogen synthetase activity (via inhibition of cyclic AMP-dependent protein kinase) was originally shown to stimulate pyruvate dehydrogenase activity in mitochondria from adipocytes ( 9 4 , but subsequent purification has indicated that the mediator that inhibits cyclic AMP-dependent protein kinase is distinct from the one that stimulates pyruvate dehydrogenase (95). The relationship between the different mediators is of obvious interest. Despite intense +

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effort in several laboratories the structure of the postulated peptide mediator(s) of insulin remains elusive. To establish that these peptides (or glycopeptides) are the mediators of insulin’s action, their complete structure must be elucidated and a synthetic preparation of the peptide must be shown to possess insulin-like properties in different insulin-sensitive systems.

VI. Comparison of Properties of Mitochondria1 a-Ketoacid Dehydrogenase Kinases and Phosphatases

Only two mitochondria1 enzymes have been shown to be regulated by reversible phosphorylation-namely, pyruvate dehydrogenase and branched-chain aketoacid dehydrogenase. Pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase have been purified to homogeneity and their structure and regulation have been studied in detail. The branched-chain a-ketoacid dehydrogenase kinase has not yet been obtained in a homogeneous state. It is tightly bound to the branched-chain a-ketoacid dehydrogenase complex and copurifies with the complex (96, 97). Branched-chain a-ketoacid dehydrogenase kinase activity is inhibited by ADP, branched-chain a-ketoacids, and thiamin diphosphate (98, 99). These effects are analogous to those observed with pyruvate dehydrogenase kinase, which is inhibited by ADP, pyruvate, and thiamin diphosphate. Unlike pyruvate dehydrogenase kinase, which is inhibited by CoA and NAD+ and stimulated by acetyl-CoA and NADH, the branched-chain aketoacid dehydrogenase kinase from ox kidney is apparently unaffected by CoA, NAD , isovaleryl-CoA, or NADH (98). The branched-chain a-ketoacid dehydrogenase kinase from rabbit liver is inhibited slightly by isovaleryl-CoA and more effectively by acetoacetyl-CoA (40% at 0.01 mM) (99). Branched-chain a-ketoacid dehydrogenase phosphatase has been purified approximately 8,000-fold from extracts of bovine kidney mitochondria (46). The highly purified phosphatase has an apparent M, of about 460,000. In contrast to pyruvate dehydrogenase phosphatase, which requires Mg2 or Mn2 and is markedly stimulated by Ca2 , the branched-chain a-ketoacid dehydrogenase phosphatase is active in the absence of divalent cations. Polyamines markedly stimulate pyruvate dehydrogenase phosphatase activity at physiological concentrations of Mg2 . By contrast, branched-chain a-ketoacid dehydrogenase phosphatase activity is not affected by polyamines. The latter phosphatase is inhibited by nucleoside di- and triphosphates and is stimulated by basic polypeptides. Both phosphatases are relatively specific for their physiological substrates. Thus, pyruvate dehydrogenase phosphatase exhibits only 0.5- 1 .O% of maximal activity with phosphorylated branched-chain a-ketoacid dehydrogenase complex as substrate. Branched-chain a-ketoacid dehydrogenase phosphatase from +

+

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bovine kidney shows about 10% of maximal activity with phosphorylated pyruvate-dehydrogenase complex as substrate. However. this latter activity is not likely to be physiologically significant because bovine kidney mitochondria1 extracts contain only about one-seventh as much branched-chain a-ketoacid dehydrogenase phosphatase activity as pyruvate dehydrogenase phosphatase activity (46).

