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) (26) is probably due to inhibition of substrate binding. At higher (>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 -S
t NADHz

+ C02

R2

in Section II,B]. Control of the branched-chain complex by reversible phosphorylation and the biological significance of this control is discussed in Sections 111 and IV. In order to put this discussion in context the biological significance of branched-chain amino acids and general aspects of the pathways of metabolic degradation are outlined in this introduction. In animals the branched-chain amino acids are the L-isomers of leucine [(CH,), CH CH, CH (NH,) COOH], isoleucine [CH, CH, CH (CH,) CH (NH,) COOH], and valine [(CH,), CH CH (NH,) COOH]. The a-keto (or 2-0x0) acids formed by transamination are referred to as ketoleucine (4-methyl-2-oxopentanoate), ketoisoleucine (~-3-methyl-2-oxopentanoate),and ketovaline (3-methyl-2oxobutyrate). In man and rat the three branched-chain amino acids are essential amino acids which means that dietary requirement at nitrogen equilibrium is determined by the degradation rate. Regulation of the reactions catalyzed by the branched-chain complex is important because they are rate limiting for degradation of branched-chain amino acids (i.e., at nitrogen equilibrium the activity of branched-chain complex determines dietary requirement). As a group branchedchain amino acids on a molar basis account for 45% of the total essential amino acid requirements in man. They are relatively common constituents of tissue proteins (14% of all amino acids in muscle, for example), contributing importantly hydrophobic side chains. In addition leucine has specific effects on metabolism. Leucine and ketoleucine stimulate insulin secretion; and leucine may activate protein synthesis and inhibit protein degradation by direct action on tissues and is also an activator of L-glutamate dehydrogenase. Autoregulation of the overall rate of protein degradation may therefore be effected in protein catabolic states by the circulating concentrations of leucine and ketoleucine. In normal animals the blood plasma concentrations of branched-chain amino and ketoacids are maintained relatively constant. In man protein meals may induce about a twofold increase in concentrations and comparable changes may be seen in protein catabolic states (e.g., starvation, diabetes, and trauma) ( 1 , 2). The importance of the activity of branched-chain complex in regulation of circulating concentrations of branched-chain amino and ketoacids is perhaps most

(I)

99

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

+ isovaleryl

leucine

ketoleucine

isoleucine

k e t o i s o l e u c t n e 3 3-methyl-butyryl

valine

ketovnline

transaminases

CoA

-b CoA

HMCCoA+

f

--+ a c e t y l

ketone bodies

CoA

i s o b u t y r y l C o b u c c i n y l CoA+glucose

branched c h a i n complex

various enzymes

FIG. 1 . Degradation of branched-chain amino acids in animal tissues.

obvious from findings in patients with Maple Syrup Urine Disease. This is an inborn error of human metabolism in which the branched-chain complex exhibits an approximately 100-fold increase in K , for branched-chain ketoacids as a result of (presumed) mutation in the E, component of the complex ( 5 ) . As a consequence the plasma concentrations of branched-chain amino and ketoacids are increased up to 70-fold (6). If the condition is not recognized in the newborn and treated by restriction of dietary intake of branched-chain amino acids, severe brain damage ensues and affected infants rarely survive beyond two years of age. The pathway of degradation ( 4 ) (summarized in Fig. 1) is initiated after cell membrane and mitochondrial transport (2, 3 ) by reversible transamination with 2-oxoglutarate, catalyzed by mitochondrial and cytosolic transaminases. In the rat there are at least three isozymes; two of them may accept all three branchedchain amino acids thus allowing (with branched-chain complex) for coordinated rates of degradation. The first irreversible reaction in the pathway is catalyzed by the branched-chain complex [Eq. ( l)] and the resultant branched-chain acyl-CoA are metabolized by further enzyme-catalyzed reactions to acetyl-CoA (ketoisoleucine), HMG-CoA (ketoleucine), and succinyl-CoA (ketovaline and ketoisoleucine). As a consequence of this pattern of degradation valine and isoleucine are potential precursors of glucose; and isoleucine and leucine are potential precursors of ketone bodies. The activity of branched-chain complex appears to be rate limiting for oxidation of branched-chain amino acids in extrahepatic tissues (i.e., transaminases may be in excess). Transamination is apparently rate limiting in liver. The activity of branched-chain complex may be rate limiting for oxidation of branched-chain ketoacids in muscle. In liver at higher rates of flux a reaction or reactions beyond this step may be rate limiting; NADH and branchedchain acyl-CoA may limit flux through the branched-chain complex by end product inhibition. More detailed accounts of the biological significance of branched-chain amino and ketoacids, the pathways of degradation, other component enzymes, Maple Syrup Urine disease, and a more complete bibliography may be found in Refs. (1-6).

100

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

II. Animal Branched-Chain Ketoacid-Dehydrogenase Complex

A. DISCOVERY The existence of a branched-chain ketoacid-dehydrogenase complex analogous to the pyruvate-dehydrogenase and 2-oxoglutarate-dehydrogenase complexes was assumed when evidence was obtained in 1955 for oxidative decarboxylation of branched-chain ketoacids to the corresponding branched-chain acyl-CoA (7). Connelly and co-workers (8-10) first obtained evidence (19681972) for branched-chain ketoacid dehydrogenase-complex activity in mitochondria towards ketoleucine, ketoisoleucine, and ketovaline, and its requirement for CoA, NAD+ , and TPP. These studies suggested that there were two such complexes, one active towards ketoleucine and ketoisoleucine and the other towards ketovaline. Subsequent studies first published in 1978 indicated that a single multienzyme complex in mitochondria of animal tissues oxidizes all three branched-chain ketoacids (11-16). The likelihood that a single complex may oxidize all three branched-chain ketoacids was also suggested by the biochemical findings in Maple Syrup Urine Disease (3, 5 , 6 ) . There is no explanation for the evidence for two complexes in the studies of Connelly and co-workers (8-10). Single complexes from ox kidney, ox liver, rat kidney, and rabbit liver have been purified to apparent homogeneity (15-28).

B.

SUBSTRATES AND COENZYMES

All purified complexes investigated require CoA, NAD , TPP, and Mg2 and utilize as ketoacid substrates, ketoleucine, ketoisoleucine, and ketovaline. Ox kidney complex also utilizes ketomethionine (the a-ketoacid corresponding to methionine), ketobutyrate, and pyruvate. Apparent K , values for ketoacids, coenzymes, and Mg2+, together with relative V,,, values for the ketoacids, are given in Table I (19). Pyruvate is a poor substrate with a high K,, and a low V,,,. For reasons that are not apparent, different studies have shown either rr-latively lower values (9-20 pl4) or relatively higher values (40-50 pl4) for apparent K,,, for branched-chain ketoacids. The holocomplex reaction is described by Eq. 1 (Section I) and the stoichiometries have been shown experimentally (20). Branched-chain complexes are inhibited by their principal end products NADH (competitive with NAD+) and branched-chain acyl-CoA (shown for isovaleryl-CoA: competitive with CoA) (11-15). The mechanism is presumed to involve reduction or reductive acylation +

+

101

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE TABLE 1

SUBSTRATES A N D COENZYMES, BRANCHELKHAIN KETOACII,-DEHYI,KO~~NASE COMPLEX, KINETIC CONSTANTS"

K,

Substrate or coenzyme

(fl)

Apparent K,,,

9-50 10-37 13-40 56

Ketoleucine ~~-Ketoisoleucine Ketovaline Ketobutyrate Ketomethionine Pyruvate CoA NAD

I 0.67 1.26 0.67 0.26

110

7 15- 1000 2.5-10 40- I09 4.2 0.35- I .2

+

Mg

Relative V,,,,,

+

TPP

18-5 1 8-19

NADH Isovaleryl CoA

Walues are for complexes from ox kidney (15, 19), ox liver ( 1 2 ) , rabbit liver (16), rat liver (11, 13), and rat heart (14). The higher apparent K,,, values for ketoacids are from (11, 15) and the lower values are from (12-14. 16. 19). The reason for differences in apparent K,,, is not known. Inhibitions by NADH and isovaleryl CoA are competitive with NAD+ and CoA, respectively.

of lipoate (see Eqs. (2) to (3, Section 11,C). The K i values for NADH and isovaleryl-CoA are shown in Table I.

C. COMPONENT ENZYMES The branched-chain complex contains three component enzymes, E, (branched-chain ketoacid dehydrogenase or decarboxylase), E, (branched-chain acyltransferase), and E, (dihydrolipoly dehydrogenase; EC 1.6.4.3). The reactions they catalyze are shown in Eqs. (2) to (3,although the evidence is either 0

OH

OH

0 'H-C-hydrolipoyl-Ez

+ El .TPP

(3)

102

P. J. RANDLE, P. A. PATSTON, AND J. ESPINAL 0 RI.

0

I1 CH-C-hydrolipoyl-E2

It

Rl\

+ CoASH

,CH-C

Rz'

-SCoA

+ dihydrolipoyl-E*

(4)

R2

Dihydrolipoyl-E2

+ NAD+

lipoyl-E2

+ NADH

(5)

incomplete or not given (15, 17).Following purification branched-chain complexes are either deficient in or devoid of E, (E, is common to the branched-chain ketoacid-, pyruvate-, and 2-oxoglutarate-dehydrogenase complexes). The M, values of the holocomplex as purified (mainly subcomplexes of E, and E,), its component enzymes, and subunits are given in Table 11. The complex has been dissociated into its component enzymes by gel filtration at high ionic strength and alkaline pH (15) thus enabling subunit M,to be assigned. El is composed of two dissimilar subunits (a,p) and the overall M, by gel filtration may be consistent with a tetramer (a2@,)exhibiting axial asymmetry (see Section 111, D). E, is of highM, (gel filtration) and on electron microscopy is a cube, suggesting that it may be composed of 24 subunits (15).Lipoamide dehydrogenase (E,) is assumed from other studies to be a dimer. The stoichiometric relationships between E l , E,, and E, are not known. Electron microscopy indicates that E, forms the core of the complex and E, (and presumably E,) are distributed on the surface of E, (15). The association constants for the combination between E, and E, and E, and E, are not known. There is evidence that liver and kidney mitochondria but not muscle TABLE I1 MOLECULAR WEIGHTVALUESFOR BRANCHED-CHAIN KETOACID-DEHYDROGENASE COMPLEXES" rn, Values for animal tissue source

Ox kidney

Ox liver

Rabbit liver

Rat liver

Rat kidney

Holocomplex

s20, w=40 S

>2x 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 <6.8 is employed, and branchedchain dehydrogenase kinase activity is inhibited by a number of factors including 2-(N-morpholino) propane sulfonate buffers used in earlier studies. Suitable methods for copurification of branched-chain complex with intrinsic branchedchain kinase have been reported (16, 17, 28-30). B . BRANCHED-CHAIN KETOACID DEHYDROGENASE KINASEREACTIONS 1 . Phosphoiylation and Inactivation When purified complexes are incubated with [ Y - ~ ~ P I A phosphorylation TP and inactivation are closely correlated (16, 17, 28). Phosphorylation is confined to seryl residues in the a-chain of E, (16, 17, 19, 28). Inactivation is a pseudo firstorder reaction (19) and the apparent first-order rate constant at saturating ATP depends on the method of preparation of the complex. The half-time for inactivation ranges from 0.23 to 0.69 min [ox kidney complex, Ref. (28)]to 1.5 min [rat kidney complex, Ref. (17 ) ] , to 25 min [rat liver complex, Ref. (16)]. The extent of inactivation is >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-

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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-

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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 <5 X 10-9M (the concentration required for half-maximum reactivation of 10 milliunits of phosphorylated complex). Clearly, identity between activator protein and the E, component of the branched-chain complex can be established only by demonstrating identity of primary structure for their individual subunits. Activator protein (i.e., free El) could be, for example, an isozyme of the form of E, present in the complex. For this reason we prefer to continue to use the term activator protein.

+

4. Properties, Mechanism, and Biological Significance Activator protein reactivates phosphorylated complex completely at saturating concentration but is without demonstrable effect on the activity of dephosphorylated complex (21, 45). If activator protein is free E l it follows that reactions catalyzed by El in dephosphorylated complex are not rate limiting in the holocomplex reaction and that El in the complex is inactivated by phosphorylation.

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111

Activator protein is present in extracts of rat liver mitochondria prepared by freezing and thawing. In such extracts branched-chain complex is associated with the membrane fraction whereas activator protein remains in the supernatant after centrifugation at 33,000 g for 5 min and is recovered in full. This might suggest that complex is located in the inner mitochondria1 membrane (it can be solubilized with Triton X-100) and activator protein in the matrix. Activator protein fully reactivates particulate (i .e., membrane bound) phosphorylated complex but has no effect on dephosphorylated complex. Therefore if activator protein is free El it is surplus to requirement for the branched-chain complex, and moreover is bound by the complex with lower affinity than E l , which copurifies with the solubilized complex or is associated with complex in the inner mitochondrial membrane (21). This may mean, for example, that there is more than one category of binding site for E, in the complex or that activator protein is an isozyme of El that has a lower affinity for the complex. Activator protein has been detected in liver mitochondria (rat, rabbit, and ox) and kidney mitochondria (rat and ox), but not in rat heart or rat hind limb skeletal-muscle mitochondria (21, 45). In particular, activator protein was not detected in heart and skeletal-muscle mitochondria in which inactive (phosphorylated) complex had been totally converted into active (dephosphorylated) complex by incubation without substrate (3, 21). This observation is important because freshly prepared heart and skeletal-muscle mitochondria contain predominantly phosphorylated complex, and might conceivably contain inactive (phosphorylated) activator protein. This stricture is unlikely to apply to mitochondria in which phosphorylated complex has been converted into active complex by dephosphorylation. Phosphorylated rat heart complex, like phosphorylated bovine kidney complex, is fully reactivated by rat liver activator protein (3, 21, 45). These observations led Espinal et al. (21) to conclude that activator protein (i.e., free E, surplus to El in the complex) is present in liver and kidney mitochondria but not in rat heart and skeletal-muscle mitochondria. This led them to conclude further that coordination of the synthesis of E,, E,, and E, is different in muscle than in liver and kidney. As an alternative explanation, reduplication of the gene for E, , possibly with mutation in one of the genes, may conceivably have occurred with expression of both genes in liver and kidney but not in muscle. An important question is the extent to which activator protein (i.e., free El) may be responsible for tissue-specific regulation of interconversion of branchedchain complex as previously defined. Activator protein (i.e., free E,) has been envisaged as a buffering mechanism that maintains holocomplex in an active form in liver and kidney but not in muscle (21). No convincing evidence has been obtained for the presence of phosphorylated and inactive activator protein in liver mitochondria even when up to 75% of branched-chain complex is in the

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inactive form (21). To this extent the evidence is consistent with the interpretation that activator protein may mediate tissue-specific regulation but is incomplete in that a mechanism is lacking. It may be that activator protein is not phosphorylated until all complex is phosphorylated and that it inhibits phosphorylation of E, in the complex. Alternatively phosphorylated activator protein may be dephosphorylated more rapidly than phosphorylated complex by branched-chain phosphatase. These possibilities are amenable to investigation. A third possible mechanism is that an unknown factor prevents phosphorylation of both complex and activator protein in liver and kidney mitochondria but evidence is perhaps against this possibility (21). The mechanism by which activator protein reactivates phosphorxlated branched-chain complex is not fully known. Evidence given in Ref. (45) suggests that activator protein does not exchange with phosphorylated E, in the complex. This might suggest that binding to additional sites on E, restores activity to the complex. This is clearly a matter for further and more detailed investigation.

IV. Biological Significance of Reversible Phosphorylation

A.

ACTIVITIES IN TISSUESIN

VIVO

There are a number of problems involved in assaying the concentrations of branched-chain complex (active and inactive forms) in tissues in vivo. 1. The activities are low and it is necessary to concentrate the complex prior to assay. 2. It is necessary to prevent interconversion of phosphorylated and dephosphorylated forms and to prevent inactivation by proteolysis to which the complex is especially susceptible (49). 3. Activator protein may interfere with the estimation of inactive complex in liver and kidney. 4. Extramitochondrial oxidase may contribute to assays based on production of 14C02 from 14C-ketoacids. In the studies of Patston et al. (49) branched-chain complex was concentrated prior to assay by isolation within mitochondria. Total complex (i.e., sum of active and inactive forms) was assayed in such mitochondria by incubation in the absence of respiratory substrate to effect conversion of inactive complex into active complex. The active form of the complex was assayed by isolating mitochondria under conditions that prevented interconversion (100 mM NaF to inhibit

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

113

the phosphatase; 5 mM ketoleucine to inhibit the kinase). Studies have shown by happenstance that inclusion of NaF blocks the activation of phosphorylated complex by activator protein in subsequent assays (21). As a consequence assay under these conditions is apparently specific for the dephosphorylated form of the complex. Branched-chain complex was assayed with appropriate controls by NADH production in the holocomplex reaction after extraction from mitochondria. Activator protein was assayed in a supernatant fraction. Proteolysis was prevented with ox serum (assays of complex) or with further addition of benzamidine and phenylmethanesulfonyl fluoride (activator protein, which involved a further 2 h of centrifugation at 150,000 g). Gillim et al. (50) concentrated complex by high-speed centrifugation prior to assay, which may be expected to separate complex from activator protein. The principal findings from our own studies in the rat (21, 49) are summarized in Table IV (branched-chain complex) and Table V (activator protein). Briefly they indicate the following: 1. In normal rats fed a standard pellet diet (17% protein w/w) total complex (sum of active and inactive forms) is comparable in liver and kidney, somewhat lower in heart, and much lower in hind limb skeletal muscle (0.035 unit/g wet wt; not shown in Table IV). 2. The activity of activator protein is about three times greater in liver than in kidney (it has not been detected in heart and skeletal muscle). 3. The proportion of complex in the active form is 55% in liver and 71% in kidney, but only 5% in heart and <20% in skeletal muscle (below the lower limit of assay). 4. In vivo the activity of complex in liver and kidney may be higher than the values obtained from assay of extracts because of activation of phosphorylated complex within mitochondria by activator protein. 5. Low protein diets or 48 h of starvation decreases total complex in liver (but not in kidney or heart), decreases the proportion of complex in the active form in liver and kidney (but not in heart), and decreases the activity of activator protein in liver and kidney. 6. Alloxan diabetes decreases the proportion of active complex in liver and kidney but increases it in heart: it decreases total complex in heart but not in liver or kidney; and it decreases activator protein in liver but not in kidney. Overall, in rats fed a standard diet approximately 70% of active complex is in the liver. Low protein diets decrease the activity of branched-chain complex in liver by 90% and in kidney by 70%; the decreases may be larger if allowance is made for the potential contribution of activator protein in vivo but this is uncertain. Approximately 70% of the decrease in whole-body branched-chain complex

TABLE IV COMPLEXES; EFFECTSOF DIETAND ALLOXANDIABETEP ACTIVITIESOF BRANCHED-CHAIN Branched-chain complex (unitslg wet wt of tissueb) Rat Liver Rat (diet) Normal (standard) Normal (80% casein) Normal (9% casein) Normal (0% casein) Normal (48 h starved) Alloxan-diabetic (standard)

Total

0.8220.06 0.79-CO.07 0.32-CO.04" 0.2520.05c 0.61?0.04c 0.7020.06

Rat Heart

Rat Kidney % active

Total

5524.6 302 1.5 14-C1.oC 2 9 t 3.3c 10-C0.7c 1920.7'

0.7720.07 0.8720.07 0.77-CO.04 0.6620.03 0.61 20.02 0.82+0.05

OData are from Ref. (41). Values are mean 2 S.E.M. for not less than 6 animals. bA unit of enzyme activity forms 1 pb4 product/min. cP
% active

71 24.9 4723.8 21 2 2 . 8 ~ 26-t5.9c 32-C-2.6~ 3325.7c

Total

0.5720.04 0.5320.03 0.5720.04 0.62 20.03 0.55-CO.02 0.4220.02C

% active

5.320.5 8.921.8 10.92 1.9 6.022.2 7.221.0 1923.8~

4.

115

BRANCHED-CHAIN KETOACID DEHYDROGENASE TABLE V ACTIVITIES OF ACTIVATOR PROTEIN: EFFECT OF DIETA N D DIABETES" Activator protein (% of control) Rat diet Normal (standard) Normal (80% casein) Normal (9% casein) Normal (0% casein) Normal (48 h starved) Diabetic (standard)

Liver

Kidney

loo+ 14 9 8 2 12 1021 b 9+1 2824 30t2

100+21 173+27 3025 1424 21+4 b 15.5226

"Data are from Ref. (41). Results are mean + S.E.M. for not less than 6 animals. K0.5 for activator protein in controls were for liver, 1.8 mg mitochondrial protein; kidney, 5 . 4 mg mitochondrial protein. "P <0.01 for difference from control (normal rat fed standard diet).

activity occasioned by low protein diets is hepatic. Details of the calculations and assumptions are given in Ref. (49). The effect of low protein diet to decrease the total activity of branched-chain complex in tissues was first suggested by the work of Wohlhueter and Harper (52), which was carried out before the discovery of reversible phosphorylation. The effect of low protein diet to lower the proportion of complex in the active form was first described by Gillim et al. (50). Qualitatively there are no important differences between the results of Patston et al. (49) and those of Gillim ef al. (50) although there are some important quantitative differences. The effect of low protein diet to lower branched-chain complex activity in liver was also reported by Hauschildt et al. (53).The low proportion of complex in the active form in heart and skeletal muscle as compared with liver and kidney was also described by Wagenmakers et al. (51, 54). In contradistinction to the work of others (49,50)Wagenmakers et al. (54) find an increase in total activity of branched-chain complex in liver and kidney after starvation; this effect is not accompanied by any change in the percentage of active complex. The reason for the different results is not fully known but Wagenmakers et al. used a longer period of starvation (72 h as opposed to 48 h) and differences in the age of animals used could also be relevant. Further work in their laboratory indicates that exercise increases the percentage of active branched-chain complex in heart and skeletal muscle but not in liver and kidney.

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B . ACTIVITIES IN TISSUES IN VITRO In perfused heart the percentage of active complex is comparable to that seen in vivo provided that physiological respiratory substrates (glucose, palmitate, and pyruvate) are available (14, 36, 41, 55, 56). If such respiratory substrates are not available the proportion of active complex may increase (55, 56). There is evidence that to be effective glucose may require insulin presumably because insulin is required for glucose entry [compare data in Ref. (56) with data in Refs. (14, 36); see also Ref. 0 1 . Inhibitors of branched-chain kinase (n-octanoate, ketoleucine, ketovaline, clofibrate, dichloroacetate, and phenylpyruvate) increase the percentage of active complex (14, 36). It is of physiological interest that concentrations of ketoleucine, ketovaline, and leucine (which may be converted to ketoleucine in the heart) are effective (14, 36, 58). We are not aware of in v i m studies bearing on reversible phosphorylation in other tissues. C.

UNRESOLVED PROBLEMS

In studies in vivo, summarized in Section IV,A, low protein diets decreased total branched-chain complex activity in liver (sum of active and inactive forms) and activity of activator protein (free El or an isozyme of E l ) in liver and kidney. These decreases in activity are assumed to reflect decreases in concentration but this requires confirmation by immunoassay. If activator protein is free E, (as opposed to an isozyme) the question arises as to whether the decrease in the total activity of branched-chain complex is due solely to a decrease in the concentration of E, , or alternatively to a coordinated decrease in the concentrations of E, and E,. This may be resolved by immunoassay. The proportion of complex in the active form is decreased by low protein diets in tissues in which activator protein is present and in which it is reduced in concentration by low protein diets (i.e., liver and kidney). In muscle, which lacks activator protein, the percentage of active complex is not altered by low protein diets. This is currently the only evidence that activator protein may regulate reversible phosphorylation and mediate tissue-specific differences. More direct studies, in particular towards understanding the underlying mechanism, are needed. Current evidence suggests that tissue-specific differences in reversible phosphorylation are important in vivo in directing degradation of branched-chain ketoacids toward the liver. In particular the rate of transamination of branchedchain amino acids in muscle (and kidney) exceeds flux through the branchedchain complex; as a consequence branched chain ketoacids are released and may be removed from the circulation and degraded in the liver (59-61). The importance of the liver and kidney in the disposal of intragastrically administered ketoleucine has been shown in a quantitative study (62); 85% of the administered load was metabolized by liver and kidney to leucine or ketone bodies. Quan-

117

4. BRANCHED-CHAIN KETOACID DEHYDROGENASE

titative information is lacking regarding the contribution of different tissues to the overall degradation of branched-chain amino and ketoacids . Current knowledge of the regulation of branched-chain dehydrogenase kinase and branched-chain dehydrogenase phosphatase may be incomplete. The only known regulators of the kinase reaction of immediate physiological interest are ADP, branched chain ketoacids, and acetoacetyl-CoA. It is conceivable that effects of branched-chain ketoacids may be to adjust activity of the complex and thus rates of degradation in relation to diet. However, known tissue concentrations of branched-chain ketoacids in the rat (5-25 nmol/g wet wt) (58) may not be sufficiently high in relation to the Ki or I,, (Table 111) to be effective inhibitors of the kinase. Inhibition of the kinase by acetoacetyl-CoA is also difficult to understand physiologically because oxidation of fatty acids may promote phosphorylation of branched-chain complex in heart, not dephosphorylation. As mentioned in Section III,C, effectors of branched-chain phosphatase of obvious physiological importance have not been described. There is also a lack of convincing evidence for hormonal regulation of reversible phosphorylation in the branched-chain complex.

D. GENERAL CONCLUSIONS The results of studies in vivo, of reversible phosphorylation, and of the action of activator protein have indicated that the activity of branched-chain complex may be regulated by four mechanisms; they have also indicated that different types of regulation may vary in their significance in different tissues. In all tissues examined activity of the complex may be regulated by reversible phosphorylation of the a-chain of the El component. Phosphorylation is inactivating. In liver and kidney, but not in heart and skeletal muscle, phosphorylated complex may be reactivated without dephosphorylation by activator protein, which is either free El (i.e., El in excess of that required to saturate high-affinity sites on E2) or an isozyme of E, . In all tissues examined phosphorylated complex may be reactivated by dephosphorylation. In liver and kidney, but not in muscle, the activity of branched-chain complex may be regulated by induction or repression of the complex and of activator protein (i.e., free E, or E, isozyme). In all tissues examined the active (dephosphorylated) form of the complex is capable of being inhibited by its products, NADH (competitive with NAD ) and branched-chain acyl-CoA (competitive with CoA). Current evidence suggests that end-product inhibition may be important in liver when rates of flux through the branchedchain complex is high. The results of studies in vivo may suggest that the major role of reversible phosphorylation in rats receiving adequate dietary protein is to restrict the activity of branched-chain complex in muscle and thereby to facilitate hepatic (and possibly renal) oxidation of branched-chain ketoacids. Current evidence suggests +

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that the rate of transamination of branched-chain amino acids in muscle and kidney exceeds flux through the branched-chain complex. As a consequence branched-chain ketoacids may be released into the circulation and removed and degraded by the liver. It is not entirely clear why regulation may be directed towards effecting predominantly hepatic degradation. The only obvious explanation is the specialized function of the liver to form ketone bodies and glucose from degradation products of branched-chain ketoacids. Glucose and ketone bodies are fuels of more general use and notably for cells in the central nervous system and such a mechanism could be significant in maximizing survival during starvation.

V. Addendum Several studies done since this review warrant mention here. The branchedchain complex has been estimated in rat heart and rat liver mitochondrial extracts by rocket immunoelectrophoresis employing a rabbit polyclonal antibody to purified ox kidney complex. Western blots on SDS-gels of ox kidney complex, of E, and E, resolved from ox kidney complex, and of rat mitochondrial extracts, and other evidence, showed that this is an antibody to E,. There were two minor bands corresponding to one component of higher M, than E, and one component of lower M,.The immunoassay (ox kidney standards) gave estimates for total complex of 9.3 milliunits/mg protein for liver mitochondria and of 6.1 milliunits/mg for heart mitochondria (rats fed normal diet) (63). These corresponded closely to the highest estimates obtained by bioassay. With the immunoassay no decrease in total complex was detected in liver mitochondria from rats fed 0% casein diet whereas bioassay showed a 70% decrease in total complex [(49),see also Table IV]. Three possible explanations for this discrepancy between immunoassay and bioassay are as follows. Because immunoassay measured E, it is possible that the concentration of E, is decreased specifically by 0% casein diet (i.e., free E, is present). However, addition of ox kidney E, to extracts of liver mitochondria from rats fed 0% casein diet did not increase holocomplex activity, i.e., it did not reveal free E,. A second possibility is that the immunoassay is not specific for E, (e.g., it might include degradation products of E, which are not detected by bioassay). This possibility was not completely excluded. The third possibility is that the apparent decrease of total complex on 0% casein diet is due to failure to achieve full conversion into the active form in the bioassay. Evidence in support of this was obtained by more prolonged incubation of liver mitochondria for up to 240 min with 5 mM ketoleucine (inhibitor of branched chain kinase). After maximum activation total complex activities (milliunits/mg protein; mean SEM) were 9.2 & 0.12 (normal diet) and 8.9 0.19

*

*

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119

(0% casein diet). These results indicate that low protein diet has no significant effect on the total concentration of branched-chain complex (sum of active and inactive forms) in liver mitochondria. These studies show that the only effect of low protein diet is to decrease the proportion of complex in the active (dephosphorylated) form by at least 90% (63). In a more recent study the activity of branched chain kinase has been assayed in extracts of rat liver and heart mitochondria; in extracts of rat liver mitochondria subjected to gel filtration on Sephacryl S300; and in branched-chain complex purified to near homogeneity from rat liver mitochondria extracts (64). Branched-chain kinase activity was approximately three-fold greater in heart as compared with liver mitochondria1 extracts (normal rats, normal diet). Feeding rats 0% casein diet for 10 days lead to a four-fold increase in branched-chain kinase activity in liver (extracts, gel filtered extracts, purified complex); and to a two-fold increase in heart. Damuni et al. (65, 66) have isolated a heat stable inhibitor of the branched chain dehydrogenase phosphatase from bovine kidney mitochondria and purified it to apparent homogeneity. It is a protein of M, = 36,000 which exhibits noncompetitive inhibition with a Kiof approximately 0.13 nM. Its physiological significance has yet to be assessed.

ACKNOWLEDGMENTS The authors’ work is supported by grants from the Medical Research Council (UK) and the British Diabetic Association.

REFERENCES 1. Harper, A. E., Miller, R. H., and Block, K. P. (1984). Annu. Rev. Nurr. 4, 409. 2. Walser, M., and Williamson, J. R., eds. (1981). “Metabolism and Clinical Implications of Branched Chain Amino and Ketoacids.” Am. Elsevier, New York. 3. Randle, P. J., Fatania, H. R., and Lau, K. S. (1984). In “Enzyme Regulation by Reversible Phosphorylation-Further Advances” (P. Cohen, ed.), p. I . Elsevier, Amsterdam. 4. Meister, E. (1965). “Biochemistry of the Amino Acids,” 2nd ed., Vol. 2, p. 743. Academic Press, New York. 5. Chuang, D. T., Nui, W. L. and Cox, R. P. (1981). BJ 200, 59. 6. Dancis, J., and Levitz, M. (1978). In “The Metabolic Basis of Inherited Disease” (J. B. Stanbury, J . B. Wyngaarden, and D. S. Frederickson, eds.), 4th ed., p. 397. McGraw-Hill, New York. 7. Coon, M. J. (1955). In “Amino Acid Metabolism” (W. D. McElroy and B. Glass, eds.), p. 431. Johns Hopkins Univ. Press, Baltimore, Maryland. 8. Connelly, J. L., Danner, M. J., and Bowden, J. A. (1968). JBC 243, 1198. 9. Bowden, J. A,, and Connelly, J. L. (1968). JBC 243, 3526. 10. Johnson, W. A,, and Connelly, J. L. (1972). Biochemistry 11, 1967.

120 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35.

36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50.

51. 52. 53. 54. 55. 56. 57.

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

Danner, D. J., Lemmon, S. K., and Elsas, L. J. (1978). Eiochem. Med. 19, 27. Parker, P. J., and Randle, P. J. (1978). EJ 171, 751. Parker, P. J., and Randle, P. J. (1978). FEES Lett. 90, 183. Parker, P. J., and Randle, P. J. (1980). FEES Lett. 112, 186. Pettit, R. H., Yeaman, S . J., and Reed, L. I. (1978). PNAS 75, 4881. Paxton, R., and Harris, R. A. (1982). JEC 257, 14433. Odessey, R. (1982). EJ 204, 353. Heffelfinger, S. C., Sewell, E. T., and Danner, D. J. (1983). Biochemistry 22, 5519. Lau, K. S., Fatania, H. R., and Randle, P. J. (1982). FEES Lett. 144, 57. Danner, D. J., Lemmon, S. K., Besharse, J. C., and Elsas, J. (1979). JEC 254, 5522. Espinal, J., Patston, P. A,, Fatania, H. R., Lau, K. S., and Randle, P. J. (1985). EJ 225, 509. Parker, P. J., and Randle, P. J. (1978). FEES Lett. 95, 153. Odessey, R., and Goldberg, A. L. (1979). EJ 178, 475. Odessey, R. (1980). FEES Lett. 121, 306. Hughes, W. A,, and Halestrap, A. P. (1981). EJ 196, 459. Lau, K. S., Fatania, H. R., and Randle, P. J. (1981). FEES Lett. 126, 66. Laemmli, U. K. (1970). Nature (London) 227, 680. Fatania, H. R., Lau, K. S., and Randle, P. J. (1981). FEES Lett. 132, 285. Lawson, R., Cook, K. G., and Yeaman, S. J. (1983). FEES Lett. 157, 54. Damuni, Z., Merryfield, M. L., Humphreys, J. S., and Reed, L. J. (1984). PNAS 81, 4335. Lau, K. S., Phillips, C. E., and Randle, P. J. (1983). FEES Lett. 160, 149. Cook, K. G., Lawson, R., and Yeaman, S. J. (1983). FEES Lett. 157, 59. Cook, K. G., Lawson, R., and Yeaman, S. J. (1983). FEES Lett. 164, 85. Cook, K. G., Lawson, R., Yeaman, S . J., and Aitken, A. (1983). FEES Lett. 164, 47. Cook, K. G., Bradford, A. P., Yeaman, S. J., Aitken, A., Fearnley, I. M., and Walker, J. E. (1984). EJE 145, 587. Paxton, R., and Harris, R. A. (1984). AEE 231, 48. Paxton, R., and Hams, R. A. (1984). AEE 231, 58. Yeaman, S. J., Cook, K. G., Boyd, R. W., and Lawson, R. (1984). FEES Len. 172, 38. Parker, P. J. (1979). Ph.D. Thesis, University of Oxford. Fatania, H. R., Patston, P. A., and Randle, P. J. (1983). FEES Left. 158, 234. Harris, R. A,, Paxton, R., and Parker, R. A. (1982). EERC 107, 1497. Randle, P. J., Denton, R. M., Pask, H. T., and Severson, D. M. (1974). Eiochem. SOC. Symp. 39, 75. Paul, H., and Adibi, S. (1982). JEC 257, 12581. Paul, H., and Adibi, S. (1983). JEC 258. 11471. Fatania, H. R., Lau, K. S . , and Randle, P. J. (1982). FEES Lert. 147, 35. Patel, T. B., and Olson, M. S. (1982). Biochemistry 21, 4259. Aftring, R. P., May, M. E., Manos, P. N., and Buse, M. G. (1982). JEC 257, 6156. Odessey, R. (1980). EJ 192, 155. Patston, P. A,, Espinal, J., and Randle, P. J. (1984). EJ 222, 71 1. Gillim, S. E., Paxton, R., Cook, G. A,, and Harris, R. A. (1983). EERC 111, 74. Wagenmakers, A. J. M., Schepens, J. T. G . , Veldhuizen, J. A. M., and Veerkemp, J. H. (1984). EJ 220, 273. Wohlhueter, R. M., and Harper, A. E. (1970). JEC 245, 2391. Hauschildt, S., Leuthje, J., and Brand, K. (1981). J . Nutr. 111, 2188. Wagenmakers, A. J. M., Schepens, J. T. G., and Veerkamp, J. H. (1984). EJ 223, 815. Waymack, P. P., De Buysere, M. S., and Olson, M. S. (1980). JEC 255, 9773. Buxton, D. B., Barron, L. L., Taylor, M. K., and Olson, M. S. (1984). EJ 221, 593. Vary, T. C., and Randle, P. J. (1984). J. Mol. Cell. Cardiol. 16, 723.