REFERENCES 1. Reed, L. J. (1974). Acc. Chem. Res. 7, 40. 2. Oliver, R. M., and Reed, L. J. (1982). In “Electron Microscopy of Proteins” (J. R. Harris, ed.), Vol. 2, p. 1. Academic Press, London. 3. Linn, T. C., Pettit, F. H., and Reed, L. J. (1969). PNAS 62, 234. 4. Hucho, F., Randall, D. D., Roche, T. E., Burgett, M. W., Pelley, J. W., and Reed, L. J. (1972). ABB 151, 328. 5 . Denton, R. M., Randle, P. J., Bridges, B. J., Cooper, R. H., Kerbey, A. L., Pask, H. T., Severson, D. L., Stansbie, D., and Whitehouse, S. (1975). Mol. Cell. Biochem. 9, 27. 6. Wieland, 0. H. (1983). Rev. Physiol. Biochem. Pharmacol. 96, 124. 7. Randle, P. J., Fatania, H. R., and Lau, K. S. (1984). Mol. Aspects Cell. Regul. 3, 1. 8. Bleile, D. M., Munk, P., Oliver, R. M., and Reed, L. J. (1979). PNAS 76, 4385. 9. Hale, G., and Perham, R. N. (1979). FEBS Len. 105, 263. 10. Kresze, G.-B., and Ronft, H. (1980). EJB 112, 589. 11. Bleile, D. M., Hackert, M. L., Pettit, F. H., and Reed, L. J. (1981). JBC 256, 514. 12. Perham, R. N., Duckworth, H. W., and Roberts, G. C. K. (1981). Nature (London) 292,474. 13. Packman, L. C., Perham, R. N., and Roberts, G. C. K. (1984). BJ 217, 219. 14. Hackert, M. L., Oliver, R. M., and Reed, L. J. (1983). PNAS 80, 2907. 15. Barrera, C. R., Namihira, G., Hamilton, L., Munk, P., Eley, M. H., Linn, T. C., and Reed, L. J. (1972). ABB 148, 343. 16. Wu, T.-L., and Reed, L. J. (1984). Biochemistry 23, 221. 17. Pettit, F. H., Roche, T. E., and Reed, L. J. (1972). BBRC 49, 563. 18. Yeaman, S. J., Hutcheson, E. T., Roche, T. E., Pettit, F. H., Brown, J. R., Reed, L. J., Watson, D. C., and Dixon, G. H. (1978). Biochemistry 17, 2364. 19. Sugden, P. H., Kerbey, A. L., Randle, P. J., Waller, C. A,, and Reid, K. B. M. (1979). BJ 181, 419. 20. Sugden, P. H., Hutson, N. J., Kerbey, A. L., and Randle, P. J. (1978). BJ 169, 433. 21. Kerbey, A. L., and Randle, P. J. (1979). FEBS Letr. 108, 485. 22. Teague, W. M., Pettit, F. H., Yeaman, S. J., and Reed, L. J. (1979). BBRC 87, 244. 23. Roche, T. E., and Reed, L. J. (1972). BBRC 48, 840. 24. Walsh, D. A., Cooper, R. H., Denton, R. M., Bridges, B. J., and Randle, P. J. (1976). BJ 157, 41. 25. Hubner, G., Neef, H., Schellenberger, A., Bernhardt, R., and Khailova, L. S. (1978). FEBS Lett. 86, 6. 26. Linn, T. C., Pelley, J. W., Pettit, F. H.,Hucho, F., Randall, D. D., and Reed, L. J. (1972). ABB 148, 327. 27. Stepp, L. R., Pettit, F. H., Yeaman, S. J., and Reed, L. J. (1983). JBC 258, 9454. 28. Pettit, F. H., Pelley, J. W., and Reed, L. J. (1975). BBRC 65, 575. 29. Cooper, R. H., Randle, P. J., and Denton, R. M. (1975). Nature (London) 257, 808.