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Acetyl-Coenzyme A Carboxylase ROGER W. BROWNSEY"

RICHARD M. DENTON?

*Department of Biochemistry University of British Columbia Vancouver, Brirish Columbia Canada V6T IWS fDepartment of Biochemistry University of Bristol Medical School Bristol BS8 ITD, United Kingdom

I. Introduction 11. Structural As A. Molecular Forms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Allosteric Regulators ............. ............. 111. Short-Term Hormonal Re with Persistent Changes in Acetyl-CoA Carboxylase Activity . . . . . . . . . . . . IV. Early Evidence for the Regulation of Acetyl-CoA Carboxylase by Reversible Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Effects of Hormones on the Level of Phosph Carboxylase within Intact Cell Preparations ............ VI. Protein Kinases That Phosphorylate AcetylVII. Protein Phosphatases That Act on Acetyl-CoA Carboxylase . . . . . . . . . . . . . VIII. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References .............................. .............

1.

123 125 125 128 130 134

135

138 141 142 143

Introduction

The de novo synthesis of fatty acids requires a supply of acetyl-CoA together with ATP and NADPH as shown in Eq. (1) for the synthesis of palmitic acid. 123 THE ENZYMES, Vol X V l l I Copynght 0 1987 by Academic Press, Inc All nghtc of reproduction in any form rererved

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ROGER W. BROWNSEY AND RICHARD M . DENTON 8 CH3CO-S-COA + 7 ATP + 14 NADPH + 14 H + + CH3(CH2),,COOH + 8 COASH + 7 ADP + 7 Pi + 14 NADP+ + 6 H2O

(1)

This overall reaction is catalyzed in two stages by the enzymes acetyl-CoA carboxylase and fatty acid synthetase. In animal cells both possess complex structural and functional properties with multiple reactive sites on a single polypeptide chain. This chapter concentrates on the regulation of the specific activity of acetylCoA carboxylase from animal tissues, especially by phosphorylation. Many aspects of the structure, enzymology, synthesis, and breakdown are not fully covered in this chapter; fuller accounts can be found in a number of reviews (15 ) . In addition, an excellent review of the enzymology of fatty acid synthetase appeared in Volume XVI of this series (6). Acetyl-CoA carboxylase [more formally acetyl-CoA: carbon-dioxide ligase (ADP-forming), EC 6.4.1.21 catalyzes the ATP-dependent carboxylation of acetyl-CoA to produce malonyl-CoA which provides all but two (omega) of the carbon atoms to be incorporated into fatty acid products by fatty acid synthetase. The first description of acetyl-CoA carboxylase by Wakil (7) followed quickly after studies that demonstrated that bicarbonate (8, 9 ) and biotin (10) were required for the reaction. The biotin prosthetic group is attached by linkage of its valeric acid side chain to an €-amino lysine residue of the apoenzyme and plays a central role in both “half-reactions” of acetyl-CoA carboxylase [and other carboxylases as reviewed in Refs. (11, 12)].The overall enzyme-catalyzed reaction proceeds in a strictly ordered sequence involving carboxylation of biotin-enzyme using bicarbonate and the coupled hydrolysis of ATP followed by transfer of the carboxyl residue onto acetyl-CoA. The evidence for the overall reaction sequence has been discussed fully elsewhere ( I ) . The generally accepted bi-bi-uniuni ping-pong mechanism is illustrated in Eq. (2). ATP HCO; Enzyme-biotin

ADP+Pi

acetyl-CoA Enzyme-biotin

malonyl-CoA Enzyme-biotin

(2)

I

C0;

The precise mechanisms involved in the carboxylation and carboxytransferase reactions of biotin are a matter of some debate (13, 14). The existence of two active sites which must be serviced by a common prosthetic group raises the intriguing problem of the mobility of the common reactive species. The biotin “arm” has the potential to swing between sites as much as 25 A apart but NMR and ESR studies indicate that the movement involved is probably less than 10 A (15, 16).

The overall reaction catalyzed by acetyl-CoA carboxylase is often designated, rather vaguely, as the rate-limiting or the major rate-limiting enzyme in fatty acid

5 . ACETYL-COA CARBOXYLASE

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synthesis. It would seem useful to summarize the major reasons why the activity of the enzyme may be important in the regulation of fatty acid synthesis. 1. Malonyl-CoA has no substantial metabolic fate other than to be utilized as substrate by fatty acid synthetase, thus acetyl-CoA carboxylase commits carbon towards fatty acid synthesis. 2. The equilibrium constant, based on that for propionyl-CoA carboxylase (12), is probably in the range 100-1000. This is orders of magnitude larger than the mass action ratio in fat and liver cells which may be estimated to be between lo-' and lop4 based on determinations of the intracellular concentration of reactants and products of acetyl-CoA carboxylase (17-19) and assuming an intracellular concentration of bicarbonate ions of about 10 mM. The step catalyzed by acetyl-CoA carboxylase thus does not approach equilibrium in the cell. 3. Alterations in rates of fatty acid synthesis in rat liver associated with starvation, refeeding, and exposure to glucagon are linearly and positively correlated with the cell concentration of malonyl-CoA whereas the reverse is broadly the case with the cell concentration of acetyl-CoA (19, 20). 4. The activity of the enzyme is subject to regulation both in the long term through changes in rates of protein synthesis and breakdown and in the short term through changes in specific activity. Short term changes may be brought about by a number of potential allosteric regulators as well as by the reversible phosphorylation of several serine residues. This latter subject is the main topic of this review. Taken together, it seems entirely reasonable to assume that acetyl-CoA carboxylase plays an important role in the regulation of fatty acid synthesis. However, the qualitative and imprecise nature of the above arguments should be noted. There is a need for a more quantitative assessment of the regulatory role of the enzyme especially in comparison to the mechanisms that determine the supply of acetyl-CoA, for example through the regulation of the activities of glucose transport, phosphofructokinase, pyruvate kinase, and especially pyruvate dehydrogenase (21).

II. Structural Aspects and Regulation by Allosteric Effectors A. MOLECULAR FORMS Acetyl-CoA carboxylase derived from both avian and mammalian sources has been shown to be activated under conditions that promote the aggregation of dimers of the enzyme into linear, filamentous polymers (such as the presence of citrate); and to be inactivated under conditions such as high salt, low tem-

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perature, or in the presence of fatty acyl-CoA esters, malonyl-CoA, or MgATP plus bicarbonate which promote depolymerization (I -5). The possible physiological importance of this form of regulation is discussed in more detail in Section II,B. There is general agreement that the inactive dimer form has a molecular weight of about 500,000 and exhibits an szo,wof 10-16 S (22-29). This species may be further dissociated into two subunits (4-5 S) in the presence of SDS or other denaturing conditions. These subunits are apparently identical, each containing 1 mol/mol of biotin and active sites both for the ATP-dependent carboxylation of biotin and for the transfer of the carboxyl group to acetyl CoA producing malonyl-CoA. The M, of each subunit as determined by SDSpolyacrylamide gel electrophoresis has been reported to be in the range 215K260K (24, 26, 27, 30-33). Some of this variability may be explained by the use of different molecular weight standards and by the enzyme from different animals having slightly different molecular weights. For example, the enzyme from rabbit tissues appears to have a slightly greater molecular weight than that from rat tissues (28, 34). However, differences may also occur because the enzyme is extremely susceptible to limited proteolysis especially during isolation from liver by conventional procedures. Such procedures (typically a combination of ammonium sulphate precipitation, calcium phosphate, DEAE-cellulose, and Sepharose-2B column fractionations) resulted in enzyme preparations which gave multiple protein bands on SDS-polyacrylamide gel electrophoresis. For example, 140K, 130K, and 117K for the avian enzyme (35) and 230K, 215K, 125K, and 118K for the rat liver enzyme (36).Such preparations were found to have a specific activity of between 8-15 unit/mg protein. However, a substantially different view has subsequently emerged in studies that employed tissues with less intrinsic proteinase activity such as mammary gland (27-29, 33) and white adipose tissue (37), and/or rapid purification procedures including polyethylene glycol precipitation (27), citrate precipitation ( 3 3 , and Sepharose-avidin chromatography (23, 24. 30) together with a suitable cocktail of proteinase inhibitors. These preparations give essentially a single band on SDS-polyacrylamide gel electrophoresis with an apparent M, of 230K to 260K and have a considerably lower specific activity of the order of 1-5 unit/mg/protein. The activating effects of limited proteolysis have been directly investigated in a number of studies (24, 31, 38). For example, Song and Kim (24) compared the properties of rat liver acetyl-CoA carboxylase prepared by Sepharose-avidin affinity chromatography with the properties of the enzyme prepared by conventional techniques. The former preparation led to a product with a subunit M, of 260K, specific activity 1.2 unit/mg and K , for acetyl-CoA of 80 pA4. Corresponding values for the product of the alternative procedure were 230K, 12 unit/mg and 8 pM, respectively. Guy and Hardie (38), have shown that controlled limited proteolysis of rabbit mammary-gland acetyl-CoA carboxylase

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with trypsin leads to cleavage of the native (subunit Mr 250K) polypeptide yielding a 225K product which has about twice the specific activity. Finally, it should be pointed out that when acetyl-CoA carboxylase is purified by the very widely employed combination of ammonium sulfate fractionation and Sepharose-avidin chromatography, its final product is nearly always contaminated with varying amounts of a protein with a subunit M, or 140K-150K. This is almost certainly pyruvate carboxylase (R. W. Brownsey and R . M. Denton, unpublished observations). Contamination can be minimized by limiting the breakage of mitochondria at the initial extraction stage. The recovery of enzyme activity through Sepharose-avidin chromatography is usually about 3060%. Although superficially this may appear entirely satisfactory, the possibility must be considered that not all forms of the enzyme are being recovered. This point is discussed in subsequent sections but it is pertinent to mention at this stage that avidin appears to bind the enzyme only when it is in the inactive dimeric form (39). The polymerized active form of acetyl-CoA carboxylase that is produced upon exposure of the enzyme to citrate appears to have a maximum szo,w of 30-60 S which corresponds to the aggregation of up to 20 inactive dimers (M,about 500K) to give filaments of up to 0.5 p.m in length. However, there is considerable evidence from sedimentation studies that intermediate polymeric forms may exist ( I , 25, 27,29, 39a) and it has been reported that the proportion of these are increased following treatment of purified avian-liver enzyme with isocitrate plus MgATP (40),exposure of rat fat cells to epinephrine (25), and phosphorylation of the rat-liver enzyme (41). These intermediate-size species may form in virro even in the presence of maximally stimulating concentrations of citrate and may exhibit submaximal enzyme activity (40, 41). It must be emphasized that the relationship between activity and polymerization is by no means straightforward. Extensive studies by Beaty and Lane (23, 42, 43) with the chicken-liver enzyme employed rapid-quench techniques and right-angle light scattering to follow the time courses of the changes in activity and polymerization on addition of citrate. These studies show that the increase in activity is faster than polymerization under all conditions and therefore precedes polymerization. Similar experiments have yet to be carried out on the enzyme from mammalian sources. The extent to which polymerization and depolymerization may occur within intact cells has been explored in a number of studies. Halestrap and Denton (44) found that after exposure of fat cells to insulin a higher proportion of acetyl-CoA carboxylase activity could be sedimented in a high-molecular-weight form. Recently similar findings have been obtained using fast-protein liquid chromatography on a Superose 6 column (442). Another approach has been to employ permeabilization of liver cells with digitonin (45, 46). With chick-liver cells the rate and extent of elution of acetyl-CoA carboxylase from permeabilized cells was found to be increased by agents that promote depolymerization such as

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ROGER W. BROWNSEY AND RICHARD M. DENTON

malonyl-CoA whereas the converse was found with citrate (45). However, in similar experiments with rat liver cells pretreated with insulin or glucagon, which alter enzyme activity, no changes in elution rates from permeabilized cells (46) or in the rate of sedimentation in a sucrose gradient (47) were found. Further studies into the extent of polymerization of acetyl-CoA carboxylase within cells are certainly needed. It has been suggested that acetyl-CoA carboxylase may undergo reversible association with components of the microsomal fraction in rat liver under conditions of increased rates of fatty acid synthesis (48).However, in a separate earlier study no detectable association of the rat liver enzyme with the microsomal fraction was found (49) and we have found no association in rat epididymal adipose tissue (unpublished observations).

B. ALLOSTERIC REGULATORS Whatever the exact relationship between polymerization and activity, there is no doubt that the activity of acetyl-CoA carboxylase can be greatly altered in vitro by a number of regulators which would appear to be of potential physiological importance. The first to be recognised were the activating effects of citrate (and isocitrate) and the inhibitory effects of fatty acyl-CoA esters. With purified preparations of acetyl-CoA carboxylase, half-maximal effects of citrate are observed in the range 0.5-2.5 mM (1-3, 33, 43, 50, 51) and of palmitoyl-CoA and other fatty acylCoA esters at free concentrations of less than lOnM (2, 52). It should be noted that the effects of both citrate and fatty acyl-CoA may be considerably altered by other components usually present in the assay medium for acetyl-CoA carboxylase. For example, effects of citrate may be altered by the presence of divalent metal ions such as Mg2+ and those of fatty acyl-CoA by albumin. At first sight, the effects of citrate and fatty acyl-CoA would appear to be examples of feedforward activation and end-product inhibition respectively. However, it has proved rather difficult to establish irrevocably the extent to which alterations in the cytoplasmic concentration of these (and the other) regulators are important in the regulation of acetyl-CoA carboxylase activity. On the whole, there have been few reports of parallel changes in fatty acid synthesis and whole tissue or cytoplasmic concentrations of citrate. These include in the liver on refeeding previously fasted rats (53)and following the incubation of chicken liver cells with medium containing acetate, octanoate, or fructose (54), or rat liver cells with medium supplemented with lactate plus pyruvate (55). In many other conditions, no positive correlation between the concentration of citrate and flux through acetyl-CoA carboxylase has been evident (44, 56-62). For example, in rat epididymal adipose tissue, stimulation of fatty acid synthesis by insulin is associated with an unchanged or decreased

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tissue concentration of citrate whereas exposure of the tissue to adrenaline which leads to inhibition of fatty acid synthesis results in a marked increase in the concentration of citrate (44, 57-59). Moreover, incubation of this tissue with pyruvate or fluoroacetate leads to 8- to 10-fold increases in the concentration of citrate but little or no increase in acetyl-CoA carboxylase activity (44, 59). Glucagon and dibutyryl cyclic AMP inhibit fatty acid synthesis in both rat and chick liver cells but only in the chick cells incubated in medium containing glucose does this occur with a parallel decrease in cell citrate content (20, 6062). In both rat and chicken liver cells incubated with medium containing fructose, pyruvate, or lactate, the content of citrate increases (20, 60-62). An obvious weakness in such studies is that changes in the whole-cell content of citrate mav be a poor guide to changes in the cytoplasmic concentration of citrate to which acetyl-CoA carboxylase is exposed. However, the cytoplasm probably contains about 50% of cell citrate and changes do appear to broadly follow changes in whole-cell content (63-65). A possible role for fatty acyl-CoA esters in the regulation of acetyl-CoA carboxylase has obtained some support from observations of inverse changes in the whole-cell or tissue concentrations and the rate of fatty acid synthesis (44, 53, 54, 56-58). Examples include decreases in concentration when fatty acid synthesis is stimulated by insulin in rat epididymal fat cells (44, 57, 58) and by refeeding in the livers of previously fasted chickens (56) or rats (53). The concentration may also increase in chick liver when fatty acid synthesis is decreased following exposure to glucagon (54). However, a similar increase in the concentration of fatty acyl-CoA esters in rat liver cells incubated with glucagon was found to be transient and disappeared at later time points when inhibition of fatty acid synthesis was still apparent (20). It has to be emphasized that the interpretation of whole-cell measurements of fatty acyl-CoA is particularly hazardous because of the great uncertainty in the distribution of these esters, not only between different intracellular components (as with citrate) but also between aqueous and hydrophobic phases and specific binding sites within each compartment. Indeed, the whole-cell concentration of fatty acyl-CoA esters are in the range 50 phi-150 phi which is orders of magnitude greater than the range of concentrations found to result in reversible inhibition of purified acetyl-CoA carboxylase (44, 52). A number of other potential regulators of acetyl-CoA carboxylase activity have been described. These include coenzyme A, [which may markedly diminish the apparent K , for acetyl-CoA (5, 66)],AMP (67), GTP and other guanine nucleotides (68, 69), and polyphosphoinositides especially phosphatidylinositol 4 3 bis-phosphate (70, 71). So far there are no studies that indicate if changes in cellular concentrations of these potential regulators do indeed correlate with activities of acetyl-CoA carboxylase and rates of fatty acid synthesis.

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Short-Term Hormonal Regulation of Fatty Acid Synthesis Associated with Persistent Changes in Acetyl-CoA Carboxylase Activity

In mammals the principal sites of fatty acid synthesis are liver, white and brown adipose tissue, and lactating mammary tissue. Rates of fatty acid synthesis can be measured rather conveniently both in vivo and in v i m by following the incorporation of 3H from 3 H , 0 (72, 73). The short-term effects of hormones on the rates of fatty acid synthesis are summarized together with references in Table I (74-93). These effects can be observed within a few minutes of exposure of the tissue to hormones and thus are very unlikely to involve changes in the amount of enzymes. It is well established that insulin stimulates fatty acid synthesis in all four tissues although insulin effects, at least in percentage terms, are greatest in white and brown adipose tissue. In contrast, hormones that increase cyclic AMP result in the inhibition of fatty acid synthesis. The best-studied example is the effect of glucagon in liver preparations (20, 55, 63, 88-92) where inhibition of between 30-70% is observed. Glucagon and adrenaline (apparently acting through padrenergic receptors) also inhibit fatty acid synthesis in rat epididymal white fat cells, particularly in the presence of insulin (51, 57, 85, 94, 95) and it has been shown that fatty acid synthesis in interscapular brown adipose tissue of coldadapted rats is inhibited by up to 70% on injection of the rats with norepinephrine (82). Inhibition of fatty acid synthesis in mammary tissue under conditions of increased-cell cyclic AMP has not been demonstrated. Glucagon does not inhibit fatty acid synthesis in this tissue apparently as the result of a lack of receptors (85). Direct evidence that these opposing short-term effects of insulin and of hormones that increase cyclic AMP are brought about through changes in the activity of acetyl-CoA carboxylase has come from the recognition that persistent differences in the activity of acetyl-CoA carboxylase can be observed in extracts of cells previously exposed to the hormones. The first to be recognized was the increased activity which persists in extracts of rat epididymal adipose tissue from rats previously exposed to insulin (76). The increase was found to be maintained in extracts despite extensive dilution or incubation of extracts in the presence of serum albumin but was no longer evident after incubation of extracts with sufficient citrate to cause maximum activation (44, 76). Subsequently similar observations have been made with isolated rat fat cells (37, 51, 59, 78, 79), liver cells (88, 90, 91), and mammary tissue acini (96). However, the effects in liver cells are rather modest and in the case of mammary tissue have been observed only when acini from lactating starved or fat-fed animals are incubated with dichloroacetate (85, 86). Activation of acetyl-CoA carboxylase can also be demonstrated in vivo following manipulation of circulating plasma insulin concentra-

TABLE I SHORT-TERM EFFECTS OF HORMONES ON THE SYNTHESIS OF FATTYACIDSWHICH ARE ASSOCIATED WITH CORRESPONDING PERSISTENT CHANGES I N ACETYL-COACARBOXYLASE ACTIVITY IN VARIOUS RAT TISSUES Changes in fatty acid synthesis rates and activity of acetyl-CoA carboxylase Tissue preparation Epididymal white adipose tissue (in vivo) Epididymal white adipose tissue (intact tissue, in vitro) Epididymal white adipose tissue (isolated cells, in vitro) Interscapular brown adipose tissue (in vivo) Mammary tissue (in vivo) Mammary tissue (acini, in v i m ) Liver (in vivo) Liver (isolated cells, in v i m )

Increases

Decreases

Insulin (74) Insulin (37,44,50,51,59,76) Insulin (37,51,79) EGF(79a) Insulin (80,81) Insulin (81,83,84) Insulin (85,86) Insulin (74) Insulin (47,88) EGF(I14) Vasopressin (89) Angiotensin

Epinephrine (75) Epinephrine (25,77,78) Epinephrine (51,77,79) Glucagon (79) Nor-epinephrine (82)

}

-

Glucagon (87) Glucagon (26,88,90-92)

a-agonists (93)

In the examples given changes in fatty acid synthesis usually measured as the incorporation of 3Hfrom 3 H 2 0 into tissue fatty acids were found to be associated with parallel changes in acetyl-CoA carboxylase activity measured in subsequently prepared extracts. Acetyl-CoA carboxylase activity was usually measured in the absence of citrate or in the presence of a non-saturating concentration. EGF: epidermal growth factor.

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ROGER W. BROWNSEY A N D RICHARD M . DENTON

tions with, for example, injections of antiinsulin serum or glucose, in both white (74) and brown adipose tissue (83,in lactating mammary tissue (83, 84) and rather less convincingly in liver (74).Epidermal growth factor (EGF) appears to have similar effects to insulin in both fat and liver (79a, 114). In contrast to the effects of insulin, treatment of tissues with hormones that give rise to increases in cyclic AMP and inhibition of fatty acid synthesis result in decreases in the activity of acetyl-CoA carboxylase. The first suggestion for this came from studies on the effects of dibutyryl cyclic AMP on rat liver slices (97). Subsequently, diminished activity has been observed in extracts of rat epididymal fat pads (25, 51, 77) and fat cells treated in vitro with adrenaline or isoproterenol (51, 77, 79) or with glucagon (79). Injection of rats with epinephrine or norepinephrine have been shown to result in decreases in activity in rat epididymal white adipose tissue (75) and interscapular brown adipose tissue (82). Similar decreases have been found in rat liver cells following exposure to glucagon (26, 88, 90-92). The sensitivity of the enzyme to activation by citrate is also decreased after exposure of tissues to hormones that increase cell cyclic AMP (77, 79, 92). It is noteworthy that no persistent changes in acetyl-CoA carboxylase activity have been shown in extracts of chick liver cells previously exposed to glucagon (61-63). The effects on both fatty acid synthesis and the activity of acetyl-CoA carboxylase of hormones (a-adrenergic agonists, vasopressin, and angiotensin) which act in the liver primarily by increasing polyphosphoinositide breakdown and hence the cytosolic concentration of Ca2+ appear to be rather contradictory. Inhibition of fatty acid synthesis has been found in mouse liver perfused with medium containing angiotensin or vasopressin (98) and in rat liver cells by norepinephrine apparently acting through a-receptors (93). In the latter case it was shown that a decrease in the activity of acetyl-CoA carboxylase persisted in subsequently prepared extracts. However, vasopressin and angiotensin have also been found to stimulate fatty acid synthesis and acetyl-CoA carboxylase activity in rat liver cells (89). Further studies into the effects of these hormones are required but it is possible that the apparent discrepancies in the findings are explained (in part) by differences in incubation conditions. All the above studies have followed the acetyl-CoA-dependent fixation of I4C from I4C-labeled bicarbonate into acid- and heat-stable products as a measure of acetyl-CoA carboxylase activity. There has been some controversy as to the extent to which pyruvate carboxylase activity might interfere. This appears to be a potential problem in liver extracted under conditions where mitochondria are broken and assays are carried out with high concentration of extract (47, 99). However, under standard conditions of assay, it has been argued that only a small fraction of the I4C measured is the result of pyruvate carboxylase activity (46,47, 79a, 100). In adipose tissue the problem is less important because under most conditions the amounts of pyruvate or intermediates that may give rise to pyruvate in extracts is negligible.

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Changes in activity in fresh extracts are likely to be the result of changes in phosphorylation or some other covalent modification, but they may also be the result of changes in the residual tight binding of some allosteric ligand, for example, fatty acyl-CoA esters. However, changes in activity of acetyl-CoA carboxylase have been found after purification of the enzyme which strongly suggests that some covalent modification is involved. Thus the diminished activity after exposure of liver cells to glucagon or fat cells to either epinephrine or glucagon are evident after purification by a combination of ammonium sulphate fractionation and chromatography on Sepharose-avidin (50, 92, 101). In the same way, activation of the white adipose tissue enzyme after treatment of the tissue with insulin was found to persist, albeit diminished, through a 100-fold purification by ammonium sulphate fractionation and differential high-speed centrifugation (37). The effects of insulin are also still clearly demonstrable after a combination of ammonium sulphate precipitation and superose-6 chromatography (44a). However, effects of insulin appear to be lost during purification by Sepharose-avidin chromatography (50). The changes in acetyl-CoA carboxylase activity brought about by insulin are not strictly the inverse of those seen with hormones that increase cyclic AMP (Fig. 1). Whereas the effects of epinephrine and glucagon are still apparent after incubation of tissue extracts with citrate, the effects of insulin are abolished (37). Conversely, in extracts of rat epididymal adipose tissue, the effects of epi-

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a I . Persistence of changes in acetyl-CoA carboxylase activity in extracts of rat epididyrnal adipose tissue. Pieces of rat-epididymal adipose tissue were exposed to either no hormone (unshaded), insulin (shaded), or epinephrine (cross hatched) and acetyl-CoA carboxylase activity determined after incubation of tissue extracts for 20-30 min at 30°C with (a) no additions (b) citrate (20 mM) (c) MgCI2 (5 mM) plus Ca2+ (about 50 N ) .Data taken from Ref. (37). FIG.

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ROGER W. BROWNSEY AND RICHARD M. DENTON

nephrine disappear if tissue extracts are incubated with Mg2+ and Ca2+ but those incubated with insulin remain (37, 77). As discussed in Section VII, partial dephosphorylation of acetyl-CoA carboxylase occurs under these conditions (77). Taken together these observations suggest that epinephrine and insulin may act through different modes of regulation which is in accord with the idea proposed in Section V that the hormones act by altering the level of phosphorylation of different sites on the enzyme.

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

The first hint that reversible phosphorylation might play a role in the regulation of acetyl-CoA carboxylase came from the finding that the enzyme purified from rat liver contained about 2 mol of alkali-labile phosphate per mol of 21 5K subunit (36). Subsequently, Carlson and Kim (102, 103) found that incubation of ammonium sulfate fractions from rat liver with MgATP led to loss of enzyme activity whereas incubation with Mg2 resulted in activation, which could be blocked by fluoride. These findings were consistent with the activity of acetylCoA carboxylase being inhibited with increasing phosphorylation and represented the first evidence for a role for reversible phosphorylation in the regulation of this enzyme. However, it was not clearly demonstrated in this early work that phosphorylation of acetyl-CoA carboxylase was actually occurring since incorporation of 32P from [ Y - ~ ~ P I A Tinto P a protein of subunit with a M, of 230K-250K was not shown. Phosphorylation of acetyl-CoA carboxylase was then demonstrated within intact epididymal fat cells by Brownsey et al. (104). In these studies, fat cells were incubated in medium containing 32Pi and acetyl-CoA carboxylase then rapidly separated from other cell proteins by either specific immunoprecipitation or affinity chromatography on Sepharose-avidin. Both techniques resulted in the isolation of 32P-labeledprotein, M,230K, which comigrated with purified [ I4Cbiotinyll-acetyl-CoA carboxylase on SDS-gel electrophoresis (104) (Fig. 2). Similar techniques have been used in later studies to show phosphorylation of acetyl-CoA carboxylase in cultured chick liver cells (105) and rat hepatocytes (26). On the basis of chromatographic separation of apparently two forms of the enzyme it had been concluded that fatty acid synthetase was also regulated by phosphorylation (106). However, in contrast to acetyl-CoA carboxylase this conclusion appears to be incorrect since no appreciable incorporation of 32Pfrom medium [32P]phosphateinto fatty acid synthetase within intact fat cells is found under conditions where phosphorylation of acetyl-CoA carboxylase is readily observed (51, 104). +

5.

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ACETYL-COA CARBOXYLASE

S

P

S

P

FIG. 2. Immunoprecipitation of ["Placetyl-CoA carboxylase from an extract of rat epididymal fat cells. Fat cells were incubated with medium containing 32Pi for 60 min and a 100,000g supernatant prepared. This was then incubated with antiserum to acetyl-CoA carboxylase for 30 min at 30°C before centrifugation at 80,000 g to give (P) antibody precipitate and (S) supernatant fractions. Proteins in equivalent samples were then separated by SDS-polyacrylamide-gel electrophoresis, stained with Coomassie blue and radioautographed. It should be noted that the protein band of M , 230K in the initial supernatant represents both acetyl-CoA carboxylase and fatty acid synthetase since the unphosphorylated component remaining in the antibody supernatant could be precipitated following incubation with antisera to fatty acid synthase. The 32P-labeled protein band of M , 130K is ATPcitrate lyase. Further details are given in Ref. (77) and (104).

V. Effects of Hormones on the level of Phosphorylation of Acetyl-CoA Carboxylase within Intact Cell Preparations

Incorporation of 32P into acetyl-CoA carboxylase in fat or liver cells during incubation with medium containing [32P]phosphatereaches a steady-state after 45-60 min. Addition of either insulin or a hormone that increases cell cyclic AMP at this point, results in modest increases in phosphorylation within 5-15 min as judged by 32P incorporation. The increase with insulin is about 10-40% in both fat (26, 104, 107, 108) and rat liver cells (109). Exposure of rat epi-

no hot-ntone insol i n er, I neDtir ine

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FIG. 3. Two-dimensional separation of tryptic 3*P-phosphopeptides derived from acetyl-CoA carboxylase labeled within intact epididymal fat cells. Pieces of rat epididymal fat pads were preincubated with medium containing 32P, for 90 min and then for a further 15 min in the same medium with additions of (a) no hormone (b) insulin (0.1 llM) or (c) isoproterenol(5pW) as appropriate. Acetyl-CoA carboxylase was isolated by immunoprecipitation, digested with trypsin, and peptides separated on thin-layer cellulose plates by electrophoresis in the first dimension and chromatography in the second dimension. Radioautographs (a)-(c) were then prepared. [Unpublished results of R. W. Brownsey and R. M. Denton, but full experimental conditions were as given in Ref. (37)].A key is shown which designates the groups of [32P]phosphopeptides observed. The histogram summarizes the effects of insulin and epinephrine on the phosphorylation of specific sites on acetyl-CoA carboxylase in fat cells found in 10 separate experiments in an earlier study (37). In this study the D-group of peptides were not observed perhaps because they were removed from the plate during electrophoresis.

5. ACETYL-COA CARBOXYLASE

137

didymal fat cells to epinephrine (77, 101) or glucagon (101) results in a 18-60% increase in phosphorylation and a similar increase has been found in liver cells incubated with medium containing glucagon (26, 39a, 92). Initially, these findings were rather surprising as insulin results in activation of the enzyme whereas epinephrine and glucagon result in diminished activity. Because the work of Hardie and his colleagues had indicated that phosphorylation of the mammary gland enzyme occurred at multiple sites (27, 110, I l l ) we explored the extent of multisite phosphorylation of acetyl-CoA carboxylase in rat epididymal fat cells (37, 108, 112). Acetyl-CoA carboxylase was isolated by immunoprecipitation, digested with trypsin and the 32P-peptides separated in a two-dimensional system (Fig. 3). A number of different peptides were evident from the radioautographs. Incorporation of 32P into one group of 2 or 3 peptides (designated A-peptides) was increased by epinephrine about twofold but was unaffected by insulin. In contrast, insulin increased incorporation into a different peptide (I-peptide) some fivefold. Neither hormone apparently affected phosphorylation of the other peptides (C-group). Estimates of stoichiometry are difficult because the steady-state specific activity of y-32Pof intracellular ATP under these conditions is only 25-30% of the original specific activity of medium phosphate (113) and because of the quantification of the recovery of phosphopeptides. Our best estimates are that total phosphorylation of the fat cell acetyl-CoA carboxylase is 2-3 mol/mol of subunit. All the phosphorylation appears to be on serines. Exposure to insulin results in an increase of phosphorylation of the I-site by about 0.5 mol/mol of subunit while exposure to epinephrine leads to an increase in overall phosphorylation of acetyl-CoA carboxylase by about 1 mol/mol of subunit of which most occurs at the A-sites. These studies showed that exposure of fat cells to insulin and epinephrine led essentially to the phosphorylation of different sites on acetyl-CoA carboxylase and suggested that these different patterns of phosphorylation might be associated with the opposite changes in activity. Broadly similar findings have been obtained by Witters et al. in which fat cell acetyl-CoA carboxylase was isolated by ammonium sulfate fractionation plus Sepharose-avidin chromatography and tryptic peptides separated by HPLC on a C,,-reverse phase column (50). Phosphopeptides were eluted in four peaks by 31, 35, 36, and 46% acetonitrile. Epinephrine increased incorporation into a phosphopeptide or peptides eluted with 3 1% acetonitrile whereas the main effect of insulin was to increase incorporation into a peptide eluted with 35% acetonitrile. The changes in peptide phosphorylation (especially with insulin) were smaller than those found following trypsin digestion of immunoprecipitates (37). This may be because not all the different phosphorylated forms of the enzyme are recovered in equal proportions through Sepharose-avidin chromatography. The overall recovery of enzyme activity in this study was less than 25%. Holland and Hardic (114) have carried out similar studies on the effects of insulin and EGF on

138

ROGER W. BROWNSEY AND RICHARD M. DENTON

isolated liver cells from fasted-refed rats. Again acetyl-CoA carboxylase was purified by ammonium sulphate fractionation plus Sepharose-avidin chromatography and tryptic peptides separated by HPLC on a C,,-reverse phase column. The pattern of phosphopeptides eluted appear to be rather different to those observed by Witters et al. (50) since [32P]phosphopeptideswere eluted at 29, 33, 41, and 42% acetonitrile. Insulin increased incorporation into the 42% peak. Further studies are needed to relate the tryptic peptides separated by HPLC in these two studies to those separated by the thin-layer two-dimensional system used in this laboratory (37, 108, 112). In some recent studies it has become evident that insulin can give rise to increases in the phosphorylation of one or two peptides in addition to the I-peptide ( 4 4 ~ )Further . studies are required to establish whether all these peptides contain the same phosphorylation site. Studies have also been carried out by Holland, Hardie, and colleagues (92, 101) using chymotrypsin rather than trypsin to generate peptides from acetylCoA carboxylase purified by Sepharose-avidin chromatography from liver or fat cells previously incubated with medium containing [32P]phosphate. In both cases, at least seven phosphopeptides were separated from the enzymes derived from cells incubated in the absence of hormone. Exposure of liver cells to glucagon increased phosphorylation of the first phosphopeptide to be eluted from the HPLC-column without major changes in the phosphorylation of the other peptides (92). However, exposure of fat cells to either glucagon or epinephrine was found to increase the phosphorylation not only of the corresponding peptide but two or three other peptides in addition (101). In all the studies discussed (37, 50, 92, 101), the overall increase in phosphorylation observed in fat or liver cells exposed to a high concentration of hormones that increase cyclic AMP was found to correspond to 0.5-1.5 mol phosphate per mol of enzyme subunit but it is evident that the actual number of phosphorylation sites that exhibit increased phosphorylation is still a matter of dispute. Although the studies employing peptide analysis clearly indicate that acetylCoA carboxylase may be phosphorylated in cells on many serines, the exact number of different sites cannot be stated with any confidence. However, it is very unlikely that acetyl-CoA carboxylase will prove to have less than six different phosphorylation sites since acetyl-CoA carboxylase purified from liver cells may contain more than 5 mol alkali labile phosphate per mol of subunit (34, 92) and the enzyme from rabbit mammary gland more than 6 mol/mol of subunit (111). The number of phosphorylation sites on acetyl-CoA carboxylase can be confidently predicted to rival the number on glycogen synthetase!

VI.

Protein Kinases That Phosphorylate Acetyl-CoA Car boxylase

Most studies have been concerned with the phosphorylation of acetyl-CoA carboxylase by the catalytic subunit of cyclic AMP-dependent protein kinase.

5. ACETYL-COA CARBOXYLASE

139

Phosphorylation of purified acetyl-CoA carboxylase was first reported for the enzyme from rabbit mammary gland (11I). This was extended with studies of the rat mammary gland enzyme which demonstrated that phosphorylation resulted in inactivation and that the inactivation could be reversed on dephosphorylation (33). Phosphorylation resulted in a decrease in V,,, and an increase in the apparent K, for citrate (33). Similar changes in activity following phosphorylation by cyclic AMP-dependent protein kinase have also been demonstrated with acetyl-CoA carboxylase purified from rat adipose tissue (115) and from rat liver (50, 116). The changes are in line with those found after exposure of intact fat or liver cells to hormones that increase cell cyclic AMP (50, 77, 79, 92, 101). Up to 2 mol of phosphate per mol of subunit have been reported to be incorporated into acetyl-CoA carboxylase (33, 92). However, Lent and Kim (117) reported that cyclic AMP-dependent protein kinase is unable to phosphorylate acetyl-CoA carboxylase purified from rat liver directly but instead phosphorylates and activates another protein kinase which in turn phosphorylates acetyl-CoA carboxylase resulting in inactivation. This second cyclic nucleotide-independent kinase appears to require CoA (117-119). Others have found no evidence for such a bicyclic cascade (120) and it seems reasonable to conclude that acetyl-CoA carboxylase is a direct substrate for cyclic AMP-dependent protein kinase but the possibility that indirect increases in phosphorylation can also occur when cyclic AMP-dependent protein kinase is activated must be considered. A number of studies have used phosphopeptide analysis to compare the sites exhibiting increased phosphorylation in cells exposed to hormones that increase cyclic AMP such as glucagon and epinephrine to those phosphorylated in purified acetyl-CoA carboxylase treated with cyclic AMP-dependent protein kinase (50, 112, 92, 101). In general reasonable but not perfect correlations have been found. For example, in an early study the major tryptic phosphopeptides derived from mammary tissue acetyl-CoA carboxylase separated on an isoelectric focussing gel had a pl of 7.0 and comigrated with the major phosphopeptide derived from enzyme isolated by immunoprecipitation from fat cells treated with epinephrine (/12). Trypsin digestion of rat mammary or adipose tissue acetyl-CoA carboxylase phosphorylated by cyclic AMP-dependent protein kinase yields a number of phosphopeptides which can be separated in the thin-layer two-dimensional system used in Fig. 3; the principal phosphopeptides run in the same positions as the A-group of peptides which exhibit increased phosphorylation when derived from fat cells treated with epinephrine. In further studies, phosphopeptides have been separated by HPLC and similar correspondences observed (50, 92, 101). However, it is noteworthy that in one study (50) only a single major peptide peak appears to be evident after digestion with trypsin whereas in another study using similar chromatographic conditions two major peptides peaks were found ( / 2 0 ) , and at least three different major peaks have been observed after digestion with chymotrypsin (92, 101). These different profiles might be in part the result of incomplete digestions but it is also compati-

140

ROGER W. BROWNSEY AND RICHARD M. DENTON

ble with cyclic AMP-dependent protein kinase resulting in phosphorylation of three or more different sites (92, 101). Further studies, in particular the determination of amino acid sequences adjacent to the phosphorylation sites, are required to resolve the obvious complexities in this area. There are apparently at least two different cyclic nucleotide independent protein kinases that phosphorylate acetyl-CoA carboxylase resulting in a diminution of activity. One as previously mentioned appears to be phosphorylated and activated by cyclic AMP-dependent protein kinase (41, 117-1 19). This kinase initially copurifies with rat liver acetyl-CoA carboxylase through polyethylene glycol fractionation and has been further purified by DEAE cellulose and Sepharose 2B chromatography. It has been reported to have a subunit with an M, of 170K and to be markedly activated by coenzyme A. The site or sites on acetylCoA carboxylase phosphorylated by this kinase are not known. The other independent kinase capable of inhibiting acetyl-CoA carboxylase activity copurifies with acetyl-CoA carboxylase prepared from rabbit and rat mammary gland (111, 120). This enzyme appears to phosphorylate acetyl-CoA carboxylase to at least 1.5 mol/mol of subunit on the same sites as those phosphorylated by cyclic AMP-dependent protein kinase (120). Protein kinases corresponding to casein kinase I and I1 have been separated from rat mammary tissue (120) and rat liver (121) and shown to phosphorylate purified acetyl-CoA carboxylase at sites that are distinct from those phosphorylated by cyclic AMP-dependent protein kinase (50, 120). No changes in activity have been found with either kinase although incorporation of up to 1.9 mol of phosphate per mol subunit has been reported for casein kinase I and up to 0.5 mol/mol for casein kinase I1 (121). Phosphopeptide analysis of tryptic peptides by HPLC has indicated that casein kinase I1 increases the phosphorylation of a single peptide whereas casein kinase I increases the phosphorylation of the same peptide together with either one or two others (50, 120). The peptide that is phosphorylated by both kinases may be the same peptide that exhibits increased phosphorylation in fat or liver cells exposed to insulin (50, 114) but of course, the peptide may contain more than one phosphorylation site. Acetyl-CoA carboxylase does not appear to be a good substrate for protein kinase C, glycogen synthase kinase 3, or phosphorylase kinase (46). There remains the important question of the identity of the kinase activity responsible for the increased phosphorylation of acetyl-CoA carboxylase in fat and liver cells exposed to insulin. Purified plasma membranes from fat cells have been found to display protein kinase activity that was capable of phosphorylating partially purified acetyl-CoA carboxylase with apparent concomittant activation of the enzyme (115). The kinase was found to employ MgATP, to be insensitive to cyclic nucleotides and the specific (Walsh) inhibitor protein, and was apparently fully active in the absence of Ca2+ (115). The changes in catalytic activity of acetyl-CoA carboxylase observed following incubation with plasma mem-

141

5. ACETYL-COA CARBOXYLASE

brane preparations were similar to those that are seen following the exposure of intact fat cells to insulin; in particular, increases were on initial activity and were no longer evident after treatment of the enzyme with citrate (115). Under those conditions phosphorylation appeared to occur at a number of sites on the enzyme but about 50% occurred at I-site as defined in Fig. 3. No appreciable change in acetyl-CoA carboxylase kinase activity was found on adding insulin direct to the fat cell membrane preparation (21). Subsequently it was observed that exposure of intact fat cell preparations to insulin resulted in an increase in protein kinase activity in subsequently prepared high-speed supernatant fractions (122). The properties of this protein kinase appeared to be similar to the cyclic AMPindependent activity associated with fat cell plasma membranes. Activation persisted during dialysis and gel filtration (122). These findings are compatible with the idea that binding of insulin to its plasma membrane receptor leads to the activation of a protein-serine kinase capable of phosphorylating acetyl-CoA carboxylase and perhaps other proteins which also exhibit increased phosphorylation in insulin-treated cells (123). We have found (123) that triton extracts of placenta membranes that are rich in insulin receptors also contain protein-serine kinase activity apparently capable of phosphorylating acetyl-CoA carboxylase on the I-peptide (as defined in Fig. 3). Under certain conditions the addition of insulin leads not only to increased phosphorylation of the insulin receptors on tyrosine residues but also to increased serine phosphorylation of acetyl-CoA carboxylase (124).

VII.

Protein Phosphatases That Act on Acetyl-CoA Carboxylase

Characterization of the protein phosphatases able to dephosphorylate acetylCoA carboxylase is in its early stages and no clear picture has emerged of the number of different protein phosphatases that may be involved in the regulation of acetyl-CoA carboxylase within intact cells. Major problems include the difficulty of obtaining acetyl-CoA carboxylase phosphorylated only on defined sites and the need to eliminate the possibility of limited proteolysis causing activation and/or loss of phosphorylation sites. A number of phosphatase preparations have been found to be capable of dephosphorylating the sites on acetyl-CoA carboxylase phosphorylated by cyclic AMP-dependent protein kinase and hence activating the enzyme. These include a number of phosphatases purified from rat liver [designated I ,2A, 2A2, and 2C by the Dundee group (33, 38, 125, 126)] and similar results have been obtained by others (127, 128). The liver phosphatase activities corresponding to 2A, and 2A, have been reported to be markedly activated by Mg2 or Mn2 (128, 128a) and to also dephosphorylate the sites phosphorylated by casein kinase I and I1 +

+

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ROGER W . BROWNSEY AND RICHARD M . DENTON

(128). The major phosphatase activity capable of dephosphorylating acetyl-CoA carboxylase in extracts of adipose and mammary tissue are also activated by Mg2+ and may be further stimulated by Ca2+ (51, 77, 129). But these tissues appear to also contain other phosphatases which copurify with acetyl-CoA carboxylase (130, 131).

VIII. Concluding Remarks The short-term regulation of acetyl-CoA carboxylase activity is an important means whereby the rate of fatty acid synthesis is controlled. Many questions concerning the relationships between phosphorylation of the various sites, the state of polymerization of the enzyme, and catalytic activity remain to be answered. In the cell, flux through acetyl-CoA carboxylase is governed by a combination of changes in substrate concentration and alterations in the levels of possible effectors (including citrate, fatty acyl-CoA esters, phosphoinositide, and CoA) as well as the modifications in phosphorylation of at least six different sites on the enzyme. Determining the relative importance of these various means of control will undoubtedly tax researchers for many years to come. Even finding appropriate in vitro assay conditions that can faithfully reflect the activity of the enzyme within intact cells is a formidable problem. The inhibitory effects of hormones that increase the cell concentration of cyclic AMP on the activity of acetyl-CoA carboxylase appear to be essentially preserved through purification to homogeneity suggesting that changes in phosphorylation by cyclic AMP-dependent protein kinase may be sufficient to explain the changes in enzyme activity. However, the role of the other acetyl-CoA carboxylase kinases that inactivate the enzyme need to be clarified. It is generally agreed that acetyl-CoA carboxylase is activated in tissues exposed to insulin and at least in liver and white adipose tissue, the activation appears to be associated with increased phosphorylation of the enzyme. Epidermal growth factor may activate acetyl-CoA carboxylase by a similar mechanism. However, in contrast to effects of the hormones acting through cyclic AMP, the effects of insulin are lost during purification of the enzyme by sepharose avidin chromatography. One possible explanation is that phosphorylation alters the response of the enzyme to a tightly bound effector (perhaps fatty acyl-CoA or a protein) which is progressively lost during purification; another is that the recovery of the activated phosphorylated form of the enzvme is less than the recovery of other less active phosphorylated forms (see Sections II,A and V). The identity of the protein kinase responsible for the increased phosphorylation is clearly of critical importance. The recognition of the means whereby insulin brings about the stimulation of protein-serine kinase activity is likely to represent an important step in the understanding of the molecular basis of insulin action since acetyl-

5.

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ACETYL-COA CARBOXYLASE

CoA carboxylase is only one of a number of proteins exhibiting increased phosphorylation on serines following exposure of cells to insulin. Others include ATP-citrate lyase, the ribosomal protein S6 and further proteins of unknown function (123, 132). In 1981 it was suggested that the binding of insulin to its plasma membrane receptors may lead to an enhancement of protein kinase activity perhaps by causing the dissociation of protein kinase activity from the inner face of the plasma membrane (123). It is becoming increasingly evident that a characteristic of insulin action on cells may be the rapid translocation of components between plasma membrane and intracellular locations; examples include both the recruitment of glucose transporters from intracellular sites and the rapid internalization of insulin-receptor complexes (133, 134). An attractive possibility is that the stimulation of protein-serine kinase activity in cells is initiated through the phosphorylation of tyrosine residues of a membrane-associated protein by the insulin receptor kinase (21, 135). The close association of the insulin receptor and a plasma membrane protein-serine kinase is indicated both by the finding that increased phosphorylation of the P-subunit of the receptor in cells exposed to insulin occurs on serine as well as tyrosine residues (136, 137) and that increased serine phosphorylation of acetyl-CoA carboxylase ( 1 2 4 , histone (137), casein (137), and actin (138) can be observed on addition of insulin to triton solubilized insulin receptor preparations.

ACKNOWLEDGMENT Support for the work in the authors’ laboratories has come from the M.R.C.s of both the United Kingdom and Canada, the British Diabetic Association, the Juvenile Diabetes Foundation, the Percival Waite Salmond Bequest, and the British Columbia Health Care Research Foundation.

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59. Brownsey, R. W., Bridges, B., and Denton, R. M. (1977). Biochem. SOC. Trans. 5, 1286. 60. Hams, R. A. (1975). ABB 169, 168. 61. Watkins, P. A,, Tarlow, D. M., and Lane, M. D. (1977). PNAS 74, 1497. 62. Clarke, S. D . , Watkins, P. A., and Lane, M. D. (1979). J. Lipid Res. 20, 974. 63. Lane, M. D., Watkins, P. A., and Meredith, M. J. (1979). CRC Crit. Rev. Biochem. 7, 121. 64. Siess, E. A , , Brocks, D. G.,Lattke, H. K., and Wieland, 0. H. (1977). BJ 166, 225. 65. Wieland, 0. H. (1983). Rev. Physiol. Biochem. Pharmacol. 96, 123. 66. Yeh, L.-A., and Kim, K.-H. (1980). PNAS 77, 3351. 67. Yeh, L.-A,, Lee, K.-H., and Kim, K.-H. (1980). JBC 255, 2308. 68. Witters, L. A,, Friedman, J. A,, Tipper, J. P., and Bacon, G. W. (1981). JBC 256, 8573. 69. Buechler, K. F., and Gibson, D. M. (1984). ABB 233, 698. 70. Heger, H. W., and Peter, H. W. (1977). Int. J . Biochem. 8, 841. 71. Blytt, H. J . , and Kim, K.-H. (1982). ABB 213, 523. 72. Windmueller, H. G.,and Spaeth, A. E. (1966). JBC 241, 2891. 73. Windmueller, H. G.,and Spaeth, A. E. (1967). ABB 122, 362. 74. Stansbie, D., Brownsey, R. W., Crettaz, M., and Denton, R. M. (1976). BJ 160, 413. 75. Lee, K.-H., and Kim, K.-H. (1979). JBC 254, 1450. 76. Halestrap, A. P., and Denton, R. M. (1973). BJ 132, 509. 77. Brownsey, R. W., Hughes, W. A,, and Denton, R. M. (1979). BJ 184, 23. 78. Lee, K.-H., Thrall, T., and Kim, K.-H. (1973). BBRC 54, 1133. 79. Zammit, V. A,, and Corstorphine, C. G. (1982). BJ 208, 783. 79a. Haystead, T. A. J., and Hardie, D. G . (1986). BJ 234, 279. 80. McCormack, J. G.,and Denton, R. M. (1977). BJ 166, 627. 81. Agius, L., and Williamson, D. H. (1980). BJ 192, 361. 82. Gibbins, I. M., Denton, R. M., and McCormack, J . G . (1985). BJ 228, 751. 83. McNeillie, E. M., and Zammit, V. A. (1982). BJ 204, 273. 84. Munday, M. R., and Williamson, D. H. (1982). FEBS Lett. 138, 285. 85. Robson, N. A,, Clegg, R. A,, and Zammit, V. A. (1984). BJ 217, 743. 86. Williamson, D. H., Munday, M. R., Jones, R. G.,Roberts, A. F. C., and Ramsey, A. J . (1983). Adv. Enzyme Regul. 21, 135. 87. Klein, G.J . , and Weiser, P. C. (1973). BBRC 55, 76. 88. Witters, L. A , , Moriarity, D., and Martin, D. B. (1979). JBC 254, 6644. 89. Assimacopoulos-Jeannet, F. D., Denton, R. M., and Jeanrenaud, B. (1981). BJ 198, 485. 90. Geelen, M. J. H., Beynen, A. C . , Christiansen, R. Z., Lepreau-Jose, M. J . , and Gibson, D. M. (1978). FEBS Lett. 95, 326. 91. Beynen, A. C., Vaartjes, W. J., and Geelen, M. J . H. (1979). Diabetes 28, 828. 92. Holland, R., Witters, L. A , , and Hardie, D. G. (1984). EJB 140, 325. 93. Ly, S., and Kim, K.-H. (1981). JBC 256, 11585. 94. Flatt, J. P., and Ball, E. G.(1964). JBC 239, 675. 95. Jungas, R. L. (1970). Endocrinology (Baltimore) 86, 1368. 96. Munday, M. R., and Williamson, D. H. (1982). FEBS Lett. 138, 285. 97. Allred, J. B., and Roehrig, K. L. (1973). JBC 248, 4131. 98. Ma, G.Y., and Hems, D. A. (1975). BJ 152, 389. 99. Davies, D. R., Van Schaftingen, E., and Hers, H.-G. (1982). BJ 202, 559. 100. Allred, J. B., and Goodson, J . (1982). BJ 208, 247. 101. Holland, R., Hardie, D. G.,Clegg, R. A,, and Zammit, V. A. (1985). BJ 226, 139. 102. Carlson, C. A,, and Kim, K.-H. (1973). JBC 248, 378. 103. Carlson, C. A,, and Kim, K.-H. (1974). ABB 164, 478. 104. Brownsey, R. W., Hughes, W. A., Denton, R. M., and Mayer, R. J . (1977). BJ 168, 441. 105. Pekala, P. H., Meredith, M. J . , Tarlow, D. M., and Lane, M. D. (1978). JBC 253, 5267.

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106. Qureshi, A. A., Jenik, R. A., Kim, M., Lornitzo, F. A., and Porter, J. W. (1975). BBRC 66, 344. 107. Brownsey, R. W., Belsham, G. J., and Denton, R. M. (1981). Biochem. SOC. Trans. 9, 232. 108. Brownsey, R. W. (1981). Biochem. SOC. Trans. 9, 515. 109. Witters, L. A. (1981). BBRC 100, 872. 110. Hardie, D. G. (1980). In “Recently Discovered Systems of Enzyme Regulation by Reversible Phosphorylation” (P. Cohen, ed.), p. 33. Elsevier/North-Holland Biomedical Press, Amsterdam. 11 1. Hardie, D. G., and Cohen, P. (1978). FEBS Lett. 91, 1. 112. Brownsey, R. W., and Hardie, D. G. (1980). FEBS Lett. 120, 67. 113. Hopkirk, T. J., and Denton, R. M. (1985). BBA 885, 195. 114. Holland, R., and Hardie, D. G. (1985). FEBS Lett. 181, 308. 115. Brownsey, R. W., Belsham, G. J., and Denton, R. M. (1981). FEBS Lett. 124, 145. 116. Tipper, J. P., and Witters, L. A. (1982). BBA 715, 162. 117. Lent, B. A,, and Kim, K.-H. (1983). ABB 225, 972. 118. Lent, B. A., and Kim, K.-H. (1982). JBC 257, 1897. 119. Lent, B. A., and Kim, K.-H. (1983). ABB 225, 964. 120. Munday, M. R., and Hardie, D. G. (1984). EJB 141, 617. 121. Tipper, J. P., Bacon, G. W., and Witters, L. A. (1983). ABB 227, 386. 122. Brownsey, R. W., Edgell, N. J., Hopkirk, T. J., and Denton, R. M. (1984). BJ 218, 733. 123. Denton, R. M., Brownsey, R. W., and Belsham, G. J. (1981). Diabetologia 21, 347. 124. Tavare, J. M., Smyth, I. E., Borthwick, A. C., Brownsey, R. W., and Denton, R. M. (1985). Biochem. SOC. Trans. 13, 734. 125. Ingebritsen, T. S., and Cohen, P. (1983). EJB 132, 255. 126. Ingebritsen, T. S., Blair, I.. Guy,P., Witters, L. A,, and Hardie, D. G. (1983). EJB 132,275. 127. Krakower, G . R., and Kim, K.-H. (1980). BBRC 92, 389. 128. Witters, L. A., and Bacon, G. W. (1985). BBRC 130, 1132. 128a. Thampy, K. G., and Wakil, S. I. (1985). JBC 260, 6318. 129. McNeillie, E. M., Clegg, R. A,, and Zammit, V. A. (1981). BJ 200, 639. 130. Krakower, G. R., and Kim, K.-H. (1981). JBC 256, 2408. 131. Hardie, D. G., and Cohen, P. (1979). FEBS Lett. 103, 333. 132. Avruch, J., Alexander, M. C., Palmer, J. L., Pierce, M. W., Nemenoff, R. A,, Tipper, J. P., and Witters, L. A. (1982). FP 41, 2629. 133. Cushman, S. W., Wardazala, L. J., Simpson, I. A,, Karnieli, E., Hissin, J. H., Wheeler, T. J., Hinkle, P., and Salans, L. B. (1983). Horm. Cell Regul. 7, 73. 134. Sonne, O., and Simpson, I. (1984). BBA 804, 404. 135. Brownsey, R. W., and Denton, R. M. (1985). In “Molecular Basis of Insulin Action” (M. P. Czech, ed.), p. 297. Plenum, New York. 136. Kasuga, M., Zick, Y.,Blith, D. L., Carlson, F. A., Haring, H. U., and Kahn, C. R. (1982). JBC 257, 9891. 137. Gazzano, H., Kowalski, A,, Fehlmann, M., and Van Obberghen, E. (1984). BJ 216, 575. 138. Machicao, F., Caracosa, J. M., and Wieland, 0. H. (1984). Diabetologia 27, 305A.

Hormone-Sensitive Lipase PETER

STRALFORS HAKAN OLSSON

PER BELFRAGE

Department of Physiological Chemistry University of Lund S-221 00 Lund, Sweden

I. Introduction . . . . . . . . . . . . . . . . . . 11. Properties .................... 111. Mechanism of Regulation of the Adipose Tissue Lipase . . . . . . . . . . . . . . . . A. Activation-Deactivation of the Isolated Enzyme by Reversible Phosphorylation .................................. B. Short-Term Hormonal Control in the Intact Fat Cell . . . . . . . . . . . . . . . . C. Mechanisms of Action for the Short-Term Control by Lipolytic Hormones and Insulin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Other Mechanisms for Regulation of Adipose Tissue Lipolysis . . . . . . . IV. Possible Role Activatable, Multifunctional

....................................... V. Conclusions and Perspectives .................. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1.

147 148 152 152 156 161 166 168

171 172

introduction

Hormone-sensitive lipase (HSL) has a major role in fatty acid mobilization from adipose tissue by catalyzing the rate-limiting step in lipolysis of stored triacylglycerols. The mobilization of fatty acids is of fundamental biological importance since they constitute the major energy fuel in mammals. The rate of this process is under tight hormonal and neural control, to a large extent exercised through regulation of the activity of HSL. 147 THE ENZYMES. Vol XVIll Copyright 0 1987 by Academic Press. Inc All rights of reproduction In any form reserved

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Originally believed to be an enzyme unique to the adipocyte, HSL now seems likely to have a wider occurrence. It (or a very similar enzyme) has been identified in steroid-producing tissues and regulation of the enzyme activity implicated as part of the hormonal control of steroidogenesis. A role for the enzyme as a hormonally regulated tissue acylglycerol lipase-cholesterol ester hydrolase controlling the rate of several important biological processes is emerging. From previous work with adipose tissue preparations fast-acting lipolytic hormones were generally thought to control HSL through cyclic AMP-mediated phosphorylation of the enzyme. The direct experimental proof for this notion was obtained following the identification of the enzyme protein (1). This chapter deals with the molecular mechanisms for short-term hormonal control of the enzyme in adipose tissue and other organs and emphasizes the current state of knowledge and the progress as the result of identification, purification, and characterization ( I , 2) of HSL. Detailed reviews of the historical background, purification, and properties of HSL have appeared elsewhere (3-5);these aspects are only briefly mentioned here. A more general view of the hormonal, neural, and pharmacological control of adipose tissue lipolysis and fatty acid mobilization can be found in other reviews (6-8).

II. Properties The first, rate-limiting, and second hydrolysis steps in the lipolytic degradation of stored adipocyte triacylglycerols to fatty acids and glycerol are catalyzed by HSL (Fig. 1). Most of the monoacylglycerols produced are hydrolyzed by a specific, hormone-insensitive monoacylglycerol lipase (9) as the result of the substrate and positional specificities of the two enzymes. HSL was detected as a hormonally regulated enzyme activity more than 20

lips/

hormone-sensitive lipase\

~

rnonoacylglycerol

adipocyte triacylglycerol

1.2-d ia c y Ig Iy c er o I

T FFA

T-

2-monoac y lg l y cero I

FFA

FIG. 1 . Main reaction sequence of adipose tissue lipolysis.

T FFA

glycerol

149

6. HORMONE-SENSITIVE LIPASE TABLE I PROPERTIES OF HORMONE-SENSITIVE LIPASEo h RAT ADIPOSETISSUE" Property Subunit M, (SDS-PAGE)" Apparent M, of detergent-solubilized enzyme< Relative sedimentation coefficient Isoelectric point (f4" C) Binding of nonionic detergent" Inhibitors'

Comment 84,000 160,000 4.5 s 6.7-6.8 Yes DFP,9 pM

HgCI2.11 W Substrate specificity

Positional specificity Specific activity Molar specific activity/ pH optimum

NaF,25 mM Long-chain tri-, I ,2-di-, and 2-monoacylglycerol, cholesterol esters (re1 v,,,, I : 10: I :I 3 I (3)-ester fourfold faster than 2-ester bonds 400 pmol fatty acid/min/mg 600 mol/mol enzyme/s 7.0

"Data are from Refs. ( I , 2 , 16, 21, 25). bUnder reducing or nonreducing conditions. ('Gel chromatography data. Probably represents an enzyme dimer, with some bound nonionic detergent. dFrom data obtained by charge-shift electrophoresis and phase separation in Triton X-I14 ( 2 5 ) . Concentration giving 50% inhibition with trioleoylglycerol as substrate at 37" C . /Emulsified monooleoyloleylglycerol (diacylglycerol analog) as substrate at 37" C. Activity related to amount of M,=84,000 subunit.

years ago (10-14) [the term HSL was coined in 1964 by Vaughan et al. ( l o ) ] it; turned out to be extremely recalcitrant to purification because of its low abundance, hydrophobic character, and general lability. Until a few years ago only partial purification of the enzyme in a particulate, lipid-associated form was possible [reviewed in Refs. (3, 4 ) ] .A procedure for detergent solubilization was developed and, after partial purification, the enzyme protein was identified (1). A protocol for preparation of 50% pure, detergent-solubilized enzyme from adipose tissue of 200 rats has been described (2, 1 3 , allowing several of its properties to be established (Table I) (16-20). Most of the purified HSL is in the dephospho form (16),probably due to rapid dephosphorylation during the first steps of the purification, when the enzyme preparations contain protein phosphatase activity (2). The purification procedure has been scaled up to make it possible to prepare lipase from adipose tissue of 500 rats at a time, and modified by substituting some of the chromatographic steps by ion-exchange chromatography using high-performance liquid chromatography (HPLC) equipment (18), so

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that 200 pg highly purified enzyme protein is obtained. More recently, procedures for preparing milligram quantities of highly purified lipase protein from bovine adipose tissue have been developed (19, 20). Neither the native structure of HSL in the intact fat cells nor its subcellular localization is known. In detergent-solubilized form it is likely that the enzyme is a dimer of noncovalently bound, identical M, = 84,000 subunits ( 2 , 21), similar to that found for lipoprotein lipase (22). In many ways HSL behaves like an intrinsic membrane protein (e.g., it associates strongly with phospholipids and requires detergents for solubilization and stabilization) ( 1 , 2 ) . Using the techniques of charge-shift electrophoresis (23) and phase separation in Triton X-114 (24) the enzyme was shown to directly bind nonionic detergent (25), although probably only a small amount since binding of nonionic detergents with widely varying aggregation number and micellar size did not markedly change the apparent Stokes radius of the enzyme-detergent complex, as measured by gelfiltration chromatography (25). In crude adipose tissue extracts HSL resides in particles or lipid-protein aggregates found in high-speed supernatants. These aggregates have been enriched 100-fold and reported to be of fairly uniform size (20-60 nm diameter, apparent M, = 7 X lo6, and density 1.08-1.09 g/ml) and to contain 50% phospholipid and some cholesterol (26, 27). However, it is not known if these structures represent the native state of the enzyme; they may have been formed during and after the homogenization of the tissue, since the isolation procedure (26) would tend to select particles of uniform size and density. A study with 3T3-Ll adipocyte-like cells indicates that HSL may be localized both in the cytosol and with membranes; the authors propose that hormonal stimulation of the cells causes redistribution of the enzyme from the high-speed supernatant to the membrane fraction (28) (further discussed in Section 111,A). However, in most work on its subcellular localization the enzyme has been quantitated by its activity; this may be erroneous when added artificial lipid emulsions are variably diluted by endogenous substrate. Especially, the considerable amount of enzyme associated with membranes residing in the floating fat layer after centrifugation is difficult to quantitate this way. Antibodies toward the lipase are now available (19, 29) making it possible to quantitate the enzyme protein in cell extracts and to directly study its subcellular localization in the adipocyte. HSL is inhibited by micromolar concentrations of diisopropyl fluorophosphate (DFP) and the inhibition is parallelled by [3H]DFP incorporation into the enzyme, indicating that a reactive serine residue is involved in its catalytic function (Table I). The presence of one or several functional sulfhydryl groups is indicated by inhibition with cysteine-directed reagents. HSL has a broad substrate specificity, hydrolyzing long-chain tri-, di-, and monoacylglycerols as well as cholesterol esters (Table I). Triacylglycerols are hydrolyzed at a much lower rate than diacylglycerol, so the first step of the

6. HORMONE-SENSITIVE LIPASE

151

lipolytic degradation sequence becomes rate limiting (Fig. l), at least in the fat cells which also contain a monoacylglycerol lipase. In contrast to HSL, this enzyme, purified to homogeneity and characterized in this laboratory ( 9 ) , lacks positional specificity, Monoacylglycerol lipase is required to hydrolyze the 2monoacylglycerols formed from the second HSL-catalyzed step; in its absence these reaction products accumulate (21). HSL has also been detergent-solubilized and purified (to about 10% protein purity) from swine adipose tissue (30).The purified swine lipase was found to be almost identical to the rat enzyme in a number of aspects [M,= 84,000 by sodium dodecyl sulfate-polyacrylmid gel electrophoresis (SDS-PAGE), the same molar-specific activity, substrate specificity, and inhibition properties]. The bovine adipose tissue HSL (19, 20) also had very similar properties except for the specific activity which was fivefold lower. This may, however, be due to inactivation during preparation rather than to a species difference. Also, enzyme from mouse 3T3-Ll adipocyte-like cells have been shown to be very similar (28, 31). In contrast, detergent-solubilized and partially purified lipase from chicken adipose tissue has been reported to be markedly different (32). Thus, the specific activity of the latter enzyme preparation was several thousandfold lower than that of HSL from rat adipose tissue (Table I) and of other tissue lipases. It seems most improbable that this should be due to a species difference. The phosphorylatable M, = 42,000 protein (SDS-PAGE), claimed to represent the HSL (32), is more likely to have been a major protein contaminant, with HSL as a small, undetected component on the SDS-PAGE gels. In several aspects (e.g., the high molar specific activity and the positional specificity) HSL is similar to other tissue lipases, such as lipoprotein lipase (22) and hepatic lipase (33). However, the relatively higher diacylglycerol lipase activity and, in particular, the ability to hydrolyze cholesterol esters distinguish HSL from these lipases. In fact, the cholesterol ester hydrolase activity of HSL is higher than the triacylglycerol hydrolase activity and could account for almost all hydrolysis of cholesterol ester in adipose tissue extracts ( 2 ) ,but there seems to be no function for this high activity in the adipose tissue. These observations were explained when it was found that HSL, or a very similar enzyme (M,= 84,000 by SDS-PAGE, the same molar specific activity, substrate and positional specificity, inhibition properties, activatability by cyclic AMP-dependent protein kinase-catalyzed phosphorylation, and similar [3H]DFP-peptidepattern after proteolytic digestion), was present in bovine adrenal cortex (34). An enzyme with several of these properties has also been found in bovine corpus luteum (35).The enzyme accounted for the well-known cytosolic, neutral cholesterol ester hydrolase activity in these tissues (34-36) [for review, see Ref. ( 5 ) ] .The function and regulation of HSL in the steroid-producing tissues and the possible existence of the enzyme also in other tissues is discussed in Section IV.

152

111.

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STRALFORS,H . OLSSON, AND P.

BELFRAGE

Mechanism of Regulation of the Adipose Tissue Lipase

A. ACTIVATION-DEACTIVATION OF THE ISOLATED ENZYMEBY REVERSIBLE PHOSPHORYLATION Early work indicated that HSL was subject to regulation (11, 12) involving cyclic AMP (14, 37). It was demonstrated that, in cell-free extracts, cyclic AMP in the presence of ATP and Mg2+ caused the enzyme activity to increase (38, 39). The specific inhibitor of cyclic AMP-dependent protein kinase blocked this activation, which was regained by addition of excess protein kinase (40). The direct involvement of cyclic AMP-dependent protein kinase was confirmed with partially purified preparations of HSL (41, 42). These findings led to the formulation of the “lipolytic activation cascade” hypothesis (43). A reversal of the cyclic AMP-dependent activation was demonstrated with different protein phosphatase preparations (44-46).