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Pratt, M. L., and Roche, T. E. (1979). JBC 254, 7191. Roche, T. E., and Reed, L. J. (1974). BBRC 59, 1341. Pettit, F. H., Humphreys, J., and Reed, L. J. (1982). PNAS 79, 3945. Cate, R. L., and Roche, T. E. (1978). JBC 253, 496. Kerbey, A. L., Randle, P. J., Cooper, R. H., Whitehouse, S., Pask, H. T., and Denton, R. M. (1976). BJ 154, 327. 35. Reed, L. J . , Pettit, F. H., Yeaman, S. J., Teague, W. M., and Bleile, D. M. (1980). In “Enzyme Regulation and Mechanism of Action” (P. Mildner and B. Ries, eds.), p. 47. Pergamon, Oxford. 36. Brandt, D. R., and Roche, T. E. (1983). Biochemistry 22, 2966. 37. Teague, W. M., Pettit, F. H., Wu, T.-L., Silberman, S. R.,and Reed, L. J. (1982).Biochemistry 21, 5585. 38. Pratt, M. L., Maher, J. F., and Roche, T. E. (1982). EJB 125, 349. 39. Davis, P. F., Pettit, F. H., and Reed, L. J. (1977). BBRC 75, 541. 40. Siess, E. A., and Wieland, 0. H. (1972). EJB 26, 96. 41. Denton, R. M., Randle, P. J., and Martin, B . R. (1972). BJ 128, 161. 42. Wu., T.-L.(1982). Doctoral Dissertation, University of Texas at Austin. 43. Damuni, Z., Humphreys, J. S., and Reed, L. J. (1984). BBRC 124, 95. 44. Veloso, D., Guynn, R. W., Oskarsson, M., and Veech, R. L. (1973). JBC 248, 4811. 45. Garfinkel, L., and Garfinkel, D. (1984). Biochemistry 23, 3547. 46. Damuni, Z., Merryfield, M. L., Humphreys, J. S., and Reed, L. J. (1984). PNAS 81, 4335. 47. Stadtman, E. R., and Chock, P. B. (1977). PNAS 74, 2761. 48. Denton, R. M., and McCormack, J . G. (1980). FEBS Lett. 119, 1. 49. Hansford, R. G. (1981). BJ 194, 721. 50. Coll. K. E., Joseph. S. K., Corkey, B. E., and Williamson, J . R. (1982). JBC 257, 8696. 51. Reinhart, P. H., Taylor, W. M., and Bygrave, F. L. (1984). BJ 223, 1 . 52. Wieland, 0. H., Siess, E. A., Weiss, L., Loffler, G., Patzelt, C., Portenhauser, R., Hartmann, U., and Schirmann, A. (1973). Symp. Soc. Exp. Biol. 27, 371. 53. Wieland, 0. H., Siess, E., Schulze-Wethmar. F. H., Funcke, H.J., and Winton, B. (1971). ABB 143, 593. 54. Wieland, 0. H., Patzelt, C., and Loffler, G. (1972). EJB 26, 426. 55. Caterson, I. D., Fuller, S. J., and Randle, P. J. (1982). BJ 208, 53. 56. Hutson, N. J . , and Randle, P. J. (1978). FEBS Lett. 92, 73. 57. Kerbey, A. L., and Randle, P. J. (1981). FEBS Lett. 127, 188. 58. Kerbey, A. L., and Randle, P. J. (1982). BJ 206, 103. 59. Hutson, N. J . , Kerbey, A. L., Randle, P. J., and Sugden, P. H. (1978). BJ 173, 669. 60. Sale, G. J., and Randle, P. J. (1980). BJ 188, 409. 61. Sale, G. J . , and Randle, P. J . (1981). BJ 193, 935. 62. Hiraoka, T., DeBuysere, M., and Olson, M. S. (1980). JBC 255, 7604. 63. McCormack, J. G., and Denton, R. M. (1981). BJ 194, 639. 64. McCormack, J. G., and England, P. J. (1983). BJ 214, 581. 65. McCormack, J. G., and Denton, R. M. (1984). BJ 218, 235. 66. Williamson, J. R. (1975). Handb. Physiol.. Sect. 7: Endocrinol. 6, 605. 67. Crompton, M., Kessar, P., and Al-Nassar, I. (1983). BJ 216, 333. 68. Hennig, G., Loffler, G., and Wieland, 0. H. (1975). FEBS Lett. 59, 142. 69. Denton, R. M., and Halestrap, A. P. (1979). Essays Biochem. 15, 37. 70. Berridge, M. J., and Irvine, R. F. (1984). Nature (London) 312, 315. 71. Hughes, W. A., Brownsey, R. W., and Denton, R. M. (1980). BJ 192, 469. 72. Severson, D. L., Denton, R. M., Bridges, B. R., and Randle, P. J. (1976). BJ 154, 209. 73. Denton, R. M . , McCormack, J. G., and Marshall, S. E. (1984). BJ 217, 441. 30. 31. 32. 33. 34.

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95

Denton, R. M., and Hughes, W. A. (1978). Int. J . Biochem. 9, 545. Marshall, S. E., McCormack, J. G., and Denton, R. M. (1984). BJ 218, 249. Farese, R. V., Larson, R. E., and Sabir, M. A. (1982). JBC 257, 4042. Honeyman, T.W., Strohsnitter, W., Scheid, C. R., and Schimmel, R. J. (1983). BJ 212,489. Koepfer-Hobelsberger, B., and Wieland, 0. H. (1984). Mol. Cell. Endocrinol. 36, 123. Koepfer-Hobelsberger, B., and Wieland, 0. H. (1984). FEBS Lett. 176, 411. Garcia-Sainz, J. A., and Fain, J . N. (1980). BJ 186, 781. Lafontan, M., and Berlan, M. (1981). Trends Pharmacol. Sci. 2, 126. Mukherjee, S. P., and Lynn, W. S. (1977). ABB 184, 69. May, J. M., and de Haen, C. (1979). JBC 254, 9017. Paetzke-Brunner, I., and Wieland, 0. H. (1980). FEBS Lett. 122, 29. Lamer, J., Huang, L. C., Brooker, G., Murad, F., and Miller, T. B. (1974). FP 33, 261. Lamer, J., Galasko, G., Cheng, K., De-Paoli, A. A., Huang, L., Daggy, P., and Kellogg J . (1979). Science 206, 1408. 87. Seals, J. R., McDonald, J. M., and Jarett, L. (1979). JBC 254, 6991. 88. Seals, J. R., and Jarett, L. (1980). PNAS 77, 77. 89. Saltiel, A., Jacobs, S., Siegel, M., and Cuatrecasas, P. (1981). BBRC 102, 1041. 90. Popp, D. A., Kiechle, F. L., Kotagal, N., and Jarett, L. (1980). JBC 255, 7540. 91. Seals, J . R.,and Czech, M. P. (1981). JBC 256, 2894. 92. Cheng, K., Galasko, G., Huang, L., Kellogg, J., and Lamer, J. (1980). Diabetes 29, 659. 93. Saltiel, A. R., Siegel, M. I., Jacobs, S., and Cuatrecasas, P. (1982). PNAS 79, 3513. 94. Jarett, L., and Seals, J. R. (1979). Science 206, 1407. 95. Thompson, M. P., Lamer, J., and Kilpatrick, D. L. (1984). Mol. Cell. Biochem. 62, 67. 96. Fatania, H. R., Lau, K. S., and Randle, P. J. (1981). FEBS Lett. 132, 285. 97. Lawson, R., Cook, K. G., and Yeaman, S. J . (1983). FEBS Lett. 157, 54. 98. Lau, K. S., Fatania, H. R., and Randle, P. J. (1982). FEBS Lett. 144, 57. 99. Paxton, R., and Harris, R. A. (1984). ABB 231, 48. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86.