Ftc. 2. Phosphopeptide mapping of HSL, 32P-phosphorylatedby cyclic AMP-dependent protein kinase and digested with Stuphylcoccus uureus V8 proteinase and trypsin. The proteolytic digest was separated by two-dimensional electrophoresis thin-layer chromatography and autoradiographed; “x’ ’ indicates the point of application. From Ref. (16).

153

6 . HORMONE-SENSITIVE LIPASE 4001

-400

E

-?

8 - 3 0 0 r: P

0

g

m

2001

6

-200

-

2-

-

0

r

-100

0

5x

Lt

W

00 -

1

2 3 1,me mln”leS

4

5

0

15 30 Time. minutes

45

FIG. 3. Time courses for the reversible phosphorylation-activation of HSL by cyclic AMPdependent protein kinase and an adipose tissue protein phosphatase. A. HSL was incubated with cyclic AMP-dependent protein kinase and [y-’ZP]ATP for the times indicated [from Ref. (26)].B. The enzyme was maximally phosphorylated by preincubation with cyclic AMP-dependent protein kinase for 30 min, the MgATP was then removed and protein phosphatase added at time zero. After dephosphorylation, MgATP and an excess of cyclic AMP-dependent protein kinase was added again as indicated by the arrow [from Ref. (47)].At the indicated time-points aliquots were withdrawn for determination of [32P]phosphate incorporation and enzyme activation: 0-0, phosphate incorporation; -0, activation of enzyme; vertical bars indicate SE.

Following the identification of the HSL protein the regulation of the enzyme through its reversible phosphorylation was established, using highly purified preparations of the enzyme ( I , 2, 16,47). Cyclic AMP-dependent protein kinase phosphorylates HSL at a single serine residue, in a site termed the “regulatory phosphorylation site,” which can be found in a proteolytic peptide of about ten amino acid residues (Fig. 2) (16). At maximal phosphorylation close to 1 mol of phosphate is incorporated per mol of lipase M, 84,000 subunit (16). The site of phosphorylation seems to be distinct from the area in contact with the substrate interface since preincubation and phosphorylation in the presence of a triacylglycerol substrate emulsion does not affect the rate of phosphorylation by cyclic AMP-dependent protein kinase (16);also, the fatty acids produced by the lipolytic reaction do not affect the phosphorylation rate (16). As described in Section 11, HSL is covalently modified and inhibited by DFP and by sulfhydryl reagents such as N-ethylmaleimide. Treatment with any of these agents has no effect on subsequent phosphorylation by cyclic AMP-dependent protein kinase (16). The specific inhibitor protein of cyclic AMP-dependent protein kinase, which was shown to block the protein kinase-catalyzed activation of HSL (40, 48), immediately also interrupts the phosphorylation of the lipase ( 2 ) , thus conclusively demonstrating that cyclic AMP-dependent protein kinase acts directly on HSL without any intervening “lipase kinase.”

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P. STRALFORS, H. OLSSON, AND P. BELFRAGE

The phosphorylation of the regulatory site by cyclic AMP-dependent protein kinase is closely paralleled by an increased activity of the lipase against an emulsified triacylglycerol substrate ( 2 , 16) (Fig. 3A). The rate of phosphorylation is comparable to that obtained in vivo in response to hormonal stimulation of cyclic AMP production (49) and to the rate of cyclic AMP-dependent protein kinase-catalyzed phosphorylation of phosphorylase kinase (16), one of the most rapidly phosphorylated physiological substrates of the protein kinase. A reversal of cyclic AMP-dependent protein kinase-catalyzed phosphorylation and activation has been demonstrated using a protein phosphatase isolated from adipose tissue (Fig. 3B) (47). The dephosphorylation of the lipase by the four different types of protein phosphatases that have been isolated from rabbit skeletal muscle (protein phosphatase-1, -2A, -2B, and -2C) (50) have been examined. Protein phosphatases-2A and -2C were severalfold more active against the lipase than was protein phosphatase-1 (the activities were related to that against glycogen phosphorylase and phosphorylase kinase), while phosphatase-2B was without activity against the lipase. From the total activity of the different protein phosphatases in the adipose tissue (51), phosphatase-2A and -1 are likely to be the predominant lipase phosphatase activities in the fat cell. In addition to cyclic AMP-dependent protein kinase HSL is phosphorylated by other protein kinases. Cyclic GMP-dependent protein kinase rapidly phosphorylates and activates the lipase (52).The phosphorylation occurs on serine residues in two distinct phosphorylation sites, both being phosphorylated at about the same rate (Fig. 4). One phosphorylation site is the same as that phosphorylated by cyclic AMP-dependent protein kinase (the regulatory phosphorylation site); this phosphorylation accounts for the increased enzyme activity. Both this and the other phosphorylation site (termed the basal phosphorylation site) are phosphorylated in vivo (see Section II1,B). The cyclic nucleotide-independent protein kinase glycogen synthase kinase 4 (53),isolated from skeletal muscle, also phosphorylates the lipase, but without affecting the enzyme activity. The phosphorylation occurs exclusively on the basal phosphorylation site (54). Regardless of any physiological importance of cyclic GMP-dependent protein kinase and glycogen synthase kinase 4 in the control of HSL these protein kinases should prove useful in the study of regulation of lipase and, especially, of the function of the basal-site phosphorylation of the enzyme in vivo. In addition, phosphorylase kinase (isolated from skeletal muscle) phosphorylates a serine residue in a third distinct phosphorylation site that has not been observed to be phosphorylated in intact adipocytes (54). Phosphorylation of the isolated rat adipose tissue HSL with cyclic AMPdependent protein kinase shifts the p1 from around 6.7-6.8 for the dephosphoform ( 2 ) to 6.5 (16).The phosphorylation results in an up to 3-fold increase in the lipase activity against triacylglycerol (47), whereas the activity against di- and monoacylglycerolis affected very little. Thus, the relative specific activity against

6. HORMONE-SENSITIVE LIPASE

155

FIG.4. Phosphopeptide mapping of HSL, 32P-phosphorylatedby cyclic GMP-dependent protein kinase and digested with S. aureus VS, proteinase, and trypsin. The proteolytic digest was separated by two-dimensional electrophoresis thin-layer chromatography and autoradiographed; ‘‘x” indicates the point of application. For explanation of “basal” and “regulatory,” see text [from Ref. ( 5 2 ) ] .

the different acylglycerol substrates is changed from 1 :10:4 to around 1 :3:1 against tri-, di-, and monoacylglycerol, respectively (cf., Table I). With the enzyme from hen adipose tissue 10-fold activation is obtained against the tri- and diacylglycerol substrate as well as cholesterol ester (44, 55, 56). Little is known about the way changes in the phosphorylation of the enzyme are expressed as changes in enzyme activity. This is, at least partly, due to the lack of a well-defined assay system. The “interface quality” of the emulsified lipid substrate (i.e., the lipid-water interface composition) and physicochemical properties are not known and can not be experimentally controlled. It is likely that variation of these properties of the substrate accounts for the varying extent of activation that has been obtained with different acylglycerol substrates and at different ionic strengths (42, 44, 56). The general finding is that a high degree of activation is associated with a low basal enzyme activity and, hence, that at a given extent of phosphorylation the magnitude of activation largely reflects the measured basal activity of the enzyme (cf., section 111,D). The only system available where the properties and composition of the lipidwater interface are known and can be controlled is the surface monolayer technique that has been extensively used in the study of pancreatic lipase (57) and lipoprotein lipase (58). The activation mechanisms described for these two lipases could serve as a model to understand the activation of HSL. The apparent

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P. STRALFORS, H . OLSSON, AND P. BELFRAGE

affinity of HSL for the substrate interface or for the substrate molecules at the interface seems to be affected by the phosphorylation state of the enzyme (44, 59). The substrate interface is likely to be covered with amphiphilic lipids and proteins and phosphorylation of the lipase could cause changes in the orientation of the enzyme in this interfacial layer to enhance the catalytic efficiency, in analogy with apolipoprotein C,, activation of lipoprotein lipase (22)and colipase activation of pancreatic lipase (60, 61). In the 3T3-Ll adipocyte-like cells isoproterenol stimulation of lipolysis was proposed to be associated with a redistribution of HSL from a soluble to a particulate fraction through an increased interaction of the enzyme with its substrate and associated membranes (28).Details regarding the organization of the lipid droplet-cytosol interface or of the lipase in the intact cell are lacking, but this proposal seems to be in accordance with phosphorylation inducing an increased affinity of the enzyme for the substrate. It is tempting to speculate that an activator function similar to that of apolipoprotein C,, or colipase resides in a domain in the larger HSL molecule [M,= 84,000 as compared with 56,000 (62) and 52,000 (61)for lipoprotein lipase and pancreatic lipase, respectively]. Triggering of the activator function by phosphorylation of HSL could then occur through a conformational change. The possibility that HSL, lipoprotein lipase, and pancreatic lipase share a common basic mechanism for activation could serve as a useful hypothesis for further work on the mechanism by which HSL is activated through phosphorylation. HORMONAL CONTROL IN THE INTACT FAT CELL B. SHORT-TERM Much early work demonstrated involvement of cyclic AMP and cyclic AMPdependent protein kinase in the short-term hormonal activation of HSL and adipose tissue lipolysis [reviewed in Refs. (3, 5 ) ] .In accordance with the general cyclic AMP-second-messenger mechanism the lipolytic activation cascade hypothesis (43) stated that a hormone-induced increase of cellular cyclic AMP activated cyclic AMP-dependent protein kinase, which in turn phosphorylated the lipase and thereby activated it. The phosphorylation and activation was proposed to be reversed by a protein phosphatase. However, the evidence was indirect and based on changes in enzyme activity in response to different factors. For an enzyme to be established as a physiological substrate for cyclic AMPdependent protein kinase the following additional criteria have to be satisfied, as defined by Krebs (63)and modified by Nimmo and Cohen (64). 1 . A protein substrate for cyclic AMP-dependent protein kinase should exist that bears a functional relationship to the process mediated by cyclic AMP. The rate of phosphorylation of that protein, in its native state, should be adequate to account for the speed at which the process occurs in vivo in response to cyclic AMP.

6. HORMONE-SENSITIVE LIPASE

157

2. The function of the protein should be shown to undergo a reversible alteration in vitro by phosphorylation and dephosphorylation, catalyzed by cyclic AMP-dependent protein kinase and a protein phosphatase. 3. A reversible change in the function of the protein should occur in vivo in response to cyclic AMP. 4. Phosphorylation of the protein should occur in vivo in response to a hormone, at the same site(s) phosphorylated by cyclic AMP-dependent protein kinase in vitro. Only the third of these criteria was strictly fullfilled by the work forming the basis for the lipolytic activation cascade hypothesis. Phosphorylation of the HSL protein, required by the first, second, and fourth criteria was not demonstrated. As described in Section III,A, the first and second criteria were met by work with the isolated lipase. Thus, the enzyme is phosphorylated by cyclic AMP-dependent protein kinase and is thereby activated (2, 16). The rate of phosphorylation is comparable to that in intact fat cells in response to lipolytic hormones (16,49), and is also comparable to the rate of phosphorylation of other substrates by the cyclic AMP-dependent protein kinase (16). Phosphorylated lipase is dephosphorylated and concomitantly deactivated by a protein phosphatase from the adipose tissue (47). In the following, work is reviewed that demonstrates that, in response to lipolytic hormones, the enzyme is phosphorylated in the intact fat cell (65) on the same site as by cyclic AMP-dependent protein kinase in vitro (66), as required by the fourth criterion. Procedures have been developed to measure changes in activity and state of phosphorylation of HSL in intact adipocytes in suspension. Under experimental conditions when the fatty acid:albumin ratio is kept low and the incubation time short, negligible reesterification takes place (67) and the rate of fatty acid release is a measure of the rate of lipolysis and hence an expression of the activity of HSL (68). This activity can be continuously monitored by pH-stat titration of protons that are released with the fatty acids in a stoichiometric amount (69, 70). The technique allows the short-term kinetics of hormone effects on HSL in isolated adipocytes in suspension to be monitored (see Fig. 9 in Section 111,D); it has been combined with sampling of cells from the same reaction vessel for parallel determination of the enzyme phosphorylation state (49, 66, 71). Thus, the effects of different hormones or pharmacological agents on the enzyme activity in intact cells can be directly related to its phosphorylation state. Phosphorylation of HSL in situ in intact cells has been demonstrated after purification of the enzyme from adipocytes that had been preincubated with [32P]phosphateand treated with noradrenaline (65), and from 3T3-L 1 adipocytelike cells treated with isoproterenol and isobutylmethylxanthine (28). To follow the changes in the extent of phosphorylation of the lipase adipocytes are prelabeled with [32P]phosphateuntil a constant specific radioactivity of [32P]ATPis

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reached, and the incorporation of 32P in the HSL M, = 84,000 phosphoprotein band is measured by SDS-PAGE (49, 66, 71). HSL incorporates 32Pin close parallel with the increase in the specific radioactivity of [32P]ATP, reaching and remaining on a steady-state level, whereas the enzyme activity is unchanged (Fig. 5A) (49). The phosphate rapidly turns over at a single serine residue in the basal phosphorylation site (Fig. 6A) (66).Assuming uniform labeling of [32P]ATP(260 dpm per pmol) the steady-state incorporation of 32P into the lipase protein (373 ? 17 dpm per 25 p.1 of packed cells) is equivalent to 60 ? 3 pmol [mean SE (n = 5)] of phosphate per ml of packed

*

B

A

Timemn

Time. min

FIG. 5 . Effects of noradrenaline, propranolol, and insulin on the extent of phosphorylation and activity of HSL. Lipase activity was measured as fatty acid release by continuous pH-stat titration. 32P-Hormone-sensitive lipase was determined as described in the text and the extent of phosphorylation related to the basal phosphorylation level (= 100%). (A) Noradrenaline (300 nM) and prowere added at the time-points indicated by vertical lines; A-A, refers to an pranolol (10 incubation without added noradrenaline or propranolol (= basal conditions). (B) Time course of effects of insulin. Following 40 min preincubation as in (A). 100 nM noradrenaline and 700 pM insulin was added as follows: -0, noradrenaline added at zero time; C W , insulin added at zero time (fatty acid release < 0.03 pmol/min/ml PCV and therefore not illustrated); A-A or continuous line labeled NA + INS, noradrenaline and insulin added together at zero time; 0-- -0or broken line, noradrenaline added at zero time, followed by insulin as indicated after 6 min. No treatment controls were as illustrated in (A) (basal conditions) [from Ref. (1901.

a)

6. HORMONE-SENSITIVE LIPASE

159

FIG. 6. Effects of noradrenaline and insulin on the extent of phosphorylation of the regulatory and basal phosphorylation sites of HSL. Cells from an experiment similar to that in Fig. 5 were sampled. (A) Before addition of hormones (at a in Fig. 5 ) ; (B) after addition of noradrenaline (at b in Fig. 5 ) ; (C) after addition of noradrenaline followed by insulin (at c in Fig. 5 ) . HSL was isolated by SDSPAGE and after digestion with S. uureus V8 proteinase and trypsin subjected to phosphopeptide mapping by two-dimensional electrophoresis thin-layer chromatography and autoradiography; “x” indicates the point of application, and the ring indicates the position of dinitrophenyllysine. For explanation of “basal” and “regulatory,” see text [from Ref. (66)].

cells. The amount of lipase-protein (from the data in Fig. 9, Section II1,D) per ml of packed fat cells is calculated to be approximately 75 pmol of lipase M, 84,000 subunit [total amount of enzyme activity is 0.5 pmol/min/ml of packed cells, HSL molar-specific activity is 600 mol/s/mol (2, 16), corresponding to 110 mol/s/mol against triacylglycerol in the presence of monoacylglycerol lipase

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(2)]. These estimates are clearly colisistent with near stoichiometric phosphorylation of the basal phosphorylation site. The fast-acting lipolytic hormones (e.g., noradrenaline, ACTH, and glucagon) rapidly increase the extent of phosphorylation of HSL and, after a time lag of 1-2 min, also increase the enzyme activity (49) (Fig. 5A). This phosphorylation, which is confined to the specific regulatory phosphorylation site (cf., Section III,A) (Fig. 6B) (66), is most likely near stoichiometric since the increase in the total phosphorylation is about twofold. The P-adrenergic antagonist propranolol rapidly reverses the phosphorylation and activity of the enzyme to the basal level (Fig. 5A) by causing a net dephosphorylation of the regulatory phosphorylation site (66). The reversibility and the lack of effect of the hormones on the specific radioactivity of the cellular [32P]ATP (49) establish that the hormonal effects indeed reflect changes in enzyme phosphorylation state and are not due to an increased turnover of enzyme-bound phosphate at the regulatory site. Previous work indicated that insulin counteracted the lipolytic effect of fastacting lipolytic hormones by converting HSL from a more active (presumably phosphorylated) to a less active (dephosphorylated) form [reviewed in Refs. (3, 5 ) ] . Using the techniques previously described we have directly examined the effects of insulin on HSL activity and state of phosphorylation. Insulin at physiological concentrations (< 1 nM), when added before the preincubation with [32P]phosphateor when the basal 32P-incorporation was at a steady state, was found to have no effect on the extent of the basal phosphorylation or on the time to reach the steady state (Fig. 5B). Thus, neither the extent nor the turnover of the basal phosphorylation is affected by insulin. However, when added before or with the lipolytic hormones insulin abolishes the anticipated increase in extent of phosphorylation and activity of HSL (Fig. 5B). Insulin added after lipase phosphorylation and activity has been maximally stimulated with noradrenaline, glucagon, or ACTH causes a rapid net dephosphorylation of HSL (49), at the regulatory phosphorylation site (Fig. 6C) (66). This reduces the total extent of phosphorylation to that of the basal steady-state level (Fig. 5B) and after a short time lag reduces the enzyme activity to the very low basal level (Fig. 5B) (49, 70). The effects of increasing concentrations of insulin have demonstrated a quantitatively close correlation between the decrease of the extent of dephosphorylation and the extent of inactivation of the lipase, with half-maximal effect of insulin at around 30 pM (5, 72). These findings demonstrate that fast-acting lipolytic hormones and insulin control adipose tissue lipolysis specifically by controlling the phosphorylation state of the same serine residue in the regulatory phosphorylation site of HSL. Growth hormone is a slow-acting lipolytic hormone that increases lipolysis with a time lag of several hours, probably through increased synthesis of enzymes involved in the lipolytic activation process (8). In addition, growth hor-

161

6. HORMONE-SENSITIVE LIPASE

mone has a fast-onset insulin-like, antilipolytic effect on adipocytes that have been preincubated for 3 h in the absence of the hormone, or directly on cells from hypophysectomized rats (71, 73, 74). Like insulin, physiological concentrations of growth hormone cause a rapid net dephosphorylation of HSL, presumably at the regulatory phosphorylation site, and this is accompanied by inactivation of the enzyme (71). c . MECHANISMS OF ACTIONFOR THE SHORT-TERM CONTROL LIPOLYTIC HORMONES AND INSULIN

BY

The demonstration that the fast-acting lipolytic hormones enhance the activity of HSL through a cyclic AMP-dependent protein kinase-catalyzed phosphorylation of its regulatory site establishes the mechanism for the control of adipose tissue lipolysis shown in Fig. 7.

CATECHOLAMINES ACTH, GLUCAGON

1

RECEPTOR

1

ADENYLATE CYCLASE

1

CAMP

1

CAMP-PrK CE C

4 FA

t

TG

1

Ir

1.2-DG + FA 2-MG

1

+

FA

1

GLYCEROL

t

FA

PROTEIN PHOSPHATASES

Frc. 7. Control of adipose tissue lipolysis by fast-acting lipolytic hormones and insulin through reversible phosphorylation of HSL. Arrow, with insulin, indicates the net dephosphorylation of the regulatory site of the enzyme, accounting for insulin’s antilipolytic effect. The dimeric structure of the enzyme is tentative. Encircled-P, phosphate at the regulatory sites; CAMP-PrK, cyclic AMPdependent protein kinase; CE, cholesterol ester; TG, DG, MG, tri-, di- and monoacylglycerol; FA, fatty acid.

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BELFRAGE

The role of cyclic AMP in the control of lipolysis was noted soon after the discovery of the nucleotide (14, 37). Cyclic AMP analogs and, in digitoninpermeabilized fat cells, cyclic AMP itself elicit the same lipolytic response (28, 37, 75-77) and increased phosphorylation of HSL (78)as the fast-acting lipolytic hormones. Although questioned at times, there is little doubt that increases of cellular cyclic AMP correlate with the lipolytic rate if appropriate conditions (incubation time, hormone and cell concentration) are used [see, e.g., Refs. (7981)].Very small increases of cyclic AMP markedly activate lipolysis; a doubling of the basal concentration is enough to obtain a maximal rate of lipolysis (82, 83) [and maximal phosphorylation of HSL, see Ref. (72)].Changes in the activity of cyclic AMP-dlependent protein kinase parallel those of cyclic AMP in studies where both parameters have been measured (84-88) [cf., cyclic AMP and cyclic AMP-dependent protein kinase in the regulation of steroidogenesis, Refs. (89941. In a carefully controlled investigation the activation of lipolysis by a number of lipolytic agents (isoproterenol, ACTH, glucagon, and forskolin) was shown to be quantitatively related to increased cyclic AMP-dependent protein kinase activity (92).Maximal lipolysis was obtained at about 30% activation of the protein kinase. Adenylate cyclase inhibitors and other agents known to reduce cyclic AMP decreased the lipolysis rate in close correlation with lowered protein kinase activity (92). The hormonal signal transduction chain involving cyclic AMP is in many respects well characterized. Most of its different components have been purified from various sources and a theoretical background describing the potentials of such systems has been given (93-97). To understand the characteristics of the system controlling HSL in the fat cell a quantitative evaluation of the components constituting the lipolytic activation cascade is pertinent. In a single fat cell it can be estimated that the order of lo3 P-adrenoceptors need to be occupied (98) to activate some lo5 adenylate cyclase catalytic subunits [assuming them to represent 10W5 of the cell protein (99)]in order to double the amount of cyclic AMP from 10 X lo6 to 20 X lo6 molecules (72, 82), an increase required to obtain maximal lipolysis. Through this increase of cyclic AMP about 30% (i.e., lo6 copies) of the cyclic AMP-dependent protein kinase (92, 100, 101) is activated and phosphorylates the regulatory site of 3 X lo6 copies of HSL. In this lipolytic activation cascade almost all of the amplification (about 102-fold) is obtained in the plasma membrane at the receptor-adenylate cyclase level; the remaining components of the signal chain and the target enzyme are present at nearly equimolar concentrations. The relatively high basal steady-state concentration of cyclic AMP in the cell [of the order of 10 pM; the same magnitude as the concentration for half-maximal binding of cyclic AMP to the protein kinase (102)]should confer a very high sensitivity to the system and explain why only a fractional increase in the cyclic AMP concentration is enough to markedly

-

-

-

6. HORMONE-SENSITIVE LIPASE

163

stimulate lipolysis [in addition, cyclic AMP-binding to the protein kinase exhibits positive cooperativitiy (103)]. Any protein kinase that is already activated by the basal steady-state concentration of the nucleotide should be blocked by the specific inhibitor protein of cyclic AMP-dependent protein kinase (102), thereby allowing an additional small increase of cyclic AMP to be expressed as protein kinase activity with maximal efficiency. It is noteworthy that most of the time lag in the hormonal response should be expected at the plasma membrane level, during receptor activation of the adenylate cyclase. The high and equimolar concentrations of all the subsequent intermediate steps should ensure an almost instantaneous phosphorylation of HSL. Moreover, cyclic AMP-dependent inhibition of protein phosphatase-1 through the phosphorylation of inhibitor- 1 (104106) (both are also likely to be found in the order of lo6 copies per fat cell) could provide additional sensitivity by coordinating increased phosphorylation with decreased dephosphorylating activity (107). A characteristic and recurrent finding has been a “peaking” of cyclic AMP in response to the lipolytic hormones (37, 82, 87, 88, 108). Lipolysis [and HSL phosphorylation (72)] reaches and remains at a maximal level, whereas the total cellular cyclic AMP reaches a peak after a few minutes and then returns to the basal level in the continued presence of the hormone. To some extent this may be a methodological artifact (101) but the above and other findings (81, 109-112), discussed in more detail elsewhere (3,indicate that cyclic AMP may be, at least functionally, compartmentalized. Such compartmentation may be particularly plausible in the fat cell, in which the thin rim of cytosol (0.2-0.4 pm, fat cell diameter 30- 100 pm) surrounding the central lipid droplet may be conceived of as an almost two-dimensional cell compartment. Insulin exerts its antilipolytic effect by shifting the steady-state distribution of the phospho- and dephospho-form of the regulatory site of HSL towards the dephospho-form (Figs. 7 and 8). The hormone must therefore act to decrease the rate of phosphorylation and/or to increase the rate of dephosphorylation of this phosphorylation site by reducing the activity of cyclic AMP-dependent protein kinase and/or activating protein phosphatase- 1 and -2A. Much evidence indicates that the insulin signal to a large part, perhaps exclusively, is mediated through reduction of the hormone-stimulated steady-state level of cyclic AMP (Fig. 8) (i.e., through reversal of the action of the fast-acting lipolytic hormones). Indeed, this was quite early proposed (82, 113-115). It has also been demonstrated that after selective P-adrenergic- (86) or ACTH- (87) stimulation of adipocytes, under appropriate conditions, the inhibition of lipolysis by insulin is parallelled by reduction of cyclic AMP and cyclic AMPdependent protein kinase activity. In another study, using adipocytes that had been prepared and incubated under conditions during which endogenously produced adenosine was controlled, and with up to maximally stimulated lipolysis, the reduction of cyclic AMP-dependent protein kinase activity was quantitatively

164

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0 CAMP-PrK

, / p 7

/ _/-

INSULIN

1 Tyr-PrK

,/’

PDE’ I

AC

n@

HSL

HSL-

Ser-PrK

FIG.8. Postulated mechanisms for the antilipolytic effect of insulin in fat cells exposed to fastacting lipolytic hormones. Insulin lowers the rate of phosphorylation of the regulatory phosphorylation site of HSL as the result of decreased cyclic AMP-dependent protein kinase (CAMP-PrK) activity, secondary to reduction of hormone-elevated cellular cyclic AMP. This can also lead to increased protein phosphatase (PrP)-I activity by decreasing the inhibitor-1 (1-1) inhibition of this phosphatase, as the result of decreased cyclic AMP-dependent phosphorylation of the inhibitor. Cyclic AMP reduction is obtained through activation of plasma membrane-bound, low K,,,phosphodiesterase (PDE), and, possibly, through inactivation of adenylate cyclase (AC). An increased rate of dephosphorylation may also be caused by cyclic AMP-independent activation of protein phosphatase-l and/or -2A, mediated, for example, through phosphorylation of a phosphatase regulator protein. The initial events in the transduction of the insulin signal at the plasma membrane level may involve activation of the insulin receptor tyrosine protein kinase (Tyr-PrK), serine protein kinase(s) (Ser-PrK), and/or a putative guanine nucleotide regulatory protein, (N,,s). Solid arrows indicate regulatory mechanisms demonstrated or supported by experimental evidence from adipose tissue; broken arrows indicate possible regulatory mechanisms. For references, see text.

correlated to the inhibition of lipolysis (116). Only with supramaximally stimulated lipolysis (cyclic AMP-dependent protein kinase activity ratio above 0.3) was a cyclic AMP-unrelated inhibition of lipolysis found (116). In addition to a decrease in the rate of phosphorylation of the regulatory site of HSL, cyclic AMP reduction could be expected to cause an increased rate of dephosphorylation of this site by lowering the extent of phosphorylation of inhibitor-1 and thus activating protein phosphatase- I (Fig. 8). The cellular concentration of cyclic AMP can be reduced by a decreased rate of its formation and/or an increased rate of degradation. Insulin has been shown

6 . HORMONE-SENSITIVE LIPASE

165

to activate the membrane-associated low K , cyclic AMP phosphodiesterase which may be partly associated with the plasma membrane (126) in adipocytes (117-127) and also in the 3T3-Ll adipocyte-like cells (128-130). In the latter cells selective inhibition of the membrane-associated but not of soluble cyclic AMP phosphodiesterase abolished the anti-lipolytic effect of insulin ( I31), indicating that the particulate low K , cyclic AMP phosphodiesterase has a crucial function in mediating insulin’s effects on HSL (Fig. 8). In very recent work the insulin-sensitive particulate cyclic AMP phosphodiesterase has been purified to apparent homogeneity from rat adipose tissue and identified with a 64 KDa protein (after SDS-PAGE) (132). In the liver insulin has also been reported to activate a plasma membrane-associated low K , phosphodiesterase, in a process dependent on cyclic AMP and on phosphorylation of the enzyme (133-135). There are conflicting reports whether the activity of adenylate cyclase is controlled by insulin. The hormone has been reported to inhibit hormonally activated adenylate cyclase (I14, 136-138), but others have failed to find any insulin effect on the enzyme (118, 139, 140). The initial events at the plasma membrane level through which insulin enhances the particulate cyclic AMP phosphodiesterase activity and, possibly, inhibits the adenylate cyclase activity are unknown. It can be calculated that maximal antilipolysis is obtained through the occupation of less than 2000 insulin receptors per fat cell [from Refs. (141, 142)].With insulin-sensitive cyclic AMP phosphodiesterase and adenylate cyclase present in the order of lo5 copies per cell (99, 118, 129, 134) the amplification required is about 102-fold (i.e., similar to that during activation of lipolysis by the fast-acting lipolytic hormones). In analogy with the mode of action of these hormones it has been proposed that insulin should reverse their elevation of cellular cyclic AMP through a specific “guanine nucleotide regulatory protein,” a putative ‘‘N,Ns’’ (143) (Fig. 8). Alternative mechanisms include modulation of the particulate cyclic AMP phosphodiesterase by the insulin receptor tyrosine protein kinase (Fig. 8), or, perhaps more likely, by an associated serine protein kinase, either directly or via the tyrosine protein kinase (144) (Fig. 8). The possibility that insulin acts to increase the rate of dephosphorylation also through a cyclic AMP-independent mechanism must be considered (Fig. 8). Insulin is known to cause increased phosphorylation of several intracellular fat cell proteins [for review, see Refs. (145, 146)] through stimulation of one or several cyclic AMP-independent protein kinases. Insulin activation of a plasma membrane-bound serine protein kinase has been demonstrated (145, 147-149); it has been proposed that the hormone causes a dissociation of such a protein kinase with subsequent release of active protein kinase into the cytosol compartment (145, 148, 149). Concievably, a protein kinase, activated and translocated to intracellular target proteins, could enhance the activity of a protein phosphatase [e.g., through phosphorylation of a regulatory protein; cf. Ref. (ISO)] (Fig. 8).

I66

P. STRALFORS, H . OLSSON, AND P. BELFRAGE

Such phosphorylation cascade mechanisms could provide for the lo3- to 104-fold amplification needed to directly control these target proteins (likely to be found in lo6 copies per cell, as previously discussed) (Fig. 8). It is possible that the cyclic AMP-unrelated inhibition of supramaximally stimulated lipolysis ( I 16), is mediated by such a protein phosphatase activation. The similarity between the time course of effects of growth hormone and insulin on the extent of phosphorylation and activity of HSL (see Section III,B) suggests that the two hormones act at least partly through the same mechanisms (71). Like insulin, growth hormone has been demonstrated to reduce catecholamine-stimulated cyclic AMP concentrations, cyclic AMP-dependent protein kinase activation (74),and lipolysis (71, 73, 74, 151); to activate low K,, phosphodiesterase (74);to accelerate glucose transport and metabolism (152); and to oxidate leucine (153). Furthermore, in skeletal muscle, it has been reported to inhibit adenylate cyclase (154). Since growth hormone is the only peptide hormone except oxytocin (155, 156) that, under certain conditions, can have an insulin-like, antilipolytic effect, studies of its mode of action could become particularly interesting for an understanding of the mechanism of action of insulin. FOR REGULATION D. OTHERMECHANISMS OF ADIPOSETISSUELIPOLYSIS

Control of lipolysis in fat cells exclusively through phosphorylation-dephosphorylation of HSL has been questioned (157-159). The question has been provoked by the fact that the high extent of activation of lipolysis in fat cells (more than 20-fold) is not preserved in extracts of the cells, and phosphorylation of isolated HSL causes a 3-fold increase of enzyme activity (see Fig. 3) (10-fold with chicken HSL). The following illustrates this and summarizes observations and calculations demonstrating that the difference in activatability may simply be the result of very different conditions prevailing during lipolysis in the intact cell, and in vitro. The subject has been treated in detail elsewhere (4). Disruption of rat fat cells by sonication for a few seconds decreases the apparent noradrenaline activation of lipolysis from more than 50-fold to 1.5- to 2-fold (Fig. 9). Most of this loss of activation results from the, at least, 10-fold elevation of the lipolysis rate in the nonstimulated fat cell preparation (Fig. 9), whereas the decrease of the hormonally stimulated lipolysis rate is only 50-70% [cf., Refs. (2, 77)]. This decrease is probably due to dephosphorylation of HSL during the disruption of the cells (NaF could not be used for inactivation of protein phosphatases since it inhibits HSL) [cf. Ref. (41. The 10-fold increase of basal lipolysis is likely to be due to a dramatic increase of substrate availability. It has been estimated ( 4 )that in the fat cell, assuming a single lipid droplet, only about 100 triacylglycerol molecules are available in the

I67

6 . HORMONE-SENSITIVE LIPASE

.-

E

3-

cell disruption b y sonication

FIG.9. Effects on HSL activity by fat cell disruption. Two samples of fat cells [ 5 % (v/v)] were identically incubated as in Fig. 5 , in the absence or presence of 200 nM noradrenaline (NA); HSL activity was followed continuously by pH-stat titration of released fatty acids. The cells were disrupted by sonication (2 s burst at maximal intensity) at the time-point indicated by the arrow; dithioerythritol (0.5 mM final concentration), leupeptin (20 pglml), and pepstatin (20 pg/ml) had been added immediately before. Dotted lines indicate initial disturbances in pH stat titration due to the sonication, PCV, packed cell volume.

substrate interfere for each HSL dimer molecule. After sonication of the cells as in Fig. 9, or under in vitro assay conditions with artificial substrate emulsions, 2000- 10,000 triacylglycerol molecules are calculated to be available for each enzyme molecule. As discussed in Section III,A, the activatability of isolated HSL depends on substrate availability; readily available substrate results in relatively high nonphosphorylated (basal) enzyme activity and little additional activation by phosphorylation. Although these estimates are crude they indicate clear-cut differences between the conditions in the intact cell and in cell-free preparations, and they demonstrate a rationale that is likely to explain the discrepancies in HSL activatability. Moreover, both dibutyryl cyclic AMP (78), and cyclic AMP in digitonin-permeabilized fat cells ( 7 3 , activate lipolysis [and phosphorylates HSL (78)]to the same extent as catecholamines. Therefore, any additional mechanism for hormonal activation of lipolysis in fat cells besides HSL phosphorylation must involve phosphorylation of another target protein. Although this cannot be entirely excluded, no evidence for such a mechanism has been provided. Wise and Jungas (157) reported that exposure of rat fat cells to adrenaline increased lipolysis of endogenous (homogenized tissue lipids) but not of exogenous substrate (sonicated trioleoylglycerol emulsion) by cell-free extracts of these cells. Incubation with cyclic AMP-dependent protein kinase, on the other

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BELFRAGE

hand, activated lipase-catalyzed hydrolysis of exogenous, but not of endogenous substrate. The data were interpreted to show the existence of a second pathway of lipolysis activation in response to adrenaline, besides that involving cyclic AMPmediated phosphorylation of HSL. This pathway, termed ‘‘substrate activation,” was suggested to involve a hormonally stimulated generation, or activation, of a factor that associated with the endogenous substrate and facilitated its hydrolysis by the HSL. An alternative explanation to the reported findings has been presented in detail elsewhere (4);in essence, it proposed that the “substrate activation” could reflect changes of the substrate interface composition that result from the hormone-induced phosphorylation-activation of the HSL, but were not the cause of the activation of the lipolysis. Thus, although it cannot be completely excluded that other mechanisms exist that act in parallel, there is little doubt that the cyclic AMP-dependent phosphorylation of HSL is sufficient to explain the effects of fast-acting lipolytic hormones on lipolysis.

IV. Possible Role as a Hormone-Activatable, Multifunctional Tissue Lipase

HSL is the most characteristic and functionally important enzyme of adipose tissue. It was believed to be unique to the adipocyte; it had not been identified in any other cell type. However, as discussed in Section 11, HSL, or a very similar enzyme (referred to as HSL in the following), is also present in the adrenal cortex and in corpus luteum. Moreover, the reported properties of neutral acylglycerol lipases and/or cholesterol ester hydrolases in a number of other cell types and tissues suggest that HSL may be even more widespread. In fact a role for the enzyme as a general, hormone-activatable tissue acylglycerol lipase-cholesterol ester hydrolase is becoming evident. A. IN STEROID-PRODUCING TISSUES In the steroid-producing tissues HSL catalyzes the hydrolysis of cholesterol ester to provide free cholesterol, an important ACTH-regulated step that is probably a major site for short-term hormonal control of the rate of steroidogenesis. More detailed accounts of this process and how it is believed to be controlled can be found elsewhere (5, 36); a short overview is given in this section. Cholesterol for biosynthesis of steroid hormones is mainly derived from lipid droplets, consisting of a mixture of cholesterol ester and triacylglycerol, stored in the cytosol of adrenal cortex cells (160). The HSL-catalyzed hydrolysis of these cholesterol esters to free cholesterol, and its subsequent transport into the mitochondria, constitute the first steps in steroidogenesis. (Fig. 10). Several lines of

169

6. HORMONE-SENSITIVE LIPASE

+

ACTH CAMP

1 CHOLESTEROL ESTER

HSL

HSL-

@

u

MITOCHONDRIA CHOLESTEROL

-D

PRDTEIN PHOSPHATASES

1 PREGNENOLONE

i

+

4 STEROIDS

FIG. 10. Proposed mechanism for ACTH regulation of HSL activity in the control of adrenal steroidogenesis.

evidence indicate the relationship between the lipase activity and the rate of steroid synthesis (160-167); ACTH has been shown to control the activity of the enzyme (36, 160-162, 164) and cyclic AMP to mediate the short-term steroidogenic response to ACTH in several types of steroid-producing cells (8991). A number of earlier reports indicated that reversible phosphorylation is involved in the activation of the enzyme by ACTH in adrenal cortex. The indications included activation in vitro by cyclic AMP-dependent protein kinase, inhibition of this activation by the specific inhibitor protein for the kinase, reduction of the activatability in tissue extracts from ACTH-treated animals, and reversibility of the activation by a protein phosphatase preparation (36, 161, 16.5169).

Identification of HSL from bovine adrenals made it possible to provide direct experimental proof for its phosphorylation and activation by cyclic AMP-dependent protein kinase (34),confirmed by work with the corresponding rat enzyme (170). Therefore, it seems probable that the adrenal cortex enzyme also is regulated by hormones through cyclic AMP-mediated, reversible phosphorylation (Fig. lo), as has been shown for the adipose tissue HSL. However, the four criteria for an enzyme to be a physiological substrate for cyclic AMP-dependent protein kinase (Section III,B) have not been met for the adrenal enzyme; only

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criterion 3 and, to some extent, criteria 1 and 2 have been fulfilled. Therefore, much experimental work remains to be done, especially with regard to the effects of ACTH on the phosphorylation of the enzyme in intact cells. These studies should be considerably facilitated by the experience gained from the studies of the adipose tissue enzyme. Steroidogenesis in other steroid-producing tissues is likely to be regulated in a similar manner to that in the adrenal cortex. Enzyme from bovine corpus luteum is also phosphorylated and activated by cyclic AMP-dependent protein kinase (35). Luteinizing hormone has been reported to activate this enzyme accompanied by increased steroidogenesis in the corpus luteum (171), via cyclic AMPmediated activation of cyclic AMP-dependent protein kinase (89). Testicular interstitial cells are also known to contain a neutral cholesterol ester hydrolase (172), and follicle-stimulating hormone controls the activity of cyclic AMPdependent protein kinase in these cells (90, 91). Thus, the ovarian and the testicular enzymes may both be regulated in a way analogous to the adrenal enzyme (Fig. 10).

B . IN OTHERTISSUES In addition to adipose tissue and steroid-producing organs the presence of HSL has been demonstrated in cultured cells of mesenchymal origin, including mouse 3T3-Ll adipocyte-like cells (28, 31), and in preadipocytes (173). The triacylglycerol lipase of brown adipose tissue is likely to be the HSL, which in response to an adrenergic stimulation of thermogenesis hydrolyzes the stored acylglycerols to fatty acids (174). Through P-oxidation these fatty acids serve as the major thermogenic fuel, they also trigger uncoupling of oxidative phosphorylation by binding to and activating an uncoupling protein in the mitochondrial inner membrane (174). It is interesting that, through activation by the second messenger cyclic AMP, the lipase generates a “third messenger”-fatty acids-for the acute hormonal stimulation of thermogenesis. Cytosolic neutral triacylglycerol lipase or cholesterol ester hydrolase activity with several of the properties of HSL has been reported to be present in other mammalian tissues, for example, heart muscle (175-179), arterial tissue (180182), liver (183), placenta (184), and macrophages (185).The properties include inhibition by sulfhydryl reagents and DFP (181-183) and, most importantly, activation by cyclic AMP-dependent protein kinase (182-185). None of these mammalian tissue lipases have been purified enough to allow identification with HSL. However, the similarity between the enzymological characteristics is striking and a further comparison is clearly indicated. The possible role of HSL in these other tissues is a matter of speculation. The ability of the enzyme, unique for a lipase, to be controlled via cyclic AMPdependent phosphorylation indicates that it could provide a hormone-sensitive

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acylglycerol-cholesterol ester lipase function to these cells, perhaps as a complement to a separate very active triacylglycerol lipase with hepatic lipase-like properties that has been found in adrenal cortex (I70) and macrophages (186). In analogy with the role of fatty acids as an uncoupling signal to the mitochondria during thermogenesis in brown adipose tissue HSL could have a more general function to generate fatty acids as a regulatory message. Fatty acids or the acylCoA esters have been shown to affect the activities of several enzymes [e.g., CTP:phosphocholine cytidylyltransferase (187),acetyl CoA carboxylase (I88), and cyclic GMP-stimulated phosphodiesterase (I89)].

V. Conclusions and Perspectives The results reviewed here establish that fast-acting lipolytic hormones control adipose tissue lipolysis by a reversible cyclic AMP-dependent protein kinasecatalyzed phosphorylation of HSL. Moreover, insulin exerts its antilipolytic effect through a net dephosphorylation of the same regulatory phosphorylation site that is specifically phosphorylated by cyclic AMP-dependent protein kinase after stimulation by the lipolytic hormones. It is concluded that the net dephosphorylation of HSL in response to insulin is mainly, or exclusively, a reversal of the cyclic AMP elevation by the lipolytic hormones. Available evidence indicates that insulin reduces the cyclic AMP concentration through activation of a specific membrane-bound cyclic AMP phosphodiesterase, perhaps with a concerted inhibitory effect on the adenylate cyclase. The molecular mechanisms whereby insulin affects the activity of cyclic AMP phosphodiesterase are unknown and clearly of greatest importance. The very similar effects of insulin and growth hormone as antilipolytic agents may prove a great help in elucidating these initial events upon hormone-receptor binding. The cyclic AMP-independent phosphorylation of HSL at a second serine residue, termed the basal phosphorylation site, does not directly affect the enzyme activity and does not seem to be influenced by the hormones. The function and importance of this “silent” phosphorylation can be experimentally examined with the site-specific glycogen synthase kinase 4 and cyclic GMP-dependent protein kinase. An examination of protein phosphatases involved in the dephosphorylation of HSL at the regulatory and at the basal phosphorylation sites, and how these phosphatases are affected by hormones, has just begun and is obviously of prime importance. The mechanisms whereby phosphorylation of the HSL regulatory phosphorylation site is expressed as an enhanced enzyme activity are poorly understood. It is likely that use of new experimental systems (e.g., monolayer tech-

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niques) in combination with lipase that has been reconstituted into phospholipid vesicles will help to answer this question. The demonstration that HSL is responsible for hormonally controlled provision of free cholesterol for steroidogenesis has broadened the view of the role of HSL considerably and it is proposed that HSL is a widespread tissue acylglycerol-cholesterol ester hydrolase that confers hormone-sensitivity on this function in many tissues. The cell type and the substrates present in a particular cell or tissue determine the physiological role of the enzyme (e.g., free fatty acids released to the blood, steroid hormone production, or thermogenesis). Identification of HSL and elucidation of its role in other tissues are of great interest. ACKNOWLEDGMENTS We wish to acknowledge contributions by our colleagues Drs. Per Bjorgell, Hans Eriksson, Gudrun Fredrikson, Nils Osten Nilsson, and Staffan Nilsson during various parts of the work in our laboratory on which this review is based, and the assistance of Dr. Lubos Zan in preparing Figs. 7, 8, and 10. Unpublished results used for preparing Fig. 5 were kindly provided by Dr. Nils Osten Nilsson. The skilful technical assistance of Ingrid Nordh, Birgitta Danielsson, Aniela Szulscynski, and Stina Fors is gratefully acknowledged, as well as the help of Ruth Lovin in preparing the manuscript. Cyclic nucleotide-independent protein kinases and protein phosphatases purified from skeletal muscle were generously provided by the laboratory of Prof. Philip Cohen in Dundee, Scotland. Financial support was obtained from the following sources: The Swedish Medical Research Council (grant No. 3362); A. Pihlsson’s Foundation, Malmo; Thorsten and Elsa Segerfalk’s Foundation for Medical Research and Education, Helsingborg; A. 0. Swird’s Foundation, Stockholm; P. Hlkansson’s Foundation, Eslov; Nordic Insulin Foundation, Copenhagen; the Swedish Diabetes Association, Stockholm; the Royal Physiographic Society, Lund; and the Medical Faculty of the University of Lund. The work with cyclic AMP-independent protein kinases was initiated in the laboratory of Prof. Cohen in Dundee, Scotland, under a FEBS short-term fellowship to Peter Strhlfors.

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6. HORMONE-SENSITIVE LIPASE 93. 94. 95. 96. 97. 98. 99. 100. 101.

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144.

145. 146. 147. 148. 149. 150. 151. 152. 153. 154. 155. 156. 157. 158. 159. 160.

161. 162. 163. 164.

165. 166. 167. 168. 169. 170. 171. 172. 173. 174. 175. 176. 177. 178. 179.

6. HORMONE-SENSITIVE LIPASE 180. 181. 182. 183. 184. 185. 186. 187. 188. 189. 190.

177

Howard, C. F., and Portman, 0. W. (1966). EEA 125. 623. Kothari, H. V., Bonner, M. J., and Miller, B. F. (1970). EEA 202, 325. Hajjar, D. P., Minick, C. R.,and Fowler, S. (1983). JEC 258, 192. Deykin, D., and Goodman, D. S. (1962). JEC 237, 3649. Chen, L., and Morin, R. (1971). BEA 231, 194. Khoo, J. C., Mahoney, E. M., and Steinberg, D. (1981). JEC 256, 12659. Khoo, J. C., Vance, J. E., Mahoney, E. M., Jensen, D., Wancewicz, E., and Steinberg, D. (1984). Arteriosclerosis 4, 34. Vance, D. E., and Pelech, S.L. (1984). TIES 9, 17. Bortz, W. M., and Lynen, F. (1963). Eiochem. Z. 337, 505. Yamamoto, T., Yamamoto, S., Manganiello, V. C., and Vaughan, M. (1984). AEE 229, 81. Belfrage, P., Donnkr, J., Eriksson, H., and Striilfors, P. (1986). In “Mechanisms of Insulin Action.” (P. Belfrage, J. Donnkr, and P. Striilfors, eds.), p. 323. Elsevier, Amsterdam.

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7

HydroxymethylglutarylCoenzyme A Reductase DAVID M. GIBSON REX A. PARKER Department of Biochemistry Indiana University School of Medicine Indianapolis, Indiana 46223

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Topology . . . . ................................ 111. Multivalent Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Overview of Cholesterol Homeostasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Regulation . . . . . . . . . . .

C. The Modulating Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Intracellular Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Early Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ..... .... . C. Feedback Control of Reversible Phosphorylation . . . D. Endocrine Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Cellular Proliferation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Other ..................................... VI. Reversible Degradation . A . Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. In Vitro Model System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Phosphorylation-Activated Proteolysis .. D. Intracellular Proteolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I79 THE ENZYMES. Vol. XVIII Copyright 0 1987 by Acndemic Press. Inc. All rights of reproduction in any l o r n reserved

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DAVID M . GIBSON AND REX A . PARKER

Introduction

Hydroxymethylglutaryl-CoA (HMG-CoA) reductase (EC 1.1.1.34), less well known as meva1onate:NADP oxidoreductase (CoA acylating), is an enzyme that is tightly bound to the endoplasmic reticulum of higher eucaryotic (metazoan) cells. The reaction it catalyzes (reduction of HMG-CoA to mevalonate) is generally regarded as the principal (but not unique) controlled step in the sequence of reactions leading to cholesterol synthesis, as shown in Scheme I. +

OH

0

I

II

OH

I

CH3- C
CH3- C - - C H 2 < H z 4 H

I CH2

I

I I

CH2

+ 2 NADPH + 2 H+>-

C 4 -

C 4 -

II

II

0

+ 2 NADP+ + CoASH

0

HMG-CoA

Mevalonate SCHEMEI

The interesting properties of HMG-CoA reductase as well as its considerable physiological significance have been the subject of more than a few reviews (110). The intent of this chapter is to emphasize findings that pertain principally to control of reductase through reversible phosphorylation. However, some accounting of other modalities of regulation are presented since all mechanisms undoubtedly are integrated in determining the expressed catalytic activity of the enzyme.

II. Topology The catalytic site of this membrane-bound enzyme is exposed to the aqueous environment of the cytosolic space in which the substrate HMG-CoA is generated in several steps from acetyl-CoA and the product mevalonate is processed to the polyisoprenoid pyrophosphates. From this point on enzymes in the metabolic pathway are again bound to the endoplasmic reticulum for the final stages of polyisoprene and sterol synthesis (11).Two key cholesterol processing enzymes are also embedded in this membranous structure: acyl-CoA:cholesterol 0acyltransferase (EC 2.3.1.26) and cholesterol 7 a-hydroxylase (EC 1.14.13.7) (5, 12). The former enzyme diverts cholesterol into storage form as cholesterol ester, the latter is the initial regulated step in the production of bile acids by the liver. Thus HMG-CoA reductase appears to share the lipid environment of the

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HMG-COA REDUCTASE

181

endoplasmic reticulum with the terminal steroid-polyprenoid processing multienzyme systems as well as with the lipid intermediates and products that are formed within the lipoprotein domain. HMG-CoA reductase activity released from isolated microsomes by freezing and thawing has been purified to homogeneity in several laboratories [reviewed by Beg and Brewer (6)).The soluble form of the enzyme (M,= 50,000-54,000, designated 53K) has served as the reference system in many studies. Earlier investigators assumed that this protein was the native enzyme and that its binding to the endoplasmic reticulum was that of a “peripheral” rather than an “integral” membrane protein. With antibodies to the 53K enzyme (purified IgG) reductase in free or bound form could be titrated to give an independent estimate of the amount of enzyme in tissues (13, 14). Ness et al. (15) in 1981 made the key observation that the yield of 53K reductase through freeze-thaw solubilization is quite limited if thiol protease inhibitors are added. This indicated that the soluble reductase is in fact a proteolytic fragment of a larger native enzyme embedded in the membrane. Shortly thereafter several laboratories employing the monospecific antibody to 53K reductase identified a 90- lOOK (97K) reductase species in the endoplasmic reticulum of experimental cell lines and in rat hepatocytes (16-20). This was accomplished by solubilizing the microsomal enzyme in the presence of detergents and protease inhibitors followed by SDS electrophoresis of the specific immunoprecipitate, or by detecting HMG-CoA reductase among microsomal proteins separated by SDS electrophoresis through the immunoblot transfer technique using a labeled antibody probe. Microsomes with intact 97K reductase can be obtained from tissues and cells if care is taken to exclude lysosomes, or better to include protease inhibitors and EGTA in the homogenizing-extraction medium. Without these precautions several immunoreactive species of reductase are generated: membrane-bound, catalytically active 60-65K (62K), the familiar soluble active 53K, and a smaller 35-40K fragment that has not been characterized (16, 18, 20-22). The three lower molecular-weight forms, with common immunodeterminants, appear to comprise the cytosolic domain of the 97K-microsomal enzyme since the anti-53K antibody, as well as other membrane-impermeant probes, inhibit the activity of intact microsomal 97K reductase (23). The soluble 53K reductase can be released from microsomes by the calcium-dependent thiol protease calpain (purified from liver or heart) (20). Proteolytic mapping (Staphylococcus aureus V8 protease) of 35S-methionine-labeled 97K, 63K, 52K, and 38K reductase species yielded peptide patterns that were quite similar (24). Therefore these forms of reductase, which share immunoreactive determinants, were derived from 97K reductase in the processing of cell extracts (24). In the labeling of cell proteins with 35S-methionine followed by recovery of microsomal reductase under conditions that preclude proteolysis only the 97K reductase is found in any

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DAVID M . GIBSON AND REX A . PARKER

significant quantity (16, 18, 25). While the 62K and 53K forms do not contribute to expressed reductase activity in vivo these reductase species may nevertheless be the first intermediates in the degradative route. Their rate of formation by single-clip proteolysis of the native 97K enzyme may be the limiting step in the degradative pathway . Compactin, a fungal metabolite, binds and severely inhibits HMG-CoA reductase (26). The cellular response is increased synthesis of the reductase protein (27, 28). Culturing cells in the presence of ever higher concentrations of compactin has permitted the selection of clones that produce 100-1000 times the normal level of the enzyme (16, 21, 29). These cells exhibit an extensive proliferation of the smooth endoplasmic reticulum (21). The underlying difference is attributed to reductase gene amplification with attending increases in reductase mRNA (19, 24, 30). With UT-1 cells (21) and C-100 cells (16) 97K reductase levels are greatly elevated. Employing in vitro translation systems with whole cell mRNA (24) or isolated reductase mRNA (18, 19) only the 97K species was identified. With the in vitro translation system Brown and Simoni (31) have shown that the 97K reductase remains with the cytosolic phase of the reticulocyte lysate unless microsomes are added thereby allowing the cotranslational insertion of the nascent reductase protein into the ER membrane. In a wheat germ translation system these authors (31) also demonstrated that the signal recognition particle (32) is required for cotranslational integration of the nascent chain. Since the in vitro translation product and the enzyme recovered from intact cells are virtually identical there does not appear to be a cleavage of an NH,-terminal signal peptide. This was proved by determining the specific positions of methionine and leucine in the NH,-terminal region; that is, the distributions were identical both in the in vitro and in vivo synthesized enzymes (31, 33). Another piece of evidence suggested that the 97K reductase protein penetrated the ER membrane to its lumenal surface. In C-100 cells radiolabeled mannose (31) was incorporated into the 97K protein (but none into the 62K fragment). The mannose was removed with endo-P-D-N-acetylglucosaminidase H, which is diagnostic of the high-mannose oligosaccharides of glycoproteins. Of interest in this regard is the observation of Volpe and Goldberg (34) showing that tunicamycin treatment of C-6 glial cells (which blocks glycoprotein synthesis) diminished reductase activity. Liscum et al. (20) noted that detergent-solubilized 97K reductase from UT-1 cells as well as from rat liver was adsorbed to concanavalin ASepharose, which was diagnostic of polysaccharide binding. UT- 1 cells incorporated labeled glucosamine into the 97K enzyme which could be removed by hydrolysis as before with the specific glucosaminidase. The hydrolytic products contained 6-8 mannose residues per N-acetylglucosamine (i .e., typical of asparagine-linked oligosaccharides prior to Golgi processing). Treatment of [3H]glucosamine-prelabeled97K reductase with calpain (Ca2 activated protease) yielded the soluble 53K form of the enzyme and a family of +

183

7. HMG-CoA REDUCTASE

smaller membrane-bound fragments (30-3510 (20). The 53K enzyme did not contain radioactivity (nor did it bind to concanavalin A). The 30-35K fragments which do not react with antisera to 53K enzymes were labeled. Taken together the data of Liscum er al. (20)and Hardeman et af. (24)indicate that a segment of the native 97K reductase protrudes into the lumen of the ER, the site of glycosylation, whereas a substantial portion extends into the cytosolic space where the immunoreactive determinants and the catalytic site are found. Cleavage of microsomal reductase with calpain yields the active 53K soluble fragment (20, 22) as well as the active, membrane-bound 62K fragment, which may be a precursor of the 53K species (35).These topological features of 97K reductase are diagrammed in Fig. l .

+H~N

(

(

8 220

CYTOSOL

192

I J -

ER

I

C

LUMEN

vuTJ

cdo887

62K

53K

FIG. 1 . Model representing the topology of 97K HMG-CoA reductase. Seven hydrophobic regions (filled rectangles) span the ER membrane and two relatively hydrophobic regions are also found in the extended cytosolic domain. An N-linked oligosaccharide is implanted in the lumen of the ER at residue 281. The site of proteolytic cleavage of the loosely bound 62K reductase species probably lies between residues 368 and 379, whereas the cut yielding the soluble 53K reductase is between residues 450 and 470 (36). [Redrawn after Liscum er al. (36)].

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DAVID M . GIBSON AND REX A . PARKER

Chin et af. (33) in a memorable tour de force have determined the complete amino acid sequence of the native 97K microsomal reductase of UT- 1 cells even though the native enzyme has never been isolated in pure form. Five overlapping cDNAs (4.8 kilobases in length) which were isolated through recombinant DNA methodology span the entire coding region of UT- I mRNA as well as portions of the 3'- and 5'-flanking regions. Both strands of the isolated cDNAs were sequenced by two separate methods from which the primary amino acid sequence was deduced. Altogether 887 amino acid residues were identified beginning with methionine at the amino terminal end. A molecular weight of 97,092 was calculated which agrees with estimates by SDS-gel electrophoresis (20). Of interest was the localization of three plausible sites for asparagine-linked glycosylation; only one of these is depicted in Fig. 1 pointing toward the ER lumen, since the other two would be in the segment projecting into the cytosolic space. The elaboration of the amino acid sequence permitted a glimpse of the secondary structure of the enzyme (36). With the scanning procedure of Kyte and Doolittle (37) seven relatively hydrophobic regions were identified in the aminoterminal third of the 97K enzyme. Each segment was at least 25 amino acid residues in length which would be sufficient to span a 40 A membrane bilayer. Looking back at the structure of the reductase gene (33, 36) each of these hydrophobic regions was related to a specific exon with a unique nucleotide sequence (separated by single introns). It was pointed out that the structure of 97K reductase is similar to that of retinal rhodopsin which spans the disc membrane seven times. Of further interest is the observation that rhodopsin covalently binds the polyisoprenoid 1 I-cis-retinal and that rhodopsin may be phosphorylated near its carboxy terminus (38) (see following sections). With the use of synthetic peptides that corresponded to certain regions of the carboxy-terminal two-thirds of the 97K structure antibodies were raised to probe the 62K and 53K fragments of the native enzyme (36). The 13-amino acid peptide corresponding to the carboxy-terminal end (residues 874-887) of 97K reductase gave rise to antibodies that interacted in an immunoblot analysis with both the 97K enzyme and the 53K soluble fragment indicating that these were coextensive at the carboxy terminus. In UT-I cells incubated under hypotonic conditions the 63K reductase fragment was released by endogenous proteolysis (20, 22). Antibodies to a synthetic peptide corresponding to residues 354-368 reacted with 97K reductase and with the membrane-bound 35K residue, but not with 62K reductase, indicating that the point of cleavage was downstream from this region. Antibodies to a 379-393 segment, however, did bind with the 62K enzyme. Within the span of 368 to 379 several likely sites for proteolytic attack were identified that could generate the 62K reductase species. Although incubation of UT-1 microsomes with calpain gave rise to the 53K soluble reductase, the 53K enzyme reacted with neither the 354-368 nor the 379-393 antibodies. The point of scission of the 53K reductase was less well defined but the data present-

7. HMG-COA REDUCTASE

185

ed (36) indicated that the 62K fragment is probably a precursor of the 53K reductase when microsomes are incubated with calpain (35,36). Present information on the topology of the native HMG-CoA reductase enzyme places it among the intrinsic glycoproteins of the endoplasmic reticulum, securely anchored to the membrane with multiple hydrophobic regions attached to a catalytically active hydrophilic tail extending into the cytoplasm. This particular deployment probably permits the observed multivectorial regulation of HMG-CoA reductase and of cholesterol synthesis.

111. Multivalent Control A. OVERVIEW OF CHOLESTEROL HOMEOSTASIS Polyisoprenoid lipids are found in membranes of the most primitive forms of present-day living cells, the Archaebacteria (39, 4 0 ) . Time- and temperaturemodified derivatives of primordial polyisoprene substances are detected in the depths of geologic sediments ( 4 1 ) . These were the antecedents of the steroids whose synthesis had to await the appearance of oxygen in the atmosphere ( 4 2 ) . Eucaryotic cells with a profusion of internal membranes evolved in this era. Thus catalytic reduction of hydroxymethyl glutarate to mevalonate would seem to be an ancient requirement of all living systems for generating classes of lipids that are necessary for proper membrane structure and function. Control of polyisoprenoid and sterol biosynthesis in the first instance must have been geared to membrane formation attending cell replication. With few exceptions (43) cholesterol is imperative for the continuity of eucaryotic cells in culture or within compact tissues of organisms. Other than saying that cholesterol in higher animals lends a certain rigidity to membranes at physiological temperatures (above 25-30') it is not known why the need is so severe. Moreover, the amount and distribution of free cholesterol in cells ( 4 4 ) appears to be regulated within tight limits in keeping with its role in defining the physical state of the several kinds of membranes. As with other critical metabolic components homeostatic mechanisms have evolved to maintain a steady-state level of free cholesterol in cells. A set point must exist that defines the narrow limits of concentration of free cholesterol (and of polyisoprenoids). The parameters that determine the steady-state level of cholesterol in cells, and must therefore be controlled, are influx of cholesterol, efflux (secretion), net formation of membranes (cell proliferation), and de nuvu biosynthesis of cholesterol. These are diagrammed in Fig. 2. Cholesterol is not metabolized in the usual sense, rather it is converted to other sterols, depending on the cell type, and these are usually secretory products. Membrane replacement synthesis (turnover) although dependent upon cholesterol availability would not deplete cellular cholesterol.

186

DAVID M. GIBSON AND REX A. PARKER MEMBRANE TURNOVER SECRETION

EXOGENOUS CHOLESTEROL

CHOLESTEROL

PROLIFERATlON r(r

I

I

/ 1

FIG.2. Diagram of principal determinants of cholesterol homeostasis in cells. Boxes numbered 1-3 represent sterol and nonsterol feedback signal molecules that are metabolic products of mevalonate. Number 4 is also an isoprene derivative that serves as a feedforward signal permitting cell replication.

Beyond the perspective of a single cell is the organism that must protect and sustain its constituent cell populations through global homeostatic control of cholesterol flux (45). Here the liver plays a central role in receiving dietary cholesterol via plasma chylomicron remnants; packaging this exogenous cholesterol and newly synthesized cholesterol as a secretory product for export to all cells in the form of plasma very-low-density lipoproteins (VLDL); receiving cholesterol back from the tissues as plasma intermediate-density lipoproteins (IDL) and low-density lipoproteins (LDL); and excreting cholesterol and bile acids to the outside as the terminal efflux of this remarkably regulated physiologic process. Since most cells in culture have metazoan origins they are equipped to receive plasma lipoproteins (LDL) through receptor-mediated endocytosis as the influx route of cholesterol, and to bind acceptor proteins, for example, high-density lipoproteins (HDL) for removal of free cholesterol from the plasma membrane. Keeping in mind the parameters in Fig. 2 one must ask how the cell can “sense” the current free cholesterol level and, if out of line, effect a change in the parametric determinants. The problem is further complicated by the fact that the distribution of cholesterol is heterogeneous among membranes in the cell (highest concentration in plasma membranes, least in the endoplasmic reticulum), among the many microdomains within any membrane, and in mo-

7. HMG-COA REDUCTASE

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lecular form (free, predominantly in membranes, or esterified to fatty acids as a depot lipid). It is not obvious how membrane-cholesterol, per se, could function as a signal molecule to a distant effector system. This cholesterol, however, can be mobilized by binding to one or more kinds of sterol carrier proteins in the cytosol (46, 47). Also minute vesicles bleb off the surface of membranes and fuse with other membranes in an orderly vectorial fashion [e.g., passage of membrane components including cholesterol from the cis to the trans sides of the Golgi layers (48)].Cholesterol movements via the sterol carrier proteins or microvesicles, possibly directed by contractile elements in the cytoskeleton, could reach sites that are important in controlling cholesterol homeostasis. At least this conveyance of cholesterol, as a signal molecule, would provide direct information on the state of those membranes from which it originated. A more subtle, indirect signaling system can be imagined that would depend on products of membrane-bound enzymes or transport proteins whose expressed activity depended on a characteristic of membrane structure which in turn was influenced by cholesterol content in the immediate vicinity of the enzymes (49, 50). Molecules related metabolically to cholesterol have also emerged as important candidates in controlling cholesterol homeostasis (51). Various hydroxy derivatives of cholesterol (and related sterols), some of which occur naturally, are potent inhibitors of cholesterol formation (7) as are nonsteroid metabolic products of mevalonate (52). Whatever mechanism is imagined for cholesterol homeostasis in cells it must affect one or more of the vectors depicted in Fig. 2. If secretion of cholesterol and/or cell proliferation increase the need for cholesterol then one or both of the ‘‘input” vectors (exogenous cholesterol and cholesterol synthesis) must be enhanced to keep the cellular cholesterol level on an even keel. This essential logic is observed. An additional parameter of importance (not shown) in liver and other cells that synthesize steroids for export is the reversible formation of cholesterol esters that function as an internal free cholesterol supply buffer (45, 53). As free cholesterol levels rise in these cells the enzyme acyl-CoA cholesterol acyltransferase is turned on to divert free cholesterol into storage form. With elevated levels of free cholesterol the flux of cholesterol into the cell is diminished by reducing the number of receptors for the cholesterol-rich plasma lipoprotein, LDL, by repressing the synthesis of a specific surface glycoprotein receptor (45, 53). Elevation of free cholesterol at the same time diminishes the expressed activity of HMG-CoA reductase (see Section 111,B). These three coordinate events (45, 53) act to bring the cholesterol level again to the normal set point. A drop in cholesterol levels would have the opposite effect on the three key determinants. Secretion of cholesterol by many cells depends on extracellular (plasma) acceptor proteins, especially HDL, or (experimentally) liposomes. In liver the limiting enzyme in bile acid production, cholesterol 7ahydroxylase, is stimulated by the concentration of its substrate in the endo-

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DAVID M . GIBSON AND REX A . PARKER

plasmic reticulum. Ingested anion exchange resins, e.g. cholestyramine, avidly bind bile acids in the intestine and prevent their reabsorption. Thus, cholesterol 7a-hydroxylase (which is dampened by bile acids) is stimulated further and diminishes the cholesterol level in liver. Cholesterol synthesis is enhanced as is the uptake of cholesterol via endocytosis of plasma lipoprotein (45, 53). Cell replication depends on new membrane formation. In proliferating cell cultures cholesterol biosynthesis is stimulated. Indeed a tight coupling of cell replication and the provision of cholesterol exists. The rate of sterol synthesis diminishes in cultures with high cell density (54-56). Endocrines may profoundly influence cholesterol homeostasis. Mitogenic growth factors signal new cholesterol synthesis. Certain cells (e.g., adrenal, ovary, and testis) that produce steroid hormones for export respond to cellspecific endocrine signals with increased cholesterogenesis, and release of free cholesterol from cholesterol ester deposits. Cholesterol traffic in liver parenchymal cells (hepatocytes) is markedly affected by diurnal variations in food comsumption. Eating is attended by release of insulin into the portal circulation which stimulates enzyme systems in liver that convert glucose and amino acid carbon into lipids, primarily triglycerides but also cholesterol. In the liver these are packaged into very-low-density lipoproteins (VLDL) for export. Cholesterol formation apparently must keep pace with VLDL production (57). If the cholesterol content of the diet is high the chylomicron remnants taken up by the liver are enriched in cholesterol and biosynthesis is depressed (50, 58). Progression toward relative starvation with diminishing levels of circulating insulin and elevated glucagon leads to a fall in the rate of triglyceride and cholesterol synthesis. Thus, a biphasic diurnal cycle of cholesterol formation is observed, especially in experimental animals subjected to a strict meal-feeding schedule ( 1 , 4).

B. REGULATION Since the expressed activity of HMG-CoA reductase is the principal determinant of the rate of cholesterol biosynthesis it is not unexpected that the enzyme protein has evolved as a sensitive regulatory site in cholesterol homeostasis (Fig. 2). However indirectly, this enzyme must sense and respond to the cholesterol level and deployment in the cell and, in more general terms, to the need for net membrane formation. Membrane turnover as a steady-state operation necessitates that HMG-CoA reductase is itself continuously tuned or coupled to the status of membrane structure and function through the information provided by cholesterol or cholesterol-related signal molecules. “Cholesterol negative feedback control” of reductase activity carries this meaning although a variety of metabolic intermediates, essentially in the cholesterol-polyisoprenoid biosynthetic pathway, undoubtedly participate as well. The expressed activity of an enzyme subject to control is the summation of two variables: ( a ) the quantity of

7.