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Branched-Chain Ketoacid Dehydrogenase PHILIP J. RANDLE PHILIP A. PATSTON JOSEPH ESPINAL Nufield Deparrment of Clinical Biochemistry University of Oxford Oxford OX3 9DU, United Kingdom

I. Introduction ........................

.............. ....................................... Dehydrogenase Kinase Reactions ..........

97 100 100 100 101 103 103 104

C. Branched-Chain Ketoacid Dehydrogenase Phosphatase Reactions . . . . . . . . . . .............. D. Activator Protein ............................... IV. Biological Significance of Reversible Phosphorylation . . . . . . . . . . . . . . . . . . A. Activities in Tissues in Viv B. Activities in Tissues in Vifro C. Unresolved Problems ..... .............. D. General Conclusions .............. .............. V. Addendum ....................................... References ................ .............................

107 109 112 112 116 116 117 118 119

A. Discovery

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

In. Regulation by

1.

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

..........

Introduction

The branched-chain ketoacid-dehydrogenase complex of animal tissues (abbreviated to branched-chain complex) is a mitochondrial-multienzyme complex 97 THE ENZYMES,Vol. XVIII Copyright 0 1987 by Academic Press, Inc. All rights of reproduction in any form reserved

98

P. J . RANDLE, P. A . PATSTON, AND J. ESPINAL

located in the inner mitochondria1 membrane. It catalyzes a thiamin pyrophosphate (TPP)- and Mg2 -dependent oxidative decarboxylation of branched-chain ketoacids with the formation of branched-chain acyl-CoA and reduction of NAD to NADH [Eq. (1): R, and R, are alkyl groups defined in this section and +

+

0 RI\

II ,CH-C-COO-

R2

+ NAD + CoASH

-

R')CH-C

TPP,Mg2+

0

II -S2x 106

-

-

El (enzyme) Ez (enzyme) E3 (enzyme) E L (subunits)

s20, w=6 S s20, w=20 S

275,000 and 2X lo6 -

-

I10,OOO

110,000

190,000 -

-

-

46,000 37,000 52,000 55,000

47,000 37,000 5 1,000 -

Protein

a

P

E2 (subunits) E3 (subunits)

46,000 35,000 52,000 55,000

-

46,000 37,000 52,000 -

Walues are taken from (15-18, 21. 22). The value for E l (enzyme) is for free E, (activator protein) (see Section 111,C) and was obtained by gel filtration on Sephacryl S-300. Values for subunits are based on SDS-PAGE.

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

103

mitochondria may contain free El in addition to El that is tightly bound to E, in the complex (21). This is discussed in detail in Section III,D.

111.

Regulation by Reversible Phosphorylation

A.