I89

HMG-COA REDUCTASE TABLE I DETERMINANTS OF THE QUANTITY A N D EXPRESSION OF HMG-COA REDUCTASE AND ITS BlOSYNTHETlC AND DEGRADATIVE SYSTEMS Component

Amount

Gene

Species-cell characteristic Amplification Mutation

mRNA

Transcription rate and RNA processing Degradation rate State of mRNA-protein complex State of RNA degradative system

HMG-CoA reductase

Translation rate and posttranslation processing Degradation rate State of reductase protein State of protein degradative system Membrane translocation

Expression Transcription rate Cell-specific control of chromatin proteins Feedback signaling Endocrine signaling Translation rate mRNA release and activation tRNA-ribosomal system State of RER membrane Feedback signaling Endocrine signaling Catalytic rate ER-membrane microenvironment Covalent modulation system Effectors Feedback signaling Endocrine signaling

enzyme protein (i.e., the number of enzyme molecules as a gene product), which depends on the balance between its rate of synthesis and degradation; and (b)the modulation of existing enzyme by agencies that affect its spatial conformation or otherwise influence its catalytic properties. Among these agencies are effectors that reversibly bind noncovalently to the enzyme, and satellite enzymes that catalyze the addition and removal of chemical groups to amino acid side chains through covalent bonds (e.g., reversible phosphorylation). Table I summarizes the several mechanisms that could affect the amount and expression of HMGCoA reductase. It is beyond the scope of this review to discuss the impact of possible combinations of these interdependent systems except to say that cholesterol feedback signals may exert an influence at all levels. For example, it has been suggested that cholesterol (or hydroxycholesterol) may act as a repressor of HMG-CoA reductase synthesis (59-61), or may inhibit the activity of reductase directly in the endoplasmic reticulum by diminishing the fluidity of the lipoprotein structure in the immediate environment of the enzyme (50, 62). One could also imagine that the cholesterol content of the endoplasmic reticulum might affect the insertion of the nascent HMG-CoA reductase protein into the membrane. In any event the cholesterol feedback components that determine cholesterol homeostasis in cells achieve this objective through regulation of both the amount and the expressed activity of HMG-CoA reductase.

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DAVID M . GIBSON AND REX A . PARKER

1. Quantity In the studies that are briefly reviewed here a number of different approaches have been employed in exploring the mechanisms by which the quantity of HMG-CoA reductase is controlled: rate of incorporation of a radiolabeled amino acid (e.g., [35S]methionine, into reductase recovered by specific immunoprecipitation and separated on SDS electrophoresis); measurement of mass of reductase by immunoblot transfer after SDS electrophoresis; estimation of reductase mRNA concentration by rate of in vitro translation (labeled reductase recovered by immunoprecipitation and electrophoresis); quantitation of reductase mRNA with specific cDNA; measurement of reductase degradation in pulse-chase experiments as well as following loss of mass by specific immunoblot recovery. a. Receptor-Mediated Endocytosis. Uptake of cholesterol-rich LDL by cells in culture by receptor-mediated endocytosis dampens HMG-CoA reductase (45). Cells remain viable since cholesterol is provided (formation of polyisoprenes such as dolichol and ubiquinone is not completely blocked). Synthesis of HMGCoA reductase is diminished and degradation is increased (22). In UT-1 cells, which grow in the presence of compactin, addition of LDL to the culture medium diminishes the level of mRNA (18) and causes a regression of the extensive proliferation of the endoplasmic reticulum (63). Uptake of LDL in normal fibroblasts increases the level of acyl-CoA:cholesterol acyltransferase, whereas the quantity of reductase and LDL receptors falls (45, 64) indicating a coordinate regulatory mechanism in the synthesis and degradation of these three proteins. Fibroblasts with genetically deficient LDL receptors exhibit elevated reductase activity (45). Removal of cholesterol from the plasma membranes of cells with HDL results in increased synthesis of reductase and decreased degradation (65, 66). Also see (201, 202). b. Mevulonak. Addition of mevalonate to cell cultures has much the same action as LDL. With amounts of mevalonate that greatly exceed its normal intracellular concentration cholesterol synthesis is increased. Reductase synthesis is impaired and degradation enhanced (17, 67) (See Section III,B,l ,f.) c. Hydroxysterols. A wide variety of hydroxysterols ( ‘‘oxysterols”) have been prepared some of which appear in small amounts normally as intermediates in cholesterol biosynthesis (7, 60). The best-studied, 25-hydroxycholestero1, brings about a decline in reductase synthesis and an increase in degradation of the enzyme (22, 68). Proliferation of cells in culture is impaired since formation of cholesterol and other mevalonate products are blocked (69). Depending on the cell type 25-hydroxycholestero1 interrupts translation [no change in reductase (mRNA)] (68) or transcription (no effect in enucleated cells) (70). The increase in the degradative rate, interestingly, is diminished in the presence of protein

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synthesis inhibitors (71, 72). In another study (73) an inhibitor of 2,3-oxidosqualene cyclase (U 18666A) causes the diversion of the normal lanosterol precursor into squalene 2,3:22,23-dioxide. Removal of the inhibitor permits the formation of “polar sterols” (probably including 24,25-0xidolanosterol), that is, ‘‘naturally occurring” oxysterols that diminish HMG-CoA reductase activity (73). Incubation of cells with squalene dioxide does in fact inhibit reductase activity (73). d. Compactin and Mevinolin. The fungal organic products compactin (26) and mevinolin (74) are potent direct competitive inhibitors of HMG-CoA reductase and thus block the production of mevalonate (and cholesterol precursors). This causes induction of reductase synthesis and impairs degradation of the enzyme protein. Both effects are reversed with added cholesterol or mevalonate (24, 45, 75). Compactin also blocks cell division (76-78). Depending on the concentration of compactin, the cell type, and stage in the cell cycle, mevalonate (but not cholesterol by itself) can rescue cell growth (76-78). In one cell type isopentenyl adenine can replace mevalonate (79, 80). This compound, or a metabolic derivative, is thus cast in the role of a positive feed-forward signal permitting the cell cycle to proceed into the S-phase (DNA replication). Human fibroblasts incorporate mevalonate into A2-isopentenyl adenine nucleotide, which is a minor base component situated near the anticodon of several tRNA species (81). Also a A2-isopentenyl pyrophosphate:5’-AMP A2-isopentenyltransferase has been identified in slime mold (82). Thus, mevalonate is the precursor of three classes of molecules: 1. Products that are needed for cell continuity (cholesterol in membrane formation, ubiquinone for electron transport, dolichol for glycoprotein synthesis, isopentenyl-tRNAs in protein synthesis) 2. Metabolic intermediates, both sterols and nonsterols, that provide negative feedback signals that control their rate of formation by limiting the flux through HMG-CoA reductase 3. A feed-forward, “all-is-ready” signal that permits the cell cycle to proceed The negative feedback signal is generated by rising levels of mevalonate products, whereas the feed-forward signal appears to be routinely produced at concentrations of mevalonate above a very low threshold. e. Cell Lines and Cloning. Several cell lines have been grown in the presence of compactin, and surviving cells cloned to face still higher levels (21, 29). In the final extremity are cells resistant to added compactin with enormously elevated levels of HMG-CoA reductase due to selection of cells with reductase gene amplification (21, 30). The endoplasmic reticulum within these cells undergoes a remarkable proliferation (“cystalloid” ER) (21, 63). With addition of

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LDL the cells revert to more normal reductase levels and membrane morphology (63).These findings recall the close correlation that exists between the generation of membranes in cell proliferation and cholesterol synthesis (HMG-CoA reductase activity) (54, 65, 77, 78, 83, 84). (See Section 111,A). f. Derivatives of Mevalonate. Derivatives of mevalonate block certain steps in the polyisoprene biosynthetic pathway. The best-studied of these, 3-fluoromevalonate, is metabolized to fluoromevalonate pyrophosphate which severely inhibits the conversion of mevalonate pyrophosphate to isopentenyl pyrophosphate. Overall, this block has the same outcome as compactin inhibition. Of special interest is the employment of this agent in cells from Drosophila embryos which do not synthesize nor require cholesterol for growth (43, 52, 85). Nevertheless, fluoromevalonate brings about a stimulation of reductase synthesis and appears to diminish its degradation. In the presence of fluoromevalonate, mevalonate pyrophosphate accumulates suggesting that a nonsteroid product of mevalonate beyond this point is ordinarily a feedback signal (52).Aside from the net effect of decreasing synthesis and enhancing degradation of reductase the mechanism of action of the mevalonate-derived signal is not known. In another cell line (Swiss 3T3 cells) labeled mevalonate, in addition to the metabolic fates noted in Section III,B,l ,d, is incorporated covalently into a set of proteins (86). Neither the proteins nor the nature of the covalently attached mevalonate product have been identified. It is thus possible that one or more enzymes that are involved in the feedback (or feedforward) system are modulated by a mevalonate metabolic derivative. Still other cellular products that originate from mevalonate are candidates for feedback repression of HMG-CoA reductase-the plant hormone abscisic acid (87) and the cyclic monoterpenes cineole and menthol (88). g. Diurnal Variation. In intact animals the diurnal rise in HMG-CoA reductase activity during feeding ( I ) is attributed principally to an increased rate of synthesis of reductase [elevated reductase (mRNA)] (25)and a diminished rate of degradation of the enzyme. This pattern is reversed as starvation ensues. Dietary supplementation with mevinolin or cholestyramine, especially if these two compounds are administered together, greatly enhances the rate of synthesis of reductase [elevated reductase (mRNA)] and decreases its rate of degradation (89-92). Examination of rat liver by electron microscopy in the latter situation reveals abundant crystalloid ER (93). Even with these elevated rates of reductase synthesis the general pattern of diurnal variation is not lost (25, 89-91) which suggests that the impact of changes in the insulin-glucagon ratio is not lost (see Section V). Dietary supplementation with cholesterol on top of mevinolin and cholestyramine also exerts an effect-diminished reductase synthesis [depressed reductase (mRNA)] and an enhancement of reductase degradation (19). It is clear from the preceding observations that control of the amount of HMG-

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CoA reductase is mediated by mevalonate product inhibition of the rate of synthesis of reductase coupled to stimulation of its rate of degradation. However, neither the mechanism of action at a molecular level nor identification of the signals is certain at this stage. In Fig. 2 four categories of mevalonate products are numbered that may function as intracellular signals. 1. The most proximal branch point is isopentenyl pyrophosphate which is the precursor of 2, 3, and 4.(Its role as a feedback inhibitor is a possibility but not established.) 2. The sterol products include oxysterols and cholesterol. 3. Nonsterol polyprenyl intermediates which are or are not on the cholesterol biosynthetic pathway would include farnesyl pyrophosphate. 4. In contrast to the three groups above are mevalonate products that comprise a feedforward signal permitting cell replication to continue (e.g., isopentenyl adenine). In routine steady-state operation of the cell the mevalonate products provide essential structural and functional molecules. Like cholesterol their homeostasis is geared to HMG-CoA reductase activity. Only by interrupting the flow of mevalonate experimentally (e.g., with compactin) do we appreciate the extent to which cellular function and viability depend on the synthetic sequelae of mevalonate, a recent example being the maintenance of fatty acid and protein synthesis (94).

h. Endocrine Signaling. Superimposed on the steady-state operation of the feedback system is the influence of endocrines that affect both the amount and (as seen later in Section V) the expressed activity of HMG-CoA reductase in cells. Hormones insure that constituent cells subserve the organism through cell proliferation (growth factors, mitogens) and specific cell functions. Cell proliferation with attending net synthesis of membranes places a demand on cholesterol synthesis that is met through the induction of HMG-CoA reductase (54, 65, 77, 78, 83, 84). On the other hand, signaled cell function that creates a need for cholesterol appears to be linked to the production of cholesterol and cholesterol derivatives for export (e.g., secretion of steroid hormones). In this sense the amount (and expressed activity) of liver HMG-CoA reductase that responds to the circulating insulin-glucagon ratio (the diurnal cycle) (4) fulfills the requirement for cholesterol in VLDL formation (57).

2 . Expressed Activity Modulation of existing HMG-CoA reductase in cells is as important a determinant of the rate of mevalonate production as control of reductase synthesis and degradation (Table I).

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a. Microenvironment in the Endoplasmic Reticulum. Our knowledge of the deployment of reductase in the ER membrane (Fig. 1) allows an appreciation of how adjacent lipids and proteins may affect the activity of the enzyme. An increase in the cholesterol content of membranes decreases the fluidity and tends to displace proteins toward the surface of the membrane (95).For example, the carboxy terminal domain of the native reductase is projected further into the cytosolic space so that it is more exposed to proteases and other enzymes that modulate reductase activity. Pertinent to this argument is the observation of Orci et al. (63) that the increase in degradation of reductase in UT-1 cells linked to endocytosis of LDL is correlated with a selective acquisition of cholesterol by the endoplasmic reticulum (as judged by the cholesterol-binding compound, filipin, visualized by electron microscopy). These authors suggest that cholesterol may signal the degradation of reductase as well as the concomitant dissolution of the UT-1 crystalloid ER. The laboratories of Mitropoulos (50)and Sabine (62) have provided evidence that reductase activity in the liver microsomal fraction from rats fed a diet supplemented with cholesterol was not only diminished but also varied in activity as a function of incubation temperature in a manner quite distinct from rats fed a control diet. In the latter case the Arrhenius plot (log reductase activity versus 1/T) showed a sharp break in the slope at 29", whereas in the cholesterol series the slope was linear. The experimental pattern was attributed to an increase in the concentration of cholesterol in the vicinity of the enzyme. Similar high-cholesterol Arrhenius plots were observed in liver microsomes of rats that received intravenous injections of mevalonate or were starved (50). Pretreating microsomes in vitro with serum (containing lipoproteins) produced the same linear relationship of reductase activity to temperature compared to microsomes exposed to lipid-depleted serum. These results were correlated with the measured cholesterol content of the membranes (50). b. Modulating Proteins and Effectors. A number of liver cytosolic proteins have been examined that influence the activity of HMG-CoA reductase (96). Aside from sterol and lipid carrier proteins their role in regulation is not certain. An interesting iron-containing protein has emerged that inhibits reductase activity (97). Disulfides such as oxidized glutathione inactivate reductase and organic thiols reactivate (e.g., dithiothreitol, which is routinely added in the preparation of the enzyme) (98).Indeed, a high-mass, inactive species of reductase has been claimed which is converted to the active 97K form under reducing conditions (99). c. Covalent Modulation. The addition of the gamma phosphate of ATP to amino acid residues of enzymes (serine, threonine, or tyrosine) through the action of specific protein kinases is now a universally observed mechanism for modulating enzyme activity, as testified by this volume. Evidence reviewed in

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Sections IV and V supports the view that reversible phosphorylation not only affects the activity of existing HMG-CoA reductase but also may in part determine the quantity of the enzyme.

IV. Reversible Phosphorylation in Vitro A.

EARLYSTUDIES

Beg et al. in 1973 (100) reported that rat liver microsomal reductase activity was diminished following incubation with MgATP and protein fractions from the liver cytosol. The loss in activity was not affected by repeated centrifugation of the microsomes in buffer. However, when the pretreated microsomes were again incubated with liver cytosolic fractions in the absence of MgATP, HMG-CoA reductase activity was restored. It was suggested that reversible phosphorylation of reductase was the underlying mechanism. Higgins and Rudney (13) in the same year presented evidence that rat liver HMG-CoA reductase responded to dietary manipulation by variation in mass as well as in degree of activation. Likewise Goodwin and Margolis (101) found that preincubation of postmitochondrial liver homogenates greatly enhanced subsequent acetate (but not mevalonate) incorporation into cholesterol. Related studies were reported over the next few years confirming the existence of the ATP-dependent modulation of microsomal reductase (102-105). Nordstrom et al. in 1977 (106) reexamined this system with purified modulating enzymes. They found that the activator enzyme was severely inhibited by fluoride and that merely isolating microsomes from liver in the absence of fluoride would bring about an activation of reductase. The inactivating system diminished the V,,, of reductase with no effect on the K , values for either substrate (104, 106). Both microsomal reductase and the soluble 53K form responded to MgATP and the inactivating enzyme (106). Finally in 1978 Ingebritsen et al. (107, 108) confirmed that reversible phosphorylation must be involved by showing that the activation system could be replaced with fluoridesensitive phosphorylase a phosphatase from liver (109). The inactivating enzyme, now termed reductase kinase, was not affected by CAMP nor the CAMPprotein kinase inhibitor (107, 108, 110) although the preparation of Nordstrom et al. (106) required ADP as well as MgATP. Further studies with reductase kinase showed that it was inactivated with the protein phosphatase (i.e., it was subject to reversible phosphorylation but in the opposite sense to reductase). A second protein kinase, reductase kinase kinase requiring ATP and Mg2 was identified, which restored activity to reductase kinase. Reductase kinase kinase was also CAMP-independent. Consequently it was realized that HMG-CoA reductase was controlled through a bicyclic system +

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RE DUCTAS E KINASE KINASE

PHOSPHATASE

REDUCTASE

PHOSPHATASE

Fic. 3. The bicyclic system for modulation of HMG-CoA reductase through reversible phosphorylation. The symbols (a) and (b) denote active and inactive enzyme species, respectively.

consisting of two protein kinases and a phosphatase (Fig. 3) (107. 108, 110). This was confirmed by Beg et al. both in rat (111) and human (112) liver. Modulation of HMG-CoA reductase through reversible phosphorylation has been demonstrated in tissues of many species and in isolated cell types [reviewed in Refs. (9, l o ) ] .Arguments that this system does not exist have been addressed and eloquently answered by Kennelly and Rodwell (9).

B . PHOSPHORYLATION For definitive proof of phosphorylation of enzymes one must demonstrate that the gamma phosphate of ATP is transferred to covalent linkage with protein residues and that a change in catalytic capacity is commensurate with the degree of phosphorylation. Three laboratories have provided this evidence. Bove and Hegardt in 1978 (113) incubated liver microsomes with [ Y - ~ ~ P ] M ~ ATP with the result that reductase activity was depressed while microsomal proteins accepted the 32P label. Treatment with protein phosphatase released the radioisotope at the same rate at which reductase activity was restored. This

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interesting result while not proving that reductase per se was phosphorylated pointed to a possible coordinate phosphorylation among microsomal proteins which deserves further study. Beg et al. (114) the same year reported a similar experiment but took pains to separate reductase from freeze-thaw extracts with specific antisera (to the homogeneous 53K enzyme). The data presented showed a direct correlation between the amount of label incorporated into the immunoprecipitate and the degree of inactivation of reductase. Keith et al. (115)and Gil er al. (116) similarly treated microsomes with reductase kinase and [Y-~~PIM~A and T Pprepared homogeneous 53K reductase from the extract. The label comigrated with the reductase band on SDS electrophoresis and with reductase activity on electrophoresis under nondenaturing conditions. In the preceding experiments an inverse correlation of microsomal HMG-CoA reductase activity and the phosphorylation state was implicit but in fact not quantified. In a recent examination of this relationship both Beg et al. (117) and Ferrer and Hegardt (118) with homogeneous reductase kinase and soluble 53K reductase were able to show that the inactivation of reductase was a function of the degree of incorporation of 32P from [ Y - ~ ~ P ] A TThe P . other side of the reversible phosphorylation cycle was established by Gil et al. (119) with 32Plabeled homogeneous reductase in the presence of purified protein phosphatase. The increase in activity was concomitant with release of inorganic 32P. Related experiments in these three laboratories corroborated that the degree of phosphorylation of HMG-CoA reductase dictated its expressed catalytic activity (1 14, 116, 119-122). Localization of the phosphorylation site(s) has not been resolved. These presumably are somewhere in the cytosolic domain of the 97K reductase (Fig. 1) which has access to the modulating enzymes (see Section V1,C). The soluble 53K fragment is the only portion of the 97K native enzyme that has been examined following treatment of microsomes with labeled ATP; and the isolated 53K enzyme is in fact modulated directly by reductase kinase and protein phosphatase. These observations do not preclude, however, the possibility that phosphorylation sites exist elsewhere in the 97K structure. Keith et al. in 1983 isolated [32P]phosphoserinefrom hydrolysates of the homogeneous phosphorylated 53K reductase (123),a finding that establishes the covalent characteristic of the protein-phosphate bond. Analysis of tryptic peptides derived from the 32P-53K enzyme suggests that there may be at least two structurally distinct phosphorylation sites per mole of the 53K species (123, 124). It is of interest that treatment of labeled reductase with purified protein phosphatase engenders full reactivation but with release of only half of the bound phosphate. The remaining phosphate is hydrolyzed without affecting reductase activity (120). Now that the amino acid sequence of 97K HMG-CoA reductase is known (33) the many serines of the cytosolic domain may be scrutinized individually.

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C. THE MODULATING ENZYMES Reductase kinase was initially defined as a cytosolic protein that was CAMPindependent (110).Nordstrom et al. (106) made the point that ADP is required for full reductase kinase activity in liver (and in fibroblast extracts) (104). Later purification of this enzyme from rat liver cytosol bore this out (125) whether using microsomal reductase, the soluble 53K form, or detergent-solubilized 97K reductase (126) as phosphate acceptor. With 53K reductase and cytosolic reductase kinase the K , for ATP was 140 cwl/l and the K , for ADP was 1.4 mM (125). Preparations of reductase kinase from microsomal extracts do not seem to show this unique allosteric requirement for ADP. The observation that reductase kinase is inactivated by protein phosphatases (107, 108, 110) was confirmed by Beg et al. (111) with purified enzymes. Reductase kinase purified to homogeneity from microsomal extracts possessed a monomeric molecular weight of 58,000 by SDS electrophoresis. With partially purified reductase kinase kinase the reductase kinase protein was phosphorylated with [Y-~~PIATP, and most of the label was removed with protein pohsphatase (111). Ferrer and Hegardt (118) also purified rat liver microsomal reductase kinase using different methods obtaining a monomeric molecular weight of 105,000. In this study as in other investigations (127, 128) reductase kinase was clearly separated from mevalonate kinase. Beg er al. (129) have demonstrated that two previously described protein kinases phosphorylate and inactivate both microsomal 97K HMG-CoA reductase and the 53K soluble enzyme. The phospholipid-dependent, calcium-activated (77K) protein kinase-C (130) and the calmodulin-dependent (50 K and 60K subunits) protein kinase (CaM kinase 11) (131) phosphorylate reductase at greater specific rates than reductase kinase. Maximal incorporation of 32P (about one phosphate per mole reductase) and the pattern of labeled tryptic peptides from reductase were the same for the three protein kinases. Incubation of phosphorylated reductase with purified rat liver cytosolic protein phosphatase was associated with loss of 32P and reactivation of enzyme activity. Protein phosphatases are ubiquitous in distribution, diverse in molecular form, and only rarely show absolute specificity with regard to phosphoprotein substrates (see Volume XVII, Chapter 8). Recent attempts to classify protein phosphatases indicate the complexity of this interesting group of enzymes (132). HMG-CoA reductase is no exception since almost any preparation of protein phosphatase (including bacterial phosphatases) will activate the enzyme. The question of which phosphatase acts on reductase in vivo and which may therefore be subject to control remains open. Reductase phosphatase activity originally identified in rat liver cytosol (106-108, 110) has also been detected in microsomes (133). High-molecular-weight phosphatases have been purified extensively under relatively mild conditions using 32P release from phosphorylated 53K reductase or reductase activation in order to follow the progress of purifica-

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tion: 480K and 310K (134), 190K and 120K (120) from rat liver cytosol, and 180K, 90K and 75K from microsomal extracts (122). No variation of any of these species uner physiological conditions in vivo as a means of controlling the state of phosphorylation of reductase has been observed.

V.

lntracellular Phosphorylation

A.

INTRODUCTION

Putative control systems must ultimately face the test of biological significance. With HMG-CoA reductase this must be addressed in the context of reversible phosphorylation. Thus, research has been directed in vivo to determine if the expressed activity varied, as well as total quantity of reductase protein, and if the change in expressed activity was due to the state of phosphorylation of the enzyme. Nordstrom et af. (106) provided the experimental approach by comparing the (‘‘expressed”) activity of microsomes isolated from liver in the presence of fluoride and EDTA (to block endogenous protein phosphatases and protein kinases) with microsomes separated and then treated with purified protein phosphatase to completely activate reductase before assay (“total” activity). These authors were the first to recognize that expressed activity in rat liver was only 20% of total activity (E/T ratio = 0.2). The extent of incorporation of 32P into the enzyme as an inverse function of expressed activity is also a necessary requirement to complete the picture. As with other interconvertible enzymes the time frame of modulation of expressed activity is short, minutes rather than the hours lapsed for changes in mass of enzyme protein in most situations. Briefly reviewed in the following sections is our state of knowledge concerning the intracellular regulation of HMG-CoA reductase by phosphorylation. Both endocrine and feedback control signals that are known to affect cholesterol homeostasis and reductase activity are examined.

B.

EARLYSTUDIES

Suspensions or monolayers of freshly isolated, intact liver parenchymal cells (hepatocytes) have been employed in many metabolic studies since these cells retain sensitivity to insulin and glucagon (135, 136). Since glucagon and CAMP seemed to inhibit cholesterol synthesis in several liver tissue preparations (137, 138) the effects of glucagon and insulin were tested in 1979 by Ingebritsen et af. (139) on hepatocytes incubated in a simple buffered medium with glucose. Expressed and total reductase were assayed in microsomes separated from sonicated cells after incubations extending over a 150 min period. Addition of glucagon caused a fall in both the expressed and total microsomal reductase

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DAVID M. GIBSON AND REX A. PARKER

activities while insulin enhanced expressed activity and diminished the spontaneous rate of decline of total activity. Expressed reductase kinase activity in the cytosolic fraction was also found to vary in response to these hormones but in the opposite direction to reductase. This was expected if the bicyclic cascade system were operative (Fig. 3), namely, insulin signaled the dephosphorylation of both enzymes and glucagon the phosphorylation of both (139). The rate of cholesterol synthesis in these cells, measured by tritiated water incorporation, was increased with insulin and decreased with glucagon in keeping with the concept that HMGCoA reductase is limiting. Observations in Rodwell’s laboratory (9) confirmed these hepatocyte studies. Changes in expressed reductase activity were detected as rapidly as 10 min after hormone addition. Also see (203, 204). These observations with hepatocytes place HMG-CoA reductase and reductase kinase among the endocrine-sensitive interconvertible enzymes of liver that acutely regulate the metabolic flow of carbon precursors into the synthesis of lipids (Fig. 4) (140-147). The CAMP-dependent protein kinase does not affect reductase or reductase kinase in vitro (110). Yet glucagon depresses the activity of reductase and stimulates reductase kinase in hepatocytes. Reductase phosphatase activity is also diminished after glucagon pretreatment of hepatocytes (155). It is possible that this action of glucagon is mediated through protein phosphatase inhibitor- 1, which is active after being phosphorylated by CAMP-dependent protein kinase (132). Also other enzymes that are phosphorylated via CAMP-protein kinase are competitive substrates for the protein phosphatase enzymes acting on reductase (155).

PHOSPHORYLATION CONTROLOF REVERSIBLE C. FEEDBACK Erickson et al. (148) administered mevalonic acid (as the mevalonolactone) to rats by stomach tube. This brought about a rapid fall (15 min) in microsomal reductase activity. Addition of protein phosphatase to the impaired experimental microsomes restored activity. The control cytosol had the same restorative capacity, but the experimental cytosol clearly possessed less phosphatase activity when assayed in this manner. It was concluded that HMG-CoA reductase was subject to acute inactivation by phosphorylation of the enzyme and that this tilt in the phosphorylation state was brought about by an inhibition of protein phosphatase activity. One hour after the mevalonolactone treatment reductase activity was severely depressed, but the enzyme failed to respond to added protein phosphatase. Arebalo et al. (149, 150) conducted similar experiments and found that the results of mevalonolactone and cholesterol feeding were essentially the same. Following up on the refractory state of reductase at one hour these investigators discovered that the total reductase protein estimated by immunotitration was

20 1

7. HMG-CoA REDUCTASE GLUCOSE -GLYCOGEN

FR UCT 0S E

BRANCHED CHAINAMINO-ACIDS ALPHA KETO ACIDS ACYL-CoA(S)

1.6 BIS-PHOSPHATE

PHOSPHATIDYL CHoLINE

TRIGLYCERIDE

PYRUVATE

FATTY -ACYL-CoA

FIG.4. Interconvertible lipogenic enzymes of liver. The 14 interconvertible enzymes numbered in circles in the diagram are presented in the dephosphorylated mode: increased enzyme activity (+), decreased (-). The dephosphorylated state facilitates the flow of glucose carbon into free cholesterol, fatty acids, triglyceride, and phosphatidylcholine. Evidence suggests that bile acid and cholesterol ester formation would be impaired (12). The enzymes listed are ( I ) glycogen synthase (140); (2) phosphorylase kinase (140); (3) glycogen phosphorylase (140); (4) 6-phosphofructo-2-kinaseI fructose-2,6-bisphosphatase(141); ( 5 ) pyruvate kinase (142);( 6 ) pyruvate dehydrogenase (143);(7) acetyl-CoA carboxylase (144); (8) cytidylyl transferase (145);(9) diglyceride acyltransferase (146); (10) reductase kinase; (1 1) HMG-CoA reductase; (12) acyl-CoA cholesterol acyltransferase (12); (13) cholesterol 7a-hydroxylase (12); (14) branched-chain a-ketoacid dehydrogenase (147). Enzymes 1-7 and 14 are reviewed elsewhere in this volume.

unchanged over this period (151). At later stages immunodetectable enzyme mass fell. It was concluded that mevalonate or cholesterol feeding acutely depressed reductase activity through phosphorylation following which irreversible changes take place that may represent initial stages of degradation of the enzyme (151). Immunoreactive but catalytically inert, soluble, 53K “reductase” has been identified in the course of purifying active 53K reductase from freeze-thaw extracts (152, 153). Beg and Brewer (6)reported experiments demonstrating that feeding mevalonolactone caused a 1.6-fold increase in the incorporation of 32P (from an intraperitoneal injection of 32Pi) into HMG-CoA reductase 20 min later at the same time that expressed reductase activity fell 36%. Virtually all of the 32Pin covalent

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linkage to the enzyme (purified to homogeneity) was released with protein phosphatase. Beg et al. (154) have provided evidence that mevalonolactone feeding not only depressed protein phosphatase activity but also significantly enhanced reductase kinase and reductase kinase kinase activities, changes which would certainly increase the degree of phosphorylation of reductase (Fig. 3). This is the first demonstration that reductase kinase kinase may be modulated. Incubation of hepatocyte suspensions with mevalonolactone produces the same pattem of changes in expressed and total reductase activities as seen in vivo (155, 156) (Fig. 5 ) . Frequent sampling revealed that the expressed activity falls dramatically over the first 15 min before any change in the total enzyme level is detectable. The total reductase activity then falls over the next 60 min. In similar experiments with hepatocytes (17) the rate of synthesis of reductase is decreased and the rate of degradation of the enzyme is increased in response to mevalonolactone. Pretreatment of hepatocytes for 15 min with mevalonolactone effects a fall in cytosolic reductase phosphatase activity to 37% of controls (156, 157). Since mevalonate or its lactone do not inhibit reductase or reductase phosphatase directly in vitro the intracellular modulator must be a metabolic derivative of mevalonate (see Section I11,B). Among these, inorganic pyrophosphate (released during squalene synthesis), mevalonate pyrophosphate, and isopentenyl pyrophosphate inhibit reductase phosphatase activity in vitro (157). Other mevalonate products may be even more potent intracellular feedback signals (52). Also see recent studies with fibroblasts (205). The preceding observations provide convincing evidence that the feedback control system, in addition to regulating the balance between synthesis and degradation of HMG-CoA reductase, also acutely affects the expressed activity of the enzyme through the bicyclic phosphorylation system. From these data also emerges the possibility that phosphorylation of reductase would hasten its degradation. Although tentative clues are at hand neither the specific nature of the mevalonate products that participate in this aspect of the feedback circuit nor the mechanism by which these interact with the satellite enzymes of the bicyclic cascade are evident.

D. ENDOCRINE CONTROL In Sections III,A and B endocrines were presented as creating a need for cholesterol synthesis by signaling the production of a cholesterol-derived secretory product or of cellular proliferation necessitating new membrane formation. Net synthesis of HMG-CoA reductase would ensue as the cholesterol feedback signal became weaker. Nevertheless, it is equally plausible that hormones or growth factors directly influence the state of reductase, acutely, in anticipation of the ultimate, holistic cellular response. The action of insulin and glucagon on the state of phosphorylation of HMG-CoA reductase is reviewed in the following paragraphs.

7. HMG-COA REDUCTASE

203

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FIG. 5. Effect of mevalonate on total and expressed activity of HMG-CoA reductase. Rat hepatocytes were incubated with 5 mM mevalonolactone. At the indicated times microsomes were isolated for determination of total (0, 0) and expressed ( 0 ,W) reductase activity. [Reprinted by permission from Gibson et a!. (156)l

Early experiments with isolated hepatocytes pointed to acute control of reductase with insulin and glucagon through reversible phosphorylation (139). Beg et al. (117)proceeded to show that 32P incorporation into liver reductase of rats that received glucagon in two injections in a one hour period was increased twofold over controls. Microsomal reductase activity declined by 35%. Arebalo et al. (158) with a single intravenous injection of glucagon into unanesthetized rats revealed that liver reductase expressed activity was diminished 43% in only 5 min. The diurnal cycle displays dramatic changes in the activity of HMG-CoA reductase (1).As-previously indicated, much of this variation can be attributed to

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DAVID M. GIBSON AND REX A. PARKER

insulin induction of reductase synthesis and subsequently to glucagon repression. Thus, during ingestion of food and release of insulin reductase levels rise. With progressive starvation these levels fall. Ignoring for the moment what effect these same hormones have on proteolytic enzyme systems the question has been asked if the state of phosphorylation of reductase is a variable in the diurnal cycle. The first answers were that it was not, namely, the ratio of expressed to total activity did not seem to change even though the total activity became elevated 10- to 20-fold during feeding (159-161). This was inconsistent with the hepatocyte experiments showing that reductase did in fact undergo endocrinesignaled changes in phosphorylation state (139). Several considerations may explain why the E/T ratio remains low and invariant in animal experiments. 1. If a short-term drop in expressed activity (E) is followed more slowly by a decay in the total enzyme activity (T), the initial rapid drop in the E/T ratio will gradually rise as the value of T continues to fall. Thus, the initial and final states of E/T ratio in this model will become equal after a period of time. It is difficult

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FIG. 6. Diurnal variation of HMG-CoA reductase activity. Microsomes were rapidly isolated from liver of rats maintained on the indicated light-dark schedule. The top curve displays total reductase activity. The ratio of expressed reductase activity to total activity ([E]/[T]) is represented by the dotted lines. The bottom curve is the ratio of expressed activity to the calculated value of the inactive, phosphorylated enzyme ([E]/[EP]). [Redrawn after Easom and Zammit (163)l.