DISCOVERY

Johnson and Connelly in 1972 (10) observed inhibition by ATP of branchedchain complex activity in damaged and permeabilized ox liver mitochondria. There was no suggestion that this inhibition was the result of phosphorylation and in view of the subsequent difficulty in demonstrating inactivation by phosphorylation in liver mitochondria of fed animals it is questionable whether phosphorylation was responsible for the loss of activity. In 1978 Parker and Randle (22) observed that the branched-chain complex activity of freshly prepared rat heart mitochondria was too low to account for rates of leucine oxidation in rat heart. This suggested the possibility of interconvertible active and inactive forms of branched-chain complex. This was confirmed when it was shown that in rat heart mitochondria incubated without respiratory substrate (to deplete ATP) activity of branched-chain complex was increased up to 20-fold. In extracts of such incubated mitochondria, ATP induced rapid inactivation of branched-chain complex; inactivation was inhibited by ketoleucine. Activation of branched-chain complex was also demonstrable in rat heart mitochondria incubated with respiratory substrates plus uncouplers of oxidative phosphorylation or with respiratory substrates plus ketoleucine. Mitochondria incubated with respiratory substrates alone maintained or acquired low activity of branched-chain complex. It was concluded that branched-chain complex may be inactivated by phosphorylation, that phosphorylation may be inhibited by ketoleucine, and that phosphorylated complex may be reactivated by dephosphorylation. At much the same time and independently Odessey and Goldberg published evidence for active and inactive forms of the complex and for inactivation by ATP in extracts of skeletal muscle (23). These observations explained why it had been difficult to detect branchedchain complex in muscle although it was known that muscle oxidizes branchedchain amino and ketoacids [for review, see Ref. (23)]. Evidence for phosphorylation of branched-chain complex was first obtained (1980-1981) in respiring mitochondria (with ’*Pi)and in mitochondria1 extracts (with [y-’*P]ATP) (24-26). It was shown by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) in Tris buffer (27) and autoradiography that 32P is incorporated into a protein corresponding in M,to the asubunit of E, and that this incorporation is inhibited by ketoleucine. The Laemmli method of SDS-PAGE (27) successfully resolved the phosphorylated asubunits of the branched-chain and pyruvate-dehydrogenase complexes. Phos-

104

P. J . RANDLE, P. A. PATSTON, AND I. ESPINAL

phorylation of the two subunits may be selectively inhibited with ketoleucine (branched-chain complex) or pyruvate (pyruvate-dehydrogenase complex). In a notable experiment Oddessy incorporated 32P into both complexes in rat kidney mitochondria and then purified the 32P-phosphorylatedbranched-chain complex to apparent homogeneity (24).In 1981, Fatania et al. (28)succeeded in copurifying ox kidney branched-chain complex and its intrinsic branched-chain dehydrogenase kinase to apparent homogeneity and showed that phosphorylation and inactivation are strictly correlated. Comparable observations for rat kidney and rabbit liver complexes were published in 1982 by Odessey (17) and Paxton and Harris (16). Early methods devised for purification of branched-chain complex from kidney or liver yielded preparations devoid of kinase and led to the conclusion (15, 20) that branched-chain complex is not regulated by reversible phosphorylation. The reasons are reviewed fully in Ref. (3),but, briefly, branchedchain kinase is lost when fractionation at pH 99% and allosteric activation of phosphorylated complex has not been described. More detailed examination of the relationship between phosphorylation and inactivation of ox kidney complex showed that this is sigmoid and that phosphorylation may continue after inactivation is complete. Both steady-state and dynamic methods of exploring this relationship have been described (31, 32). The results suggested the possibility of multisite phosphorylation, which was confirmed when it was shown that three phosphopeptides could be separated from tryptic digests of 32P-phosphorylatedox kidney complex (or of complex in rat liver, kidney, or heart mitochondria) by high-voltage paper electrophoresis at pH 1.9 (3, 31-33). The electrophoretic mobilities relative to N6dinitrophenyllysine (ox kidney complex) were 1.53 k 0.03 (TA), 1.07 k 0.02 (TB), and 0.65 ? 0.01 (TC) (mean f SE) in the studies of Lau et al. (31). The

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

105

corresponding phosphopeptides in the studies of Cook et al. (32) were described as T1 (= TA), T2 (= TB) and T3 (= TC). Relative rates of phosphorylation were TA > TB > TC and inactivation was correlated mainly (66%) with the appearance of TA. Multisite phosphorylation has been further clarified by amino acid sequence analysis of the tryptic phosphopeptides from ox kidney complex (34, 35). These studies have shown two sites of phosphorylation. The sites are frequently recovered in three tryptic phosphopeptides because the second site of phosphorylation is a seryl residue linked to arginine, and cleavage of -Arg-Ser(P)- by trypsin is generally slower than cleavage of -Arg-Ser-. The sequence of T2 (= TB) is shown in Fig. 2 and comprises 24 residues; T1 (= TA) is residues 1-14 of T2 and includes phosphorylation site-1; T3 (= TC) is residues 15-24 of T2 and includes phosphorylation site-2. When phosphorylation is confined to site- 1, Arg( 14)-Ser(15) is cleaved and site- 1 is recovered in residues 1- 14 (T 1 = TA). When site-1 and -2 are phosphorylated two phosphopeptides (TI = TA and T3 = TC) are only obtained when tryptic cleavage of Arg( 14)-Ser(P)(15) is complete; if tryptic cleavage is incomplete then T2 (= TB) is also present and three phosphopeptides are obtained. Inactivation of the complex during phosphorylation and reactivation of the complex during dephosphorylation appear to be correlated with occupancy of phosphorylation site- 1 (31-35). The function of phosphorylation site-2 (if any) is not known. 2.

Substrate Specificity

Ox kidney branched-chain dehydrogenase kinase phosphorylates branchedchain dehydrogenase with MgATP. Inactivation has been observed with preparations of ADP and GTP but these effects could be explained by their content of ATP (19). The K,,, for ATP (Table 111) was 12.6 I.M (19). With rabbit liver complex the K,,, for ATP was 25 pA4 (16). 3 . Regulation of Branched-Chain Dehydrogenase Kinase Reaction(s) Results from the principal studies of kinase regulation are shown in Table 111. The upper panel refers to studies with purified ox kidney complex (19); the lower panel to rabbit liver complex (36, 37). The studies with ox kidney complex were based on measurement of pseudo-first-order rate constants for ATP-dependent inactivation and were confirmed by measurements of 32P phosphorylation with Site I

site 2 -

(NHZ)ILE-GLY-HIS-HIS-SER(P)-THR-SER-ASP-ASP-SER-SER-ALA-TYR-ARG-SER(P)-VAL-ASP-GLU-VAL-ASN-TYR-TRP-ASP-LYS(COOH) 1 2 3 4 5 6 7 8 9 10 I 1 12 13 14 15 16 17 18 19 20 2 1 2 2 2 3 24

FIG. 2. Amino acid sequence of tryptic phosphopeptide T2 (32-35) [T2 = TB in Ref. (31)]from fully phosphorylated ox kidney branched-chain ketoacid-dehydrogenase complex. T1 (= TA) is residues 1-14; T3(= TC) is residues 15-24.

106

P. J. RANDLE, P. A. PATSTON, AND J. ESPINAL TABLE I11 BRANCHED-CHAIN KETOACIDDEHYDROGENASE KINASEREACTION, SUBSTRATE, AND INHIBITOR KINETICS

Compound

Function

K,, (N)

Substrate Inhibitor Inhibitor Inhibitor Inhibitor Inhibitor

12.62 I .O

K , (mM) and type of inhibition

Ox Kidney ComplexcJ

Mg ATP ADP Ketoleucine DL-Ketoisokucine Ketovaline TPP Compound Rabbit Liver Complexb Ketoleucine DL-Ketoisokucine Ketovaline a-Ketovalerate a-Ketoadipate n-Octanoate

140

(d)

0.07 0.5 2.5

0.5 2 0.5

0.27+0.03 (competitive) 0.48 t0 .0 6 (noncompetitive) 0.92t0.14 (noncompetitive) 8.9 k3.2 (noncompetitive) 0.004 t0.00 1 (uncompetitive)

-

-

Compound

Acetoacetyl-CoA Methylmalonyl-CoA Clofibrate Phenylpyruvate Dichloroacetate NADP Heparin +

140

(d)

0.01 0.2 0.33 I .7 3 1.5 12 plml

OFrom Ref. (19). Analysis based on pseudo-first-order rate constants for ATP-dependent inactivation [taken from Ref. (19)]. "From Refs. (16, 36, 37). Analysis based on protein-bound 32P after 20 min of incubation with [y -32PIATP. I,, is concentration required for 40% inhibition of incorporation at 75 W - A T P [taken from Refs. (36, 371.

[ Y - ~ ~ P I A TThe P . studies with rabbit liver complex were based on protein-bound 32P after 20 min of incubation with [y3*P]ATP. Inactivation correlates with phosphorylation of site- 1, whereas 32P incorporation may include variable amounts of site-2 phosphorylation (dependent on the degree of inactivation, see Section III,B,I). The kinase activity of preparations of ox kidney complex is much greater (>10-fold) than that of rabbit liver complex, thus allowing much shorter incubation times. Kinase reaction(s) were inhibited competitively by ADP (K;, ox kidney 270 pM; rabbit liver 130 tLM>. All three branched-chain ketoacids were inhibitors of the kinase reaction. With ox kidney complex inhibition was noncompetitive (for Kivalues see Table 111). With rabbit liver enzyme the kinetics of inhibition of the kinase reaction were more complex and results were given as Z40 (the concentration required for 40% inhibition of the kinase reaction) (see Table 111). Both studies showed relative inhibitor potency to be ketoleucine > ketoisoleucine > ketovaline. The study with ox kidney complex showed that the Ki values for