7. HMG-COAREDUCTASE

205

to define the time frame since the primary endocrine signals are themselves changing during the diurnal cycle in the intact animal ( i . e . , the insulin/glucagon ratio). 2. Severe endocrine disruptions attending the killing of animals (e.g., catecholamine release) cannot be disregarded in obtaining liver samples. Fluoride and EDTA preserve the expressed activity of reductase only from the time of homogenization of tissue. Studies with isolated hepatocytes preclude this particular interference and thus are able to reflect the immediate insulin-glucagon response (139). The latter issue has been addressed by Easom and Zammit (162, 163) who examined reductase in samples of liver rapidly removed and processed from anesthetized rats at intervals during the diurnal cycle (Fig. 6). The data of Easom and Zammit (163) show the expected rise and fall of total reductase activity (top curve in Fig. 6). In this study the value of the E/T ratio (middle curve) also varied in a striking manner indicating that in the first (feeding) phase the enzyme became dephosphorylated while the total reductase activity rose to a maximum level. Significantly at the beginning of the second phase the E/T value fell before the total reductase activity declined as if progressive phosphorylation presaged net enzyme degradation. The E/EP ratio (bottom curve), the ratio of expressed enzyme activity to the calculated portion of total enzyme phosphorylated (T - E), also showed this relationship. E.

CELLULAR PROLIFERATION

The state of activation of HMG-CoA reductase appears to increase in cells under conditions that stimulate proliferation. For example, the expressed activity of reductase and the rate of sterol synthesis diminish if cells are cultured at high cell density (54, 164). In three different implanted Morris hepatomas the state of activation of reductase (E/T) ranged between 0.53 and 0.73 in contrast to normal liver (0.15 and 0.25) (165). Solid hepatomas in rats are virtually insensitive to dietary cholesterol supplementation (166, 167). The E/T ratio varies in hepatocytes obtained during pre- and postnatal development in rats (168) as well as during liver regeneration (169). In the latter study it is of interest to note that the E/T ratio rose at the same time as the total reductase activity and fell preceding the observed decline in the total activity.

F. OTHERSITUATIONS The expressed activity of HMG-CoA reductase varies in response to a number of experimental interventions. Incubation of cultures of rat ileal epithelial cells for an hour with bicarbonate brought about a decrease in expressed reductase activity and a drop in the rate of cholesterol biosynthesis (170). Chronic ascorbic

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DAVID M . GIBSON AND REX A. PARKER

acid deficiency and, paradoxically, excessive ascorbic acid intake depressed the level of expressed reductase activity in liver but had no effect on the total enzyme level ( I 71). The hypoglycemic drug dichloroacetate decreased liver expressed reductase activity within an hour after its administration ( I 72).

VI. Reversible Phosphorylation and Degradation A.

BACKGROUND

In Section V emphasis was placed on reversible phosphorylation of HMGCoA reductase as a mechanism for acute control of the enzyme as mediated through endocrine signaling (glucagon and insulin), negative feedback control (sterol and nonsterol products of mevalonate), and cell proliferation (including liver embryogenesis and regeneration). The interaction of the several signal systems and the modulating enzymes that determine the degree of phosphorylation of reductase was uncertain both in terms of the intracellular messenger and the molecular targets. Insulin and glucagon appeared to affect the bicyclic system through the protein phosphatase activity. The still ill-defined products of mevalonate metabolism that comprise the feedback signal were found to impinge on both protein kinases and the phosphatase. The influence of cholesterol itself, by modulating the fluidity of the endoplasmic reticulum, possibly could have affected the degree of reductase phosphorylation by nudging the enzyme toward the cytosolic space. These vectors that acutely determine the activity of HMG-CoA reductase also influence the rate of degradation of the enzyme. As reviewed in Sections III,B, 1 and V the feedback modalities (sterol and nonsterol) and glucagon increased the rate of reductase degradation as determined by direct measurements of the enzyme protein mass. Although the mechanism of action of the endocrine and feedback signals was not defined, it was clear that reductase degradation was not simply an impairment of replacement synthesis. Cholesterol content of the endoplasmic reticulum was cited as being conducive to degradation, perhaps by displacement of the enzyme toward the cytosolic surface for proteolytic attack. In certain intracellular systems an increase in phosphorylation of HMG-CoA reductase appeared to herald degradation of the enzyme measured in terms of its total potential catalytic activity or immunoreactivity (Section V). This was the case with mevalonate products (including cholesterol and 25-hydroxysterol); glucagon, per se, as well as during the diurnal cycle; and in liver regeneration. B. I N VITROMODELSYSTEM

In view of the indications discussed in the previous section experiments were undertaken by Parker et al. (35)to test if the phosphorylation state of reductase

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could influence the rate of degradation of the enzyme in vitro. Prior to these studies it had been shown that the calcium-dependent thiol protease calpain (173, 174) would release the 53K soluble form of reductase from the microsomal 97K enzyme (20, 22, 36) (Section 11). With calpain-I1 purified from rat liver cytosol (174) and rat liver microsomes isolated in the presence of the thiol protease inhibitor leupeptin and EGTA it was found that both the membrane-bound 62K fragment and the soluble 53K reductase were generated from the 97K native enzyme (35). Pretreatment of these microsomes with MgATP and reductase kinase (which inactivated the microsomal reductase) enhanced up to sixfold the rate of conversion of 97K reductase to the 53K reductase by calpain-I1 in comparison to microsomes pretreated with protein phosphatase. This action of calpain-I1 was followed both by changes in reductase total activity and in enzyme mass (specific imrnunoblot transfer). Leupeptin as well as purified calpastatin, the specific cytosolic protein inhibitor of calpain (179, blocked the formation of membrane-bound 62K and the soluble reductase. Later studies indicated that the 62K reductase is probably a precursor of 53K, although the latter species may also be released directly from the 97K native-enzyme (192). Whether or not a calpain system participates in regulating reductase levels in vivo remains to be determined.

C. PHOSPHORYLATION-ACTIVATED PROTEOLYSIS At least three proteins are degraded preferentially in the phosphorylated state. The first to be described was the L-type pyruvate kinase of liver that is inactivated by CAMP-dependent protein kinase. In vitro the phosphorylated form of the enzyme is degraded by bacterial subtilisin at a 10-fold greater rate than the dephosphorylated species (176). Pyruvate kinase prelabeled with 32P is cleaved in vitro with calpain to give a fragment possessing the 32P-labeled site (177, 178). Two forms of pyruvate kinase have been isolated from rat liver during starvation: 56K and 51K. Since only the former is capable of being phosphorylated in vitro, it is considered to be the precursor of the 51K species (179). Incubation of yeast cells with glucose causes rapid phosphorylation and inactivation of fructose- 1,6-bisphosphatase which leads within an hour to irreversible inactivation of the enzyme (180). This finding was interpreted to mean that the phosphorylated form of the enzyme was more susceptible to endogenous proteolysis. A calcium-activated, neutral protease from cardiac muscle degrades phosphorylated troponin I in vitro 3-fold more rapidly than the dephosphorylated species (181). Still other examples that may fall into this pattern are yeast glutamate dehydrogenase (182) and slime mold ornithine decarboxylase (183). Other kinds of covalent modification of proteins increase their rates of degradation: attachment of the protein ubiquitin (184), glycosylation (185), and oxidation (186). The common thread is that a change in protein conformation has been induced

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by phosphorylation which although reflected principally in decreased or increased catalytic capacity is also manifest in the interaction with proteases. Even though the phosphorylation state of the soluble 53K reductase cannot be distinguished with its polyclonal antibody (187) one would expect that monoclonal antibodies would be useful probes for detecting conformational changes in the native 97K reductase. In any event it is reasonable to assume that phosphorylation of reductase exposes or sensitizes specific sites for proteolytic cleavage. Adjacent pairs of basic amino acids are frequently targeted for proteolytic cleavage (183, 188). Calpain seems to prefer tyrosine, methionine, or arginine residues preceded by leucine or valine (189). Of interest is the consideration that basic amino acids are often in the vicinity of serine phosphorylation sites (183, 190). Whether or not phosphorylation of serine in this particular relationship would influence an adjacent proteolytic site remains to be seen. Nevertheless, such a constellation exists in the reductase primary sequence (33, 36): valine-376, arginine-377, arginine-378, serine-379 near the point of scission of the 62K fragment (36). Beyond serine 379 are clumped 10 additional serines between residues 383 and 462. The realization that the phosphorylated and dephosphorylated forms of an interconvertible enzyme may be degraded at different rates with the same endoprotease presents the prospect that the turnover of proteins subject to reversible phosphorylation must be viewed differently. This is especially true of an enzyme such as HMG-CoA reductase which is readily modified by phosphorylation and exhibits a relatively rapid turnover (tlR < 3 h). With this in mind a model steady-state system for turnover of interconvertible enzymes has been proposed in which distinct fractional degradative rate constants may be assigned to the phosphorylated and dephosphorylated species (191). In this context the phosphorylation state of enzymes is a meaningful determinant of enzyme concentration in cells.

D. INTRACELLULAR PROTEOLYSIS The pathway of HMG-CoA reductase degradation in liver and other cells is not known. In suspensionsof hepatocytes spontaneous loss of total reductase activity and the more rapid decline in the presence of mevalonolactone are partially blocked with propylamine, monensin, amino acid mixtures, and insulin (192). In many studies these agents have been deemed diagnostic of autophagic-lysosomal degradation (193-195). Further, glucagon which enhances reductase degradation stimulates autophagic vacuole formation (195, 196). Athough it has been recognized for some time that lysosomal membranes can be phosphorylated (197, 198) it is not known if lysosomal activity is affected. Nevertheless, endocrine signaling in liver may engender degradation of reductase both through activation of the proteolytic apparatus and phosphorylation of the reductase protein substrate.

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The limiting event in reductase degradation must precede complete hydrolysis in lysosomes. The principal argument is that the first step is not observed in cells. The soluble 53K and membrane-bound 62K forms of reductase do not accumulate in intact cells, but do accumulate in broken cell preparations (Section 11). The observation that the thiol protease inhibitor leupeptin and EGTA block solubilization of reductase (release of the 53K fragment) pointed directly to the calcium-sensitive, neutral thiol protease calpain (Section 11). Calpain is a cytosolic enzyme in most tissues and is frequently bound to its specific inhibitor calpastatin (175). Calpain is an endoprotease that in liver at least is activated by contact with protein substrates in the presence of only 1-5 p M Ca2 (199). It is known to attack membrane-bound proteins (175). The fact that calcium-signaled protein kinase-C phosphorylates and inactivates reductase (129) leaves open the possibility that this kinase and calpain may act cooperatively in the degradation of reductase. Cytosolic proteases other than calpain cannot be excluded at this juncture until tested with 97K microsomal reductase. Of interest are neutral proteases that are deployed on the cytosolic surface of lysosomes (200). +

E.

BIOLOGICALSIGNIFICANCE

Through feedback and endocrine signaling reductase activity in liver is first acutely modulated then sustained in the same control mode by long-term determinants of enzyme concentration. Thus, insulin acutely activates reductase through dephosphorylation and the activation state is maintained by way of net synthesis, the resultant of increased transcription and diminished degradation, to a higher steady-state concentration of enzyme. The mass (concentration) of HMG-CoA reductase and its expressed activity may be linked coordinately to other enzymes catalyzing controlled steps in lipid biosynthesis (Fig. 4). The phosphorylation state of the functionally tied troika that regulates cholesterol formation, storage, and disposal in liver, respectively HMG-CoA reductase, acyl-CoA cholesterol acyltransferase and 7a-cholesterol hydroxylase, has been emphasized by Scallen and Sanghvi (12). A broader canvas would include all interconvertible enzymes in Fig. 4 that shunt metabolites into lipids. As pointed out earlier, the liver is exceptional in that it forms lipid products for export, an assignment well beyond most cell types that are concerned with routine replacement synthesis of membranes and, when signaled, with proliferation. Membrane elaboration requires balanced synthesis of specific protein and lipid molecules and must depend on feedback control of the metabolic and synthetic assembly lines. Cholesterol being a key ingredient in the mix of lipids comprising membranes may be the most versatile feedback signal and consequently is called upon to represent the status of membranes (i.e., providing information on

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whether or not more membrane should be produced by the cell). Biosynthetic precursors of cholesterol (sterol and nonsterol) share this responsibility. HMGCoA reductase in its role of limiting cholesterol synthesis is the logical sensor of cholesterol and thus of membrane requirements. Endocrine signaling subserving the organism would impose upon and perhaps override this basal cellular regulatory circuitry. While the mechanisms remain to be detailed it is probable that the phosphorylation state of HMG-CoA reductase is sensitively poised to respond to feedback and endocrine information and thus constrain or stimulate membrane formation.

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147. Harris, R. A,, Paxton, R., Powell, S. M., Goodwin, G. W., Kuntz, M. J., and Han, A. C. (1986). Adv. Enz. Reg. 25, 219. 148. Erickson, S. K., Shrewsbury, M. A , , Could, R. G., and Cooper, A. D. (1980). BBA 620,70. 149. Arebalo, R. E., Hardgrave, J. E., Noland, B. J., and Scallen, T. J. (1980). PNAS 77, 6429. 150. Arebalo, R. E., Hardgrave, J. E., and Scallen, T. J. (1981). JBC 256, 571. 151. Arebalo, R. E., Tormanen, C. D., Hardgrave, J. E., Noland, B. J., and Scallen, T. J. (1982). PNAS 79, 5 1. 152. Kleinsek, D. A,, Ranganathan, S., and Porter, J. W. (1977). PNAS 74, 1431. 153. Whitehead, T. R., Vlahcevic, 2. R., Beg, 2. H., and Hylemon, P. B. (1984). ABB 230,483. 154. Beg, Z. H., Stonik, J. A., and Brewer, H. B., Jr. (1984). PNAS 81, 7293. 155. Parker, R. A,, Ingebritsen, T. S., Geelen, M. J. H., and Gibson, D. M. ColdSpring Harbor Conf. Cell Proliferation 8, 609. 156. Gibson, D. M., Parker, R. A , , Stewart, C. S., and Evenson, K. J. (1982).Adv. EnzymeRegul. 20, 263. 157. Parker, R. A,, Evenson, K.J., and Gibson, D. M. (1983). In “Isolation, Characterization and Use of Hepatocytes” (R. A. Harris and N. W. Cornell, eds.), p. 609. Elsevier/North-Holland,

New York. 158. Arebalo, R. E., Hardgrave, J. E., Sena, G. R., and Scallen, T. J. (1987). J . Lipid Res. (in 159. 160. 161. 162. 163. 164. 165. 166. 167. 168. 169. 170. 171. 172. 173. 174. 175. 176. 177. 178. 179. 180. 181. 182. 183. 184. 185.

press). Brown, M. S., Goldstein, J. L., and Dietschy, J. M. (1979). JBC 254, 5144. Kleinsek, D. A,, Jabalquinto, A. M., and Porter, J. W. (1980). JBC 255, 3918. Jenke, H.-S., Lowel, M.,and Bemdt, J. (1981). JBC 256, 9622. Easom, R. A,, and Zammit, V. A. (1984). BJ 220, 733. Easom, R. A,, and Zammit, V. A. (1984). BJ 220, 739. Sexton, R. C., Panini, S. R., and Rudney, H. (1982). FP 41, 1388. Feingold, K. R., Wiley, M. H., Moser, S. H., and Siperstein, M. D. (1983). ABB 226, 231. Siperstein, M. D. (1984). J. LipidRes. 25, 1462. Gregg, R. G., and Wilce, P. A. (1983). In “HMG CoA Reductase” (J. R. Sabine, ed.), p. 245. CRC Press, Boca Raton, Florida. Leoni, S., Spagnuolo, S . , Conti-Devirgilis, L., Dini, L., Mangiantini, M. T., andTrentalance, A. (1984). J . Cell. Physiol. 118, 62. Trentalance, A., Leoni, S., Mangiantini, S . , Spagnuolo, S., Feingold, K., Hughes-Fulford, M., Siperstein, M., Cooper, A. D., and Erickson, S. K. (1984). BBA 794, 142. Panini, S. R., and Rudney, H. (1980). JBC 255, 11633. Holloway, D. E., Peterson, F. J., Prigge, W. F., and Gebhard, R. J. (1981). BBRC 102, 1283. Stacpoole, P. W., Hanvood, H. J., Jr., Varnado, C. E., and Schneider, M. (1983). J . C/in. Invest. 72, 1575. Nishiura, I., Tanaka, K., Yamato, S . , and Murachi, T. (1978). J. Biochem. (Tokyo) 84, 1657. DeMartino, G. N., and Croall, D. E. (1983). Biochemistry 22, 6287. Murachi, T . (1983). Calcium Cell Funct. 4, 377. Bergstrom, G., Ekman, P., Humble, E., and Engstrom, L. (1978). BBA 532, 259. Ekmdn, P., and Eriksson, I. (1980). Acta Chem. S c u d . , Ser. B B34, 419. Dahlqvist-Edberg, U., and Ekman, P. (1981). BEA 660, 96. Hall, E. R., McCully, V., and Cottam, G. L. (1979). ABB 195, 315. Miiller, D., and Holzer, H. (1981). BBRC 103, 926. Toyo-Oka, T. (1982). BBRC 107, 44. Hemmings, B. A. (1984). Mol. Aspects Cell. Regul. 3, 155. Kuehn, G. D. (1984). Mol. Aspects Cell. Regul. 3, 185. Hershko, A , , and Ciechanover, A. (1982). Annu. Rev. Biochem. 51, 335. Barbaric, S., Mrsa, V., Ries, B., and Mildner, P. (1984). ABB 234, 567.

7.

HMG-COA REDUCTASE

215

Rivett, A. J. (1985). JBC 260, 300. Rogers, D. H., and Rudney, H. (1982). JBC 257, 10650. Holzer, H., and Heinrich, P. C. (1980). Annu. Rev. Biochem. 49, 63. Sasaki, T., Kikuchi, T., Yumoto, N., Yoshimura, N., and Murachi, T. (1984). JBC 259, 12489. 190. Cohen, P. (1980). Mol. Aspects Cell. Regul. 1, 255. 191. Gibson, D. M., Hamilton, J. A., and Parker, R. A. (1984). J. Bioenerg. Biomembr. 16,433. 192. Parker, R. A., Miller, S. J., and Gibson, D. M. In “Drugs Affecting Lipid Metabolism” (D. Kritchevsky, ed.) Springer Verlag, Heidelberg (in press). 193. Mortimore, G. E. (1982). Nutr. Rev. 40, 1. 194. Seglen, P. O., and Gordon, P. B. (1982). PNAS 79, 1889. 195. Schworer, C. M., and Mortimore, G. E. (1979). PNAS 76, 3169. 1%. Ballard, F. J. (1980). In “Biochemical Actions of Hormones” (G. Litwack, d.), Vol. 7, p. 91. Academic Press, New York. 197. Zahlten, R. N., Hochberg, A. A., Stratmen, F. W., and Lardy, H. A. (1972). PNAS 69,800. 198. Wells, W. W., Collins, C. A , , and Kurtz, J. W. (1981). I n “Lysosomes and Lysosomal Storage Diseases” (J. W. Catlahan and J. A. Lowden, eds.), p. 17. Academic Press, New York. 199. Pontremoli, S., and Melloni, E. (1986). Ann. Rev. Biochem. 55, 455. 200. Pontremoli, S., Melloni, E., Michetti, M.,Salamino, F., Sparatore, B., and Horecker, B. L. (1982). BBRC 106, 903. 201. Chin, D. J., Gil, G., Eaust, J. R.,Goldstein, J . L., Brown, M.S., and Luskey, K.L. (1985). Mol. Cell Biol. 5, 634. 202. Gil, G., Faust, J. R., Chin, D. J., Goldstein, J. L., and Brown, M. S. (1985). Cell 41, 249. 203. Henneberg, R., and Rodwell, V. W. (1985). Physiol. Chem. Pfzys. Med. NMR 17, 35. 204. Gibbons, C . F., Bjomsson, 0. G., and Pullinger. C. R. (1984). JBIBC 259, 14399. 205. Beg, Z. H., Reznikov, D. C., and Avigan, J. (1986). ABB 244, 310. 186. 187. 188. 189.

This review is dedicated to the memory of John W . Porter, University of Wisconsin, whose discoveries related to the control of lipogenesis have provided many insightsfor future explorations.

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Aromatic Amino Acid Hydroxy lases SEYMOUR KAUFMAN Laboratory of Neurochemistry Alcohol, Drug Abuse, and Mental Health Administration National lnstiiuie of Mental Health Beihesda, Maryland 20205

.............................. ............ A. Physical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Phenylalanine Hydroxylase Phosphatase ..........................

218 22 1 22 1 225

C. Phosphorylation of Rat Liver Phenylalanine Hydroxylase in Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . nine Hydroxylase Activity in

226

D. Effect of GI

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

230

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

235

Phenylalanine Hydroxylase ..................................... G. Physiological Significance of the Regulation of Phenylalanine Hydroxylase Activity by Phosphorylation-Dephosphorylation . . . . . . . . 111. Tyrosine Hydroxylase . . . . . . . . . . . . . . . . .. A. Introduction ..................... .. B. Physical Properties ............ C. Activation by Phos ............ D. The stability of the Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Tryptophan Hydroxylase A. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

217 THE ENZYMES, Vol. XVllI

239 242 24 8 248 249 25 1 270 27 1 27 1 272

218

SEYMOUR KAUFMAN C. Physical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Regulation by Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1.

274 275 277

Introduction

The three pterin-dependent enzymes, phenylalanine, tyrosine, and tryptophan hydroxylase, share enough properties, both physical and catalytic, that it is useful to consider them as members of a family of enzymes (1). This classification has predictive value since there have been many instances where the discovery of a regulatory property for one of the enzymes-often first with phenylalanine hydroxylase-has led to the finding that this property is shared by the others. The general characteristics of this group of enzymes were first established from studies carried out with hepatic phenylalanine hydroxylase. The feature that clearly distinguishes them from other enzymes is that they all require a naturally occurring unconjugated pterin. This nonprotein cofactor was isolated from rat liver extracts on the basis of its ability to stimulate the conversion of phenylalanine to tyrosine in the presence of an active hydroxylating system (2, 3 ) , and shown to be the reduced form of the unconjugated pterin, 2-amino-4-hydroxy-6-[1,2-dihydroxypropyl-(~-erythru)-pteridine] (4, whose trivial name is biopterin (“pterin” is the name for a 2-amino-4-hydroxypteridine). The structure for tetrahydrobiopterin (BH,), the active form of the coenzyme, is shown in Fig. 1. Although biopterin had been isolated previously from human urine (3,its function, if any, in mammals was obscure. The demonstration that BH, is an essential component of the phenylalanine hydroxylating system established the first metabolic role for any unconjugated pterin and the first coenzyme role for BH,. In addition to the naturally occurring coenzyme, several synthetic tetrahydropterins, such as 6-methyltetrahydropterin (6MPH,) and 6,7-dimethyltetrahydropterin (DMPH,), were shown to be active with the phenylalanine hydroxylating system ( 3 , 6 ) . In fact, the maximum velocity of the unactivated hydroxylase

FIG. 1. Structure of tetrahydrobiopterin.

219

8. AROMATIC AMINO ACID HYDROXYLASES

0 II

O2 +

H

Phenylalanine

Phenylalanine Hydroxylase

Dihydropteridine Reductase

NADH

+

H+

H

FIG.2. The phenylalanine hydroxylating system: the reactions catalyzed by phenylalanine hydroxylase and dihydropteridine reductase.

from rat liver (3, 7) and from human liver (8, 9) is much greater with these synthetic compounds than it is with BH,. In addition to BH,, the other essential components of the phenylalanine hydroxylating system are phenylalanine hydroxylase and dihydropteridine reductase (DHPR). The reactions catalyzed by these two enzymes with 6MPH, as the coenzyme are illustrated in Fig. 2. As can be seen, phenylalanine hydroxylase catalyzes a coupled reaction in which L-phenylalanine is oxidized to L-tyrosine and the tetrahydropterin is oxidized to an extremely unstable quinonoid dihydropterin (10); molecular oxygen is the electron acceptor and is normally reduced to water (11). The second essential enzyme of the system, DHPR, catalyzes the reduction of the quinonoid dihydropterin back to the tetrahydro level, utilizing a reduced pyridine nucleotide as the electron donor (12). This reaction serves to regenerate the active form of the pterin coenzyme and thus allows the coenzyme to function catalytically. After the roles for BH, and DHPR had been established in the phenylalanine hydroxylating system, they were shown to be essential components of the tyrosine (13, 14) and tryptophan (15) hydroxylating systems. An example of how our knowledge of the characteristics of these other two hydroxylating systems relies heavily on prior results obtained with the phenylalanine hydroxylating system is provided by the fact that the oxidized pterin product that is formed during the course of tyrosine and tryptophan hydroxylation has not been rigorously identified as the quinonoid dihydropterin. That there can be little doubt that it is the same product as the one formed during phenylalanine hydroxylation is based on the demonstration, mentioned previously, that DHPR is an essential component of the tyrosine and tryptophan hydroxylat-

220

SEYMOUR KAUFMAN

ing systems. It is also known that in the presence of BH, and DHPR, the stoichiometry of the enzymic conversion of tyrosine to 3 ,bdihydroxyphenylalanine and of tryptophan to 5hydroxytryptophan catalyzed by tyrosine hydroxylase (14) and tryptophan hydroxylase ( 1 3 , respectively, is the same as that described for phenylalanine hydroxylation (Fig. 2). All three hydroxylation reactions are accurately described by the general reaction shown in Eq. 1 , where RH is the amino acid substrate, ROH is the hydroxylated amino acid product, and q-BH, is quinonoid dihydrobiopterin. RH

+ BH4 + 0 2

--*

ROH

+ q-BH2 + H2O

(1)

Although the idea that these enzymes are members of a family with many common properties has been of value, it should not be pushed too far. Thus, although the general mode of regulation of all three enzymes may be similar, the precise manner in which a given regulatory property is expressed is often unique for each enzyme. There are also regulatory mechanisms that appear to operate with only one of the enzymes. Some of the diversity in the regulatory behavior of these enzymes can be understood if one looks at the different hydroxylases from the perspective of the role that they play in the functioning of the organism. When viewed in this way, it is clear that the role of phenylalanine hydroxylase is different from that of the other two enzymes. Phenylalanine hydroxylase catalyzes the rate-limiting step in a catabolic pathway, the only pathway that leads to the complete oxidation of phenylalanine to CO, and water (16). It would be expected, therefore, that the activity of phenylalanine hydroxylase would be responsive to blood levels of phenylalanine, which in turn are determined in part by the dietary intake of this amino acid. The expectation has been realized-there is evidence that elevated levels of phenylalanine activate phenylalanine hydroxylase (7, 17-24 both directly, by altering the enzyme’s conformation, and indirectly, by increasing the enzyme’s level of phosphorylation. In contrast to phenylalanine hydroxylase, it is clear the neither tyrosine hydroxylase nor tryptophan hydroxylase are involved in the catabolism of their amino acid substrates; the catabolism of these amino acids involves other pathways. Rather, these hydroxylases catalyze steps in neurotransmitter synthesis and one would expect that the enzymes would not be particularly responsive to tissue levels or dietary intake of their substrates, but rather to the needs of the organism for neural transmission. It also seems likely that the way in which activation of these enzymes is expressed may be different because it will depend on the relationship between K,,, values for substrate and coenzyme and tissue levels of these compounds. There are indications that the K,, values of phenylalanine hydroxylase for both phenylalanine and tetrahydrobiopterin are not too far from their concentrations in liver (22). In such a situation, only limited degrees of activation of the enzyme

8. AROMATIC AMINO ACID HYDROXYLASES

22 1

could be achieved by mechanisms that led to a decrease in the enzyme’s K, for substrate or cofactor. It would seem reasonable, therefore, that activation of this enzyme would be expressed mainly by an increase in V,,,. There is evidence that this is the case. By contrast, the K, values of both tyrosine and tryptophan hydroxylase for tetrahydrobiopterin appear to be considerably larger than brain concentrations of this pterin (1). In addition, the K, value of tryptophan hydroxylase (determined in the presence of tetrahydrobiopterin) also appears to be larger than normal brain concentrations of this amino acid (1). Based on these considerations, it would seem likely that activation of both of these hydroxylases would be expressed, in part, by a decrease in K, for tetrahydrobiopterin and that for tryptophan hydroxylase, it might also involve a decrease in K, for tryptophan. Again, with these enzymes, the results appear to be a consonant with most of these expectations.

II. Phenylalanine Hydroxylase A.

PHYSICAL PROPERTIES

Essentially pure phenylalanine hydroxylase isolated from rat liver by the procedure of Kaufman and Fisher was shown to be a mixture of two different polymeric forms (23). From a determination of their Stokes radii and sedimentation constants, the molecular weights of these two species were estimated to be 110,000 and 210,000 (23). Since the molecular weight of the subunit(s) of the enzyme is in the range of 49,000 to 51,000 (23), these two major forms of phenylalanine hydroxylase correspond to the dimer and tetramer, respectively. Through the use of high-performance gel-permeation chromatography (HPGPC), our original finding that rat liver phenylalanine hydroxylase exists as a mixture of tetramers and dimers has been confirmed (24). At 25°C in 0.1 M potassium phosphate, pH 6.8, the major peak was found to correspond to a molecular weight of 200,000, whereas the minor peak eluted as a M, = 100,000 species. Analysis by HPGPC of 10 different preparations of rat liver phenylalanine hydroxylase purified either by the method of Kaufman and Fisher (23) or by that of Shiman et af. (20), showed that the tetrameric species accounts for 78 6% of the total hydroxylase protein (24). Although Daskeland et al. (25) have also reported that the rat liver enzyme exists as a mixture of dimers and tetramers, conflicting results have been reported by others. Nakata and Fujisawa (26) and Shiman (27), for example, reported that it exists solely as tetramers, whereas Tourian (18) reported that it exists solely as dimers. A partial explanation for some of these divergent findings may be provided by

*

222

SEYMOUR KAUFMAN

our observation that the oligomeric composition of the hydroxylase can change during long periods of frozen storage, with more dimers being formed at the expense of tetramers (24).It is also known that preincubation of the enzyme with phenylalanine can alter the oligomeric composition of the enzyme ( l a ) ,leading to the formation of even higher molecular weight forms of the enzyme than M, = 200,000 (12). The finding that the percentage of dimers increases on frozen storage of the enzyme makes it seem likely that the predominant form of the freshly prepared enzyme is the tetramer. As was mentioned previously, the maximum velocity of rat liver phenylalanine hydroxylase is much greater in the presence of synthetic analogues of BH,, such as 6MPH, and DMPH,, than it is in the presence of BH,. This relatively low hydroxylase activity with BH,, however, can be selectively increased 20- to 30-fold by a wide variety of reversible and irreversible modifications of the enzyme, including interaction with phospholipids (28, 29), limited proteolysis (28, 29), and alkylation of a sulfhydryl group by N-ethylmaleimide (30). Since activations of this type have little, if any, effect on hydroxylase activity measured in the presence of 6MPH, or DMPH,, the ratio of activity in the presence of BH, and one of these model cofactors serves a useful index of the state of activation of the hydroxylase with either the pure enzyme or with the enzyme in crude liver extracts. This index, for example, provided the first hint that the hydroxylase undergoes a still poorly understood activation during the course of its purification from rat liver extracts (31, 32). Activation of rat liver phenylalanine hydroxylase by phosphorylation catalyzed by CAMP-dependent protein kinase follows the pattern that had been established with the other types of activation, i.e., it leads to a selective 2- to 4fold increase in the activity in the presence of BH, but not in the presence of DMPH, or 6MPH, (33). Activation by phosphorylation, does not change the relationship between initial velocity and phenylalanine concentration from sigmoid to hyperbolic as do the kinds of activation previously mentioned. Furthermore, the activation of the hydroxylase by phosphorylation does not alter the apparent K , for BH, or for phenylalanine (33). Activation of the hydroxylase in the presence of [y32P]ATP,cAMP,Mg2 , and CAMP-dependent protein kinase is accompanied by the incorporation of approximately 0.70 mol of 32Piper M, = 50,000 subunit (33). The finding that less than stoichiometric amounts of phosphate are incorporated into the pure hydroxylase in vitro made it seem likely that the enzyme isolated from rat liver is already partially phosphorylated. Determination of the amount of protein-bound phosphate in the hydroxylase confirmed this possibility; in five different preparations of the enzyme that had been isolated by the Kaufman and Fisher procedure (23), the average phosphate content per mol of M, = 50,000-subunit was 0.31 mol with a range of 0.23 to 0.42 (33). Subsequently, it was found (34) that phenylalanine hydroxylase isolated from +

8 . AROMATIC AMINO ACID HYDROXYLASES

223

rat liver by the procedure of Shiman et al. (20), a procedure that involves adsorption of the enzyme on phenyl-Sepharose, contains about one-third the amount of protein-bound phosphate-about 0.07 mol Pi per M, = 50,000 subunit-than does the enzyme isolated by the Kaufman and Fisher procedure, an indication that phosphorylated species of the enzyme might be selectively retained by the hydrophobic matrix. This possibility was strongly supported by the finding that hydroxylase prepared by the Kaufman and Fisher procedure (0.28 mol Pi per M, = 50,000 subunit) after adsorption to and elution from phenylSepharose contained only one-third of its original protein-bound phosphate (0.10 mol Pi per M, = 50,000 subunit) (34.’ The amino acid sequence at the site of rat liver phenylalanine hydroxylase that was phosphorylated by incubation with [32P]ATPin the presence of the catalytic subunit of pig muscle CAMP-dependent protein kinase has been reported to be Ser-Arg-Ly~-Leu-[~~P]SerP-ASX-Phe-Gly-Glx-Glx (38).From their finding that the amount of this peptide in their sample was at least twice that calculated from the radioactivity of the sample, they concluded that the peptide contained a substantial amount of endogenous phosphate. These results, therefore, provide independent evidence in support of the conclusion that hepatic phenylalanine hydroxylase in untreated rats is a mixture of phosphorylated and nonphosphorylated forms. The observation that rat liver phenylalanine hydroxylase is a mixture of phosphorylated and nonphosphorylatedspecies provided an explanation for the earlier observation of Barringer and co-workers (39) that three forms of phenylalanine hydroxylase are detectabIe when crude rat liver extracts are chromatographed on cellulose-calcium phosphate gel columns. Although there was no evidence to support the idea that these three forms had arisen from genetically determined differences in the primary structure of the enzyme, they were nonetheless designated “isozymes.” The results that have just been reviewed suggested the possibility that the ‘Shiman er al. (35) have reported that the hydroxylase purified from control rats by their phenylSepharose procedure (20) contains the same amount of protein-bound phosphate, namely 0.24 +. 0.05 m o l h o l of hydroxylase subunit, as that originally reported by Abita er al. (33)for the enzyme purified by the Kaufman and Fisher procedure (23).They suggested that the finding (34) of only about one-third of this amount of protein-bound phosphate in enzyme isolated by the phenylSepharose procedure was due to the action of a phosphatase. This explanation for the discrepancy between these two results ignores the finding that essentially pure phenylalanine hydroxylase isolated by the Kaufman and Fisher method (23) and containing 0.28 mol phosphate/mol of hydroxylase has been found (34) to lose two-thirds of its protein-bound phosphate on being adsorbed to and eluted from phenyl-Sepharose according to the procedure of Shiman et al. (20). It also does not take into account the report that phenylalanine hydroxylase isolated by the Kaufman and Fisher procedure has no detectable “phenylalanine hydroxylase phosphatase” activity (36).As far as this discrepancy is concerned, it is noteworthy that D~skelandef al. (37), reported that phenylalanine hydroxylase isolated from rat liver by the phenyl-Sepharose procedure of Shiman et al. (20) contains 0.07 mol phosphatehol of hydroxylase subunit, the same value reported by Parniak er al. (34).