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

107

branched-chain ketoacids in the kinase reaction were much higher (20- to 400fold) than the corresponding K,,, values in the branched-chain-dehydrogenase complex reaction (19). This difference is also evident from studies with rabbit liver complex (16,36).The kinase reaction was also inhibited by TPP (ox kidney complex; see Table III), but no consistent effect of the other substrates for the holocomplex reaction (NAD+ and CoA) were seen (19,36). The products of the branched-chain holocomplex reaction (NADH, branched-chain acyl-CoA) have no consistent effects on the kinase reaction (19, 36). Inhibition of the rabbit liver branched-chain dehydrogenase kinase reaction has been observed with a-ketovalerate, a-ketoadipate, n-octanoate, acetoacetylCoA, methylmalonyl-CoA, clofibrate, phenylpyruvate, dichloroacetate, NADP, and heparin [see Table I11 and Refs. (36, 37)]. A wide range of other fatty acids, other metabolites, and other CoA thioesters (including acetyl-CoA) were without significant effect (19, 36, 37). 4. Molecular Aspects of Branched-Chain Kinase

Branched-chain kinase has not been separated from the complex and its M, and subunit composition are unknown. It is reported to be associated with the E, component of the complex and to remain attached to E, when El and E, components of the El-E, subcomplex are dissociated (38).

5. Studies in Mitochondria Both phosphorylation sites become phosphorylated when rat heart, kidney, or liver mitochondria are incubated with respiratory substrate and 32Pi, and it is known that phosphorylation is confined to the a-chain of the E, component (25, 26, 33). Inactivation of branched-chain complex by phosphorylation in mitochondria is inhibited by all three branched-chain ketoacids and their relative effectiveness is compatible with Ki values given in Table 111 (14, 39). C. BRANCHED-CHAIN KETOACID PHOSPHATASE REACTIONS DEHYDROGENASE The mitochondria1 branched-chain ketoacid dehydrogenase phosphatase (branched-chain phosphatase) has been detected (40) and purified (30). Detection and purification employed well-washed ox kidney mitochondria and cytosolic phosphatases were therefore unlikely to be present. Reactivation of phosphorylated purified branched-chain complex by dephosphorylation was first shown with a rat liver cytosolic phosphoprotein phosphatase (41). 1. Ox Kidney Branched-Chain Phosphatase Fatania et al. (40) first showed that branched-chain phosphatase is present in purified preparations of ox kidney branched-chain complex. The activity was low and tl,, for reactivation of phosphorylated complex ranged from 13 to 70

108

P. J . RANDLE, P. A. PATSTON, AND J . ESPINAL

min. Reactivation was measured in the presence of 0.5 mM ADP (formed by hydrolysis of ATP used in phosphorylation) and under these conditions phosReactivation was phatase activity required Mg2+ ( K , approximately 1 a). correlated with dephosphorylation and was inhibited completely by 50 mM NaF. No stimulation of reactivation by Ca2 was detected in contradistinction to pyruvate dehydrogenase phosphatase (42). Damuni et al. (30) have purified branched-chain phosphatase approximately 8000-fold from ox kidney mitochondria and to apparent homogeneity by fractional precipitation, ion-exchange chromatography (DEAE cellulose), and chromatography on ADP-Sepharose. The purified phosphatase exhibited an M , of about 460,000 by gel filtration on Sephacryl S-400 and at high dilution exhibited a M , of about 230,000. The results of SDS-PAGE were not given. Branchedchain phosphatase is clearly distinct from mitochondria1 pyruvate dehydrogenase 150,000; two subunits, M , 97,000 and 50,000). The phosphatase ( M , specific activity was 0.24 unit/mg protein (based on Pi release), and the phosphatase induced coordinated release of 3zPi and reactivation of 32P-phosphorylated ox kidney branched-chain complex. The phosphatase exhibited some activity with 32P-phosphorylatedpyruvate-dehydrogenase complex (about 10% of the activity with phosphorylated branched-chain complex). The highly purified phosphatase did not require Mg2+ in the absence of nucleotides. Phosphatase activity was inhibited by inorganic phosphate, pyrophosphate, and nucleoside di- and triphosphates (half-maximum inhibition 60-400 pA4) and the inhibition was reversed by Mg2+ [cf. Fatania et al. (40)]. Phosphatase activity was also inhibited by CoA and various acyl-CoA compounds and this inhibition was not reversed by Mg2+. There were no effects of nucleotides or of NAD or NADH. Poly-L-lysine, poly-L-arginine and histone H3 stimulated phosphatase activity but spermine and spermidine were inactive. Branched-chain ketoacids were without effect. There is perhaps no evidence for regulation by effectors that may be of physiological interest, except that inhibition by nucleoside di- and triphosphates in mitochondria may confer a requirement for Mg2+. Paul and Adibi (43, 4 4 ) described a protein factor in skeletal muscle and serum that may activate branched-chain phosphatase in liver mitochondria but its significance is not known. +