224

SEYMOUR KAUFMAN

different forms of the enzyme seen on calcium phosphate gels were being separated on the basis of their different degrees of phosphorylation. In support of this idea, it was found that highly purified preparations of the two major forms of the hydroxylase, designated I1 and 111, differed in their ratios of BH,-dependent and 6MPH,-dependent hydroxylase activities. This ratio for form 111 was about twice as high as that for form I1 (40),an indication that form 111 was more highly activated than form I1 (i.e., it was more highly phosphorylated). Subsequently, it was shown that the kinase-catalyzed phosphorylation of phenylalanine hydroxylase in vitro not only activates the hydroxylase, but also leads to a marked change in its elution pattern from calcium phosphate gels. Specifically, it was found that upon phosphorylation, forms I1 and 111 are converted to a new form (designated form IV) that is bound more tightly to calcium phosphate than are the other forms (36, 40).These results supported the idea that a major structural feature determining the elution pattern of at least the two major forms of the enzyme is their content of protein-bound phosphate. Direct determination of the amounts of protein-bound phosphate in purified forms 11, 111, and IV provided additional support for this notion (36). It was found that form I1 has about 0.25 mol of phosphate per 50,000-M, monomer, form 111 has about 0.50 mol of phosphate per 50,00044, monomer, and form IV has about 1.0 mol of phosphate per 50,000-M, monomer. From these results, a model for the subunit structure of phenylalanine hydroxylase was formulated, shown in Fig. 3, where form I1 is the monophosphorylated tetramer, form 111 is the half-phosphorylated tetramer, and form IV is the fully phosphorylated tetramer. In this scheme, form I, which is a minor species accounting for only 5 to 10% of the total hydroxylase activity in crude extracts, is shown as the nonphosphorylated form of the enzyme. Despite the fact that phosphate analysis carried out on form I showed that it contains very little protein-bound phosphate (about 0.05 mol per mol M, = 50,000 subunit) (34), there are indications that it does not form part of a coherent series with species 11,111, and IV. Thus, on phosphorylation with ATP and CAMP-dependent protein kinase, it is not converted to any of the other three forms of the enzyme (41). These findings indicate that form I differs from forms 11, 11, and IV by more than just its phosphate content and suggest that form I has undergone an additional structural modification, either in vivo or during the course of its purification. This modification has not been identified.

88 €BPp83p

pp%pp

I

II

Ill

IV

FIG.3 . Model for the various phosphorylated forms of rat-liver phenylalanine hydroxylase

8. AROMATIC AMINO ACID HYDROXYLASES

B.

225

PHENYLALANINE HYDROXYLASE PHOSPHATASE

The observation that phenylalanine hydroxylase isolated from rat liver extracts by the procedure of Kaufman and Fisher (23) contains less than 1 mol phosphate/mol of hydroxylase subunit (33) provided the earliest indication that a phosphatase might be present in liver that can catalyze the dephosphorylation of phosphorylated phenylalanine hydroxylase. The existence of such an activity was also indicated by the finding, to be discussed in detail in Section C , that the acute activation of phenylalanine hydroxylase that is observed on administration of glucagon to rats and which is due to in vivo phosphorylation of the enzyme, is transient, lasting only about two hours (42). A phosphatase has been purified from rat liver extracts on the basis of its ability to catalyze the dephosphorylation of phenylalanine hydroxylase (43, 44). Although specificity studies carried out with partially purified preparations of this enzyme suggested that it might be rather specific for phosphorylated phenylalanine hydroxylase (43), subsequent studies with highly purified preparations have shown that it can catalyze the dephosphorylation of a variety of phosphorylated proteins, including glycogen synthase and phosphorylase a (45). The properties of the purified enzyme, including its relative lack of sensitivity to protein phosphatase inhibitor-1 and -2 (46, 47) and its activity in the absence of added cations (45), indicate that the preparation of purified phenylalanine hydroxylase phosphatase would be classified according to the system devised by Ingerbritsen and Cohen (48) as a type 2A phosphatase. Phenylalanine hydroxylase phosphatase has been used to study in greater detail the relationship between the phosphate content of the hydroxylase and its BH,-dependent hydroxylase activity. When 32P-labeled hydroxylase containing about 1 mol of phosphate/mol M, = 50,000 subunit, prepared by phosphorylation of the hydroxylase in v i m with [32P]ATP and CAMP-dependent protein kinase, was incubated with the phosphatase, it was found that as protein-bound phosphate was cleaved from the hydroxylase by the action of the phosphatase, a simultaneous decrease in BH,-dependent enzyme activity was observed. This relationship between dephosphorylation of the enzyme and decrease in BH,dependent hydroxylase activity (the 6MPH,-dependent activity is relatively uneffected by the state of phosphorylation of the enzyme), however, is not a linear one. Figure 4 shows a plot of the ratio of BH,-dependent to 6MPH,-dependent hydroxylase activity against the phosphate content of the hydroxylase. It is clear that the loss of 70-75% of the protein-bound 32P is accompanied by a progressive loss of BH,-dependent hydroxylase activity. In contrast, the release of the remaining 20-25% of the protein-bound 32Pleads to little further decrease in BH,-dependent activity. These results indicate that fully dephosphorylated hydroxylase molecules and those containing 25-30% of the maximum phosphate do not differ in their BH,-dependent phenylalanine hydroxylase activities (44). Furthermore, from the results shown in Fig. 4,the most marked changes in BH,-

226

SEYMOUR KAUFMAN

PROTEIN-BOUND,*P REMAINING I % )

FIG.4. The dephosphorylation of fully phosphorylated phenylalanine hydroxylase (phosphorylated in v i m ) by phenylalanine hydroxylase phosphatase; effect of dephosphorylation on BH4dependent and 6MPH4-dependent hydroxylase activity. Data from Ref. (44).

dependent hydroxylase activity would be expected to occur on interconversion of the quarter-phosphorylated tetramers (form 11) and the half-phosphorylated tetramers (form 111). The conclusion that nonphosphorylated phenylalanine hydroxylase is not only not devoid of activity but has essentially the same BH,-dependent hydroxylase activity as the enzyme containing about 0.25 rnol of phosphate/mol of subunit (form 11) is in accord with the observation that essentially pure phenylalanine hydroxylase isolated by the procedure of Shiman et al. (20) has a high specific activity even though it contains only 0.06 to 0.07 mol phosphate/mol M, = 50,000 subunit (34). It also agrees with our observation that when hydroxylase containing 0.26 mol phosphate/mol subunit is absorbed to and eluted from phenyl-Sepharose according to the procedure of Shiman et al. (20), it loses about 75% of its phosphate (0.07 rnol phosphate/mol subunit), but it loses little if any of its BH,-dependent activity (34). OF RAT LIVERPHENYLALANINE C. PHOSPHORYLATION HYDROXYLASEIN VIVO

Studies on the modulation of the activity of purified rat liver phenylalanine hydroxylase by in vitro phosphorylation and dephosphorylation suggested that hydroxylase activity could be regulated in vivo by similar mechanisms. Since it was known that hormones such as glucagon could increase hepatic levels of CAMP(49), the effect on phenylalanine hydroxylase activity of administration of

8.

AROMATIC AMINO ACID HYDROXYLASES

221

this hormone to rats was studied. It was found that the intraperitoneal injection of glucagon to rats led to a 4-fold activation of hepatic phenylalanine hydroxylase (42). The effect was rapid, with activation being detectable within 30 min, and transient, with hydroxylase activity returning to basal levels within 2 h. The finding that the glucagon stimulation could be elicited at least one more time by a repeat injection of glucagon indicated that the decay of the activated state was due to dephosphorylation of the hydroxylase rather than to its proteolytic degradation. As with activation of the hydroxylase in vitro by the action of CAMPdependent protein kinase in the presence of ATP, this effect of glucagon was detectable when the hydroxylase was assayed in the presence of BH, but not in the presence of synthetic cofactor analogues such as DMPH,. When the glucagon was administered intraperitoneally, the dose required to produce half-maximal effects was 270 Fg/kg (42). When the glucagon was administered intravenously, the half-maximal response was observed at only 15 pg/kg (50). That the glucagon-mediated activation is due to an increase in the extent of phosphorylation of the hydroxylase in vivo was demonstrated in experiments where rats were given radioactive inorganic phosphate following the administration of the glucagon. It was found that the stimulation of enzyme activity was accompanied by the incorporation of 32Pi into the hydroxylase to the extent of 0.7 mol/mol of hydroxylase subunit (42).Furthermore, when liver extracts from glucagon-treated rats were chromatographed on cellulose-calcium phosphate columns, most of the hydroxylase migrated in the same position as form IV, the fully phosphorylated species (1 .O mol phosphate/mol hydroxylase subunit) (42). Finally, as can be seen in Fig. 5, when fully phosphorylated hydroxylase obtained from livers of glucagon-treated rats was treated in vitro with crude phosphatase, more than 90% of the hydroxylase was eluted as form I1 (44). Additional evidence that phenylalanine hydroxylase can be phosphorylated by CAMP-dependent protein kinase in vivo was provided by the observation that treatment of rats with phosphodiesterase inhibitors such as aminophylline converted all of the hydroxylase to form IV, the fully phosphorylated species (44). The failure of Brand and Harper (51) to observe any increase in hepatic phenylalanine hydroxylase activity in rats treated for 10 days with glucagon (1.25 mg/kg every 12 h, subcutaneously) was most likely due to their use of DMPH, in their assays of phenylalanine hydroxylase activity. As previously pointed out, the stimulation of phenylalanine hydroxylase by phosphorylation is detectable when the enzyme is assayed with BH, but not with DMPH, (33). The level of phosphorylation of phenylalanine hydroxylase in livers from untreated rats represents a balance between the rates of phosphorylation and rates of dephosphorylation of the hydroxylase. The observation that the basal enzyme contains about 0.25 to 0.30 mol phosphate/mol M, = 50,000 subunit (33) indicates that in control rats dephosphorylation is the dominant process. From these considerations, it was anticipated that the glucagon-mediated activation

228

SEYMOUR KAUFMAN

1

TUBE NUMBER

FIG.5. Calcium phosphate-cellulose chromatographic patterns of in vivo phosphorylated phenylalanine hydroxylase before and after treatment with phenylalanine hydroxylase phosphatase. Phosphorylation was mediated by intraperitoneal injection of glucagon. Incubation with phosphatase was carried out for 60 min at 250°C. Control extract patterns were obtained after incubations under the same conditions but in the absence of added phosphatase: (0) control extracts; ( 0 )extracts after treatment with phosphatase. Data from Ref. (44).

(and phosphorylation) of the hydroxylase would be even more transient than it is. In an attempt to understand the mechanism by which the hydroxylase remains elevated for as long as 60 min after an intraperitoneal glucagon injection (42), the activities of phenylalanine hydroxylase phosphatase and CAMP-dependent protein kinase in livers from glucagon-treated rats was studied. It was found that the latter activity was elevated 4-fold within 15 min of an intraperitoneal injection of glucagon ( 2 mg/kg), returning to basal levels 30 min after the injection. Thirty minutes after the glucagon injection, phenylalanine hydroxylase phosphatase activity was decreased 29%, was still decreased 15 min later, and had returned to, or exceeded slightly, preinjection levels after another 15 min (i.e., 60 min after the glucagon injection) (52). These results indicate that although activation of protein kinase lasts for only a short time after the glucagon injection, phenylalanine hydroxylase is maintained in an activated and phosphorylated state after this time, in part because of a decrease in the hepatic activity of phenylalanine hydroxylase phosphatase. In contrast to these results obtained in vivo, it has been reported that in rat hepatocytes, the activity of phosphorylase phosphatase (the enzyme that, as previously mentioned, may be the same as phenylalanine hydroxylase phosphatase) has been reported to be unaffected by glucagon (53).It should be noted, however, that in these experiments, the hepatocytes were exposed to glucagon for only 2 min, a period that might have been too short to elicit the small inhibition of the phosphatase observed in the intact rat.

8. AROMATIC AMINO ACID HYDROXYLASES

229

Activation of phenylalanine hydroxylase by glucagon has also been demonstrated in hepatocytes isolated from rats (54, 55). Our finding that glucagon increases the extent of phosphorylation of the hydroxylase has also been confirmed with rat liver hepatocytes (55, 56). Fisher and Pogson (57) have studied in greater detail the glucagon-stimulated phosphorylation of phenylalanine hydroxylase in hepatocytes. They found that the rate of incorporation of 32Pi into phenylalanine hydroxylase was actually slower in the presence of glucagon than in its absence. By contrast, when glucagon was added 75 min after the addition of 32Pi, phosphorylation of phenylalanine hydroxylase was stimulated for a period of 5 min, at which time a new steady-state of phosphorylation was attained. During this 5-min burst of phosphorylation, the amount of protein-bound [32P]phosphate increased from 0.29 mol/mol of hydroxylase subunit to 0.56 mol/mol of subunit (i.e., under these conditions glucagon stimulated the incorporation of only an additional 0.27mol PJmol subunit). Dibutyryl-CAMP stimulated the incorporation of 32Piinto phenylalanine hydroxylase to about the same extent as did glucagon. The glucagonstimulated phosphorylation of the hydroxylase correlated with about a 2-fold increase in phenylalanine hydroxylase activity measured in situ. From an extrapolation of these results, the authors concluded that there is a linear correlation between phenylalanine hydroxylase activity and extent of phosphorylation, with the extrapolated hydroxylase activity of the nonphosphorylated species being equal to about 8% of the maximum activity, the latter value taken to be the activity expressed by the enzyme containing about 0.60 mol 32Pi/molof subunit. The conclusion of Fisher and Pogson (57) that there is a linear relationship between extent of phosphorylation of the enzyme and its expressed hydroxylase activity is in sharp contrast to the conclusion reached from our studies of the dephosphorylation of pure phenylalanine hydroxylase, which, as discussed previously, demonstrated that the hydroxylase from which all of the previously incorporated 32Pihad been removed by the action of phenylalanine hydroxylase phosphatase has about one-third the hydroxylase activity of the fully phosphorylated enzyme (see Fig. 4). It should be emphasized that the actual data that were extrapolated in the study of Fisher and Pogson (57) were limited to the measured increase in phenylalanine hydroxylase activity where the amount of 32Piincorporated was increased from 0.29 to 0.56 mol/mol of subunit. The discrepancy between results obtained with the pure hydroxylase and those obtained with hepatocytes (57) raises the possibility that the extrapolation used in the latter study may not be valid. It is also important to note that the range of phosphorylation in which a correlation between phosphorylation and hydroxylase activity was observed is close to the range in which the hydroxylase activity of pure enzyme correlates more closely with its level of phosphorylation (see Fig. 4). The puzzling finding that glucagon, when added together with 32Pi, actually inhibits the rate of phosphorylation of the hydroxylase in hepatocytes (57) could be explained by our observation, previously mentioned, that administration of

230

SEYMOUR KAUFMAN

glucagon to rats leads to a transient inhibition of hepatic phenylalanine hydroxylase phosphatase. If a substantial fraction of the hydroxylase in control hepatocytes were fully phosphorylated, incorporation of 32Pi into the enzyme could be dependent on the prior dephosphorylation of the enzyme, a reaction that might be inhibited by glucagon in hepatocytes as it is in the intact rat. Because glucagon and insulin generally have opposing metabolic effects in the organism, the observation that glucagon can activate phenylalanine hydroxylase in rats (42) led to the expectation that a relative lack of insulin, as in diabetes, might affect the activity of phenylalanine hydroxylase in the same way as does excess glucagon (i.e., it might lead to activation of the hydroxylase). In this regard, the activity of phenylalanine hydroxylase measured in situ in hepatocytes from rats that had been made diabetic by streptozotocin injections has been reported to be about twice as high as it is in hepatocytes from control rats. The addition of glucagon to the hepatocytes from diabetic rats stimulated this elevated activity another 50% (55). These results were confirmed in studies in which phenylalanine hydroxylase activity was determined in liver extracts from diabetic rats. A 2-fold increase in the BH,-dependent activity and a smaller (32%) increase in the DMPH,-dependent activity was observed (58).Although it is likely that the activation of the hydroxylase seen in diabetic rats was the result of a higher level of phosphorylation, this point was not examined in either of the above studies. Donlon and Beirne (58)did show, however, that the pattern of elution of the enzyme from calcium phosphate-cellulose columns was consistent with the enzyme being more highly phosphorylated in the diabetic state. Both the higher BH,-dependent activity and the altered chromatographic patterns were reversed or prevented by the administration of insulin to the diabetic animals. We have obtained similar results with diabetic rats. As can be seen in Table I, in our experiments the BH,-dependent hydroxylase activity was increased almost 4-fold over control values, whereas the 6MPH,-dependent hydroxylase activity was increased about 30%. Administration of insulin to the diabetic rats depressed the enhanced activity but not down to control levels. As found by Donlon and Beirne (58),the pattern of elution from calcium phosphate-cellulose columns of hydroxylase in liver extracts prepared from diabetic rats indicated that the hydroxylase in these extracts was more highly phosphorylated than in extracts from control rats (50). D. EFFECTOF GLUCACON ON PHENYLALANINE HYDROXYLASE IN VIVO ACTIVITY The demonstration that there is a glucagon-mediated activation of phenylalanine hydroxylase that is manifest in liver extracts prepared from glucagontreated rats (42) made it likely, but did not prove, that this mode of activation

23 1

8. AROMATIC AMINO ACID HYDROXYLASES TABLE I EFFECTOF INSULINTREATMENT ON BH,DEPENDENT PHENYLALANINE HYDROXYLASE ACTIVITY IN EXTRACTSFROM STREPTOZOTOCIN DIABETIC RATsQJ’ Specific activity (nmol tyrosine 30 min/mg) Treatment -Insulin +Insulin Control

6MPH4

BH4

1071 t 173 1190 f 59 825 2 134

56.0 t 11.8 21.3 t 3.5 15.9 t 0.3

aWhere indicated, approximately 2 units of insulin (or buffer) were given subcutaneously. Animals were killed 24 h later, liver extracts were prepared and BH4- and 6MPH4-dependent phenylalanine hydroxylase activities were determined. bn = 4; values are average t S D for assays with 4 animals.

would be expressed in vivo. The possibility remained, however, that the activation might be blunted or not expressed at all in the whole rat. In order to explore the question of the in vivo expression of the activated hydroxylase, a method for assaying phenylalanine hydroxylase in vivo was needed. Furthermore, because of the marked 10- to 30-fold activation of the enzyme by its substrate, phenylalanine, it was important to be able to assay the hydroxylase at, or near, basal phenylalanine concentrations so that phenylalanine activation could be minimized. For this purpose, we used a constant perfusion technique in which rats are perfused intravenously with phenylalanine until a new, higher, steady-state level of blood phenylalanine is approached. To prevent full activation of the enzyme by phenylalanine, a perfusion rate is selected so that the new steady-state level of phenylalanine is less than three times the basal (i.e., preinfusion) level. If glucagon were indeed capable of activating phenylalanine hydroxylase in vivo, its administration to the rats that were being perfused with phenylalanine would be expected to lead to a decrease in serum levels of phenylalanine, the decrease being a function of the new higher rate of phenylalanine hydroxylation. As can be seen in Fig. 6 , the intravenous injection of glucagon (dose, 2.0 mg/kg) led to a rapid decrease in serum phenylalanine levels, the maximum effect being a 34% decrease (p < 0.001) about 9 min after the glucagon had been given. Concomitant with this decrease in phenylalanine levels, there was a comparable glucagon-mediated increase in serum tyrosine levels (50). The latter

232

SEYMOUR KAUFMAN

0

10

20 TIME (min)

FIG. 6. The effect of intravenous glucagon injection (2 mgikg) on serum phenylalanine and tyrosine concentrations. R.ats were perfused intravenously at a constant rate with phenylalanine ( 1 mg/kg/min). At various times, samples of arterial blood were collected. After deproteinization, phenylalanine and tyrosine were determined by automated amino acid analysis.

effect strengthens the conclusion that glucagon stimulated phenylalanine hydroxylation rather than another enzymic reaction that led to the net disposal of phenylalanine, such as transamination. The conclusion that this in vivo glucagon effect reflects an activation of phenylalanine hydroxylase was strongly supported by results of experiments similar to the one illustrated in Fig. 6 in which a bolus of [14C]phenylalanine was injected 5 min after the glucagon had been administered. Samples of blood were withdrawn at 1 min intervals, the phenylalanine and tyrosine isolated on the amino acid analyzer and their specific radioactivities determined. It was found that I min after the [ 14C]phenylalaninehad been injected, the specific radioactivity of serum tyrosine was 75% higher in glucagon-treated rats than in control rats that had not received glucagon. The specific radioactivity of the tyrosine remained elevated for several minutes (50). These results indicate that glucagon stimulated the rate of phenylalanine hydroxylation in vivo by at least 75%.

8.

AROMATIC AMINO ACID HYDROXYLASES

233

It is also possible to estimate the magnitude of the glucagon stimulation of phenylalanine metabolism from the data in Fig. 6. On the assumption that the phenylalanine that is injected at a constant rate distributes rapidly and evenly in all of the major tissue compartments of the rat, the concentration of phenylalanine in serum would, in the absence of any reaction that leads to the metabolism of phenylalanine, increase linearly during the perfusion period. Any negative deviation from this expected linear increase, such as that shown in Fig. 6, would most likely reflect the net metabolic disposition of the infused phenylalanine, which, at these levels of phenylalanine, would mainly be due to its conversion to tyrosine (22). On the basis of these assumptions, it can be estimated that during the 10-min period following its injection, glucagon stimulated the rate of phenylalanine hydroxylation by 75%, a value that is in excellent agreement with the previously discussed stimulation by glucagon of the rate of conversion of [14C]phenylalanine to [14C]tyrosine. It should be noted that the lower serum levels of phenylalanine seen in the presence of glucagon (Fig. 6) are not likely to be due solely to a glucagon-mediated acceleration of entry of phenylalanine into a major tissue compartment. In that case, tyrosine levels would have been expected to change in parallel with those of phenylalanine, whereas there was a glucagon-mediated increase in serum tyrosine concentrations (see Fig. 6). Although the two ways of estimating the glucagon stimulation of phenylalanine hydroxylase are in good agreement, and together strongly support the conclusion that glucagon is capable of activating phenylalanine hydroxylase in the whole rat with at least part of this higher activity being expressed in vivo, the activation in vivo is much less than the 3- to 4-fold activation that can be demonstrated in liver extracts prepared from glucagon-treated rats (42). In an attempt to explain this discrepancy, we have studied the expression of the activated state of purified phosphorylated phenylalanine hydroxylase in v i m under conditions that mimic more closely those of the enzyme within the hepatocyte, rather than under our standard assay conditions for measuring activation of the hydroxylase by phosphorylation (25"C, in the presence of approximately saturating concentrations of BH, and phenylalanine, 20 and 1 mM, respectively). As can be seen in Table 11, at 25°C with saturating concentrations of BH, and phenylalanine, the activity of the fully phosphorylated enzyme is more than five times greater than that of the native enzyme. Under conditions that approximate those attained during the perfusion experiment-namely, 37"C, 0.2 mM phenylalanine (see Fig. 6) and 6 p.M BH,, the latter value being estimated from the reported concentration of biopterin in rat liver (59)-the activity of the phosphorylated enzyme is only 2.2 times greater than that of the native enzyme. This latter value is only slightly greater than the glucagon-mediated stimulation of the in vivo hydroxylase activity. The results shown in Table I1 identified three variables that have marked

234

SEYMOUR KAUFMAN TABLE I1 EFFECT OF BH4 AND PHENYLALANINE CONCENTRATIONS ON THE CAPACITY OF PHOSPHORYLATED PHENYLALANINE HYDROXYLASE TO EXPRESSACTIVATION^ Phenylalanine (mM)

Temperature

BH4 (W)'

0.08 (Basal)

0.2 (Infusion)

1.0 (Saturation)

25OC

2 6 36 2 6 36

5.1 3.2 2.5

2.3 3.7 3.7 1.2 2.2 3.2

1.7 3.0

37°C

1.1 3.9 4.8

5.4

1.1 1.2 1.5

OValues are expressed as the specific activity of the fully phosphorylated enzyme divided by the specific activity of the native enzyme. 'The different BH4 concentrations were selected because they represent the approximate K,, value of the hydroxylase for BH4 (2 W), the approximate rat liver concentration of BH4 (6 M ) ,and a saturating concentration of BH4 (36 @).

effects on the extent of activation of phenylalanine hydroxylase by phosphorylation. Except for the values measured at 25°C at 0.08 mM phenylalanine, there is less activation of the phosphorylated enzyme at low BH, concentration and less activation at 37°C than at 25°C. Furthermore, at 37"C, at all concentrations of BH, that were tested, there was less activation at saturating (1 .O mM) concentrations of phenylalanine. The lower degree of phosphorylation-mediated activation that is expressed at the higher temperatures and at higher phenylalanine concentrations probably reflect the fact that the nonphosphorylated enzyme is activated by high phenylalanine concentrations (7, 17-21) and by exposure to elevated temperatures, at least up to 37" (35). The results shown in Table I1 differ in some important respects from those reported for phenylalanine hydroxylase in hepatocytes, where it was found that glucagon stimulated in situ phenylalanine hydroxylase activity about twofold at basal phenylalanine concentrations (about 0.05 mM), but had little effect (1015% stimulation) at phenylalanine concentrations greater than 0.25 mM (55).By contrast, the activity of the phosphorylated pure enzyme is 2.2-fold higher than that of the control enzyme under similar conditions (37°C 0.2 mM phenylalanine, 6 pkl BH,) (Table 11). Since the magnitude of this phosphorylationinduced activation of the pure enzyme is similar to the 75% stimulation of phenylalanine hydroxylase by glucagon in the intact rat, it is clear that the interrelationship between phenylalanine and glucagon activation of the hydroxylase in hepatocytes (55) is not the same as it is either in the intact rat or with the pure enzyme. The reason(s) for these differences are not known.

235

8. AROMATIC AMINO ACID HYDROXYLASES

E. EFFECTOF LIGANDSON PHOSPHORYLATION OF PHENYLALANINE HYDROXYLASE Phenylalanine and BH, have been found to have opposite effects on the rate of phosphorylation and activation of pure rat liver phenylalanine hydroxylase: physiological concentrations of the naturally occurring 6R diastereoisomer of BH, (6 to 8 pM) markedly inhibit (-80%) phosphorylation and activation, whereas 200 pM L-phenylalanine stimulates both processes to a modest extent. Perhaps of greater physiological significance, phenylalanine can overcome completely the inhibition caused by BH, (60). These effects were observed when the phosphorylation-activation of the hydroxylase was catalyzed by either bovine heart CAMP-dependent protein kinase or rat liver protein kinase catalytic subunit. The inhibition of phosphorylation of the hydroxylase by BH, shows remarkable specificity. As can be seen in Table 111, the unnatural 6s diastereoisomer of BH, is much less active than is the 6R diastereoisomer and relatively large concentrations of either 6MPH, or DMPH, do not inhibit (60). Although the quantitative aspects were somewhat different, Doskeland ef al. (37), reported effects of BH, and L-phenylalanine on the rate of phosphorylation of rat liver phenylalanine hydroxylase that were essentially the same as those reported earlier by Phillips and Kaufman (60). They found, for example, a greater stimulation by phenylalanine (maximum of twofold stimulation at 0.5 mM) and less inhibition by BH, (50% maximum inhibition at 0.03 mM). As far as this last finding is concerned, it should be noted that Doskeland er al. (37) used a mixture of the two diastereoisomers of BH, (70% 6R-BH4, 30% 6S-BH4). It is possible that the presence of the 6s isomer decreased the inhibition by the 6R isomer (see Table 111). With respect to the effect of L-phenylalanine, they observed a 50% stimulation at approximately basal serum phenylalanine concentrations (0.05 mM), lending support to the idea that this may be a physiologically significant effect. They confirmed the previous finding (35)that the phosphorylated form of phenylalanine hydroxylase required less phenylalanine to be actiTABLE I11 SPECIFICITY OF THE INHIBITION OF PHENYLALANINE HYDROXYLASE PHOSPHORYLATION B Y TETRAHYDROFTERINS Mol of phosphate/ subunit/ 10 min

Relative rate

Addition None 8.6 w (6R)-BH4) 8.6 fl (6S)-BH4) 150 6MPH4 200 w DMPH4

0.096 0.024 0.060 0.091 0.096

100.0 24.9 62.4 95.0 99.8

(%)

236

SEYMOUR KAUFMAN

51 t6 .1t8 FIG. 7. Scheme showing the effects of phenylalanine and BH4 on the interconversion of various forms of phenylalanine hydroxylase.

vated than the nonphosphorylated form, values of 29 and 51 pkf being required to obtain half-maximal activation of the phosphorylated and nonphosphorylated forms, respectively. These results reported by Phillips and Kaufman (60) and D@skelandet al. (37) describing the effects of phenylalanine and BH, on the rate of phosphorylation of phenylalanine hydroxylase are consistent with numerous proposals (7, 17-19, 27, 29, 30, 33, 61-63) that phenylalanine hydroxylase exists as an equilibrium mixture of high-activity and low-activity conformations. This type of proposal is illustrated schematically in Fig. 7 where E is a conformation of the hydroxylase that is inactive or has very low activity, E’ is an active conformation, and E-P and E’-P are the phosphorylated forms, respectively, of the low-activity and the high-activity conformations of the enzyme (for simplicity, these forms will be referred to as the inactive and active forms). The active conformation can be stabilized by the binding of phenylalanine to a regulatory site, whereas the inactive form can be stabilized by the binding of BH, in the absence of, or prior to, phenylalanine binding. The inhibition of phosphorylation in the presence of BH, and the stimulation in the presence of phenylalanine implies that the active enzyme, E’, is a better substrate for phosphorylation than is the inactive enzyme, E (i.e., reaction 7 is faster than reaction 5). The physical basis of this inhibition by BH, is unclear at present. Whatever for the phosphorylation of the basis might be, it has been found that V,,,IK,,, phenylalanine hydroxylase is reduced by a factor of 5 in the presence of BH, (60). Since V,,,IK, represents the apparent second-order rate constant for association of enzyme and substrate to form the prodctive E*S complex (649, these results suggest that phenylalanine hydroxylase, when complexed with 6R-BH4, associates with the protein kinase catalytic subunit at a rate one-fifth that of free phenylalanine hydroxylase.

8. AROMATIC AMINO ACID HYDROXYLASES

237

As shown, the scheme assumes that both E and E‘ are substrates for the protein kinase (i.e., it assumes that both reactions 5 and 7 occur). It is conceivable, however, that the rate of reaction 5 is exceedingly slow so that essentially all of the enzyme is phosphorylated via the sequence of reactions 1 and 7 (i.e., that E-P is not a significant intermediate in the phosphorylation pathway). In that case, it would be necessary to assume that even in the absence of added phenylalanine, there is a small amount of E’ in equilibrium with E and that it is this activated conformation of the enzyme that is the substrate for the kinase. This last assumption is necessary because the results of Phillips and Kaufman (60),as well as those of Doskeland et al. (37) clearly show that ligands such as phenylalanine and BH, affect the rate but not the extent of phosphorylation. There is, however, evidence in favor of the existence of E-P as a discreet species of the enzyme. Some of the evidence on this point comes from studies of the effects of BH, and phenylalanine on the rate of dephosphorylation of phosphorylated phenylalanine hydroxylase catalyzed by ‘‘phenylalanine hydroxylase phosphatase.” It has been found that, just as it does with the phosphorylation of phenylalanine hydroxylase catalyzed by CAMP-dependent kinase, 6R-BH4 also inhibits the dephosphorylation reaction. In contrast to the results with phosphorylation, however, where phenylalanine stimulates, phenylalanine was also found to inhibit the phosphatase-catalyzed release of phosphate from the enzyme. Neither BH, nor phenylalanine inhibited the dephosphorylation of [32P]histone. Thus, these effects are due to BH, and phenylalanine binding to phenylalanine hydroxylase. Furthermore, the inhibitory effect of BH, is specific for the naturally occurring (6R)-BH,; (6s)-BH,, 6MPH,, and DMPH, have little or no effect (60). These findings suggest that free phosphorylated enzyme is a better substrate for the phosphatase than either the phenylalanine-activated enzyme or the BH,-hydroxylase complex (i.e., phosphatase-catalyzed reaction 6 is faster than reaction 8). Similar results were obtained when phosphorylated phenylalanine hydroxylase was proteolyzed by a-chymotrypsin (65). The above results have implications regarding the mechanism of activation of phenylalanine hydroxylase by phosphorylation. Clearly, the inhibition of both the dephosphorylation and proteolysis of phosphorylated phenylalanine hydroxylase by both activating ligands (i.e., phenylalanine) and deactivators (i.e., BH,) cannot be explained simply as a result of an alteration of the equilibrium between active (E’) and inactive (E) conformations (Fig. 7). Rather, these data imply that the phosphorylated enzyme must exhibit a conformation that is intrinsically more susceptible to proteolysis and dephosphorylation, distinct from either the fully activated or inactive native enzyme. It would be desirable to be able to assign maximum BH,-dependent hydroxylase activities to each of the four species of the enzyme shown in Fig. 7. Unfortunately, there is still a considerable amount of disagreement on this point. One of the sharper disagreements concerns the activity of the unactivated

238

SEYMOUR KAUFMAN

enzyme, E, compared to that of the phosphorylated species, E-P. Shiman et al. (35) have reported that phosphorylated phenylalanine hydroxylase that has not been activated by phenylalanine has only 5 to 7% of the activity of the phosphorylated enzyme that has been fully activated by preincubation with phenylalanine. Based on these findings, they concluded that “in the absence of activation, the phosphorylated enzyme has little or no catalytic activity. According to these workers, the only role that phosphorylation plays in the regulation of hydroxylase activity is to facilitate activation of the hydroxylase by phenylalanine. Dgskeland et al. (37), however, could not confirm the finding of Shiman et al. (35) that the phosphorylated enzyme is essentially inactive. By contrast, they found that at the lowest concentration of phenylalanine that they examined, well below 0.01 mM, the phosphorylated enzyme had about 50% of the activity of the phosphorylated enzyme that had been fully activated by preincubation with phenylalanine. However, it should be mentioned that the extent of activation of the hydroxylase by phosphorylation reported by Dgskeland et al. (37), about 20-fold, is considerably greater than the 3- to 4-fold activation that Kaufman and colleagues found (33, 42) (see Table 11). Finally, it should be evident that any attempt to compare the activities of the unactivated enzyme, E, and the phosphorylated enzyme, E-P, may be complicated by many variables that can affect the hydroxylase activity of the phosphorylated enzyme (see Table 11). Despite the fact that there are some differences in the estimates of the intrinsic