-

-

+

2. Other Phosphatases Pyruvate dehydrogenase phosphatase displays little if any activity towards branched-chain complex (30).Cook et al. (35)investigated dephosphorylation of phosphorylated ox kidney branched-chain complex by the catalytic subunits of protein phosphatase- 1 and -2A and protein phosphatase-2B (rabbit muscle) and rat liver protein phosphatase-2C. Phosphatase- 1 and -2B were essentially inactive. Phosphatase-2A dephosphorylated complex at approximately 40% of the rate at which it acts on glycogen phosphorylase, whereas phosphatase-2C de-

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

109

phosphorylated complex at approximately 25% of the rate with the a-subunit of phosphorylase kinase. The relative rates of dephosphorylation of the two sites in branched-chain complex (site-1 and -2) were 5 to 1 with phosphatase-2A and -2C. Relative rates of dephosphorylation of the two sites with mitochondrial branched-chain phosphatase have not been described.

D. ACTIVATOR PROTEIN Activator protein is the name given by Fatania et al. (45) to a protein in rat liver and kidney mitochondria that reactivates phosphorylated branched-chain complex without dephosphorylation. In order to place it in context the evidence for tissue-specific regulation is first reviewed. 1. Tissue-Specific Regulation

The early studies with mitochondria and mitochondrial extracts showed that active branched-chain complex is readily obtained from freshly prepared liver and kidney mitochondria (11-15, 23). Freshly prepared heart and skeletal-muscle mitochondria contained predominantly inactive complex which could be converted into active complex by incubation of mitochondria without substrate (22). Interconversion of active and inactive forms was readily demonstrable in heart and skeletal-muscle mitochondria; less readily in kidney mitochondria; but not in liver mitochondria unless incubated in hypotonic media or depleted of divalent metal ions (14, 25, 26, 46, 47). In extracts of muscle mitochondria ATP-dependent inactivation of branched-chain complex was rapid and essentially complete; in extracts of liver and kidney mitochondria inactivation was slower and incomplete (22, 26, 48). Subsequent experience with complex and kinase copurified from liver and kidney showed that ATP-dependent phosphorylation and inactivation was rapid and complete upon purification of the complex (16, 17, 19, 28-30). These observations suggested that a factor or factors may operate in liver and kidney mitochondria but not in heart or skeletalmuscle mitochondria to prevent inactivation of the complex by phosphorylation. The suggestion received further support from measurements of the concentrations of active and of total complex (sum of active and inactive forms) in rat tissue in vivo. In muscle approximately 90-95% of complex is in the inactive form, whereas in liver and kidney approximately only 10-50% of complex is in the inactive form in normal rats on a normal diet (49-51). 2. Discovery of Activator Protein

In 1982, Fatania et al. (45) observed that phosphorylated ox kidney branchedchain complex was rapidly reactivated by extracts of rat liver mitochondria from which branched-chain complex had been removed by sedimentation at 150,000 g for 2 h. The reactivation was instantaneous, not progressive, displayed the con-

110

P. J . RANDLE, P. A. PATSTON, AND J . ESPINAL

centration rate relationship of an activator, and was not associated with release of 32Pifrom 32P-phosphorylatedcomplex. Similar activity was detected in a comparable fraction of rat and ox kidney mitochondria but not in a comparable fraction of rat heart or skeletal-muscle mitochondria. The material was thermolabile, inactivated by trypsin, precipitated by (NH,)*SO,, and on gel filtration displayed an apparent M, > 100,000. It was termed activator protein. 3. Purification and Characterization of Activator Protein Activator protein has been purified > 1000-fold and to apparent homogeneity from rat liver mitochondria1 extracts by high-speed centrifugation, (NH,),SO, fractionation, and high-performance liquid chromatography (HPLC) on DEAE-5PW by Espinal et al. (21). SDS-PAGE showed two subunits ofM,47,700 and 36,300, which were indistinguishable from the M,of the a-and P-subunits of the E, component of branched-chain complex. Gel filtration on Sephacryl S-300 gave an apparent M, of 190,000 suggesting the possibility that activator protein (and E, component of the complex) may be a tetramer (a$,). The estimated M, (from the subunit M,)for a tetramer is 168,000 so if activator protein and E, are tetramers then they display axial asymmetry. Yeaman et al. (38) in studies with partially purified activator protein [(NH,),SO, fraction; approximately 1% pure by criteria in Ref. (21)] obtained four lines of evidence that activator protein is free El: (a)restoration of complex activity to E, E,; (b) inactivation after incubation with ATP and phosphorylated branched-chain complex; ( c ) inactivation by [-Y-~*P]ATP and E,-kinase complex associated with incorporation of 32P into a protein of M, 46,000 on SDS-PAGE; and (d)inhibition of activator protein by thiamine thio-thiazolonepyrophosphate. The first two of these points of evidence have been demonstrated with highly purified activator protein (21). Based on a M, of 168,000, for purified activator protein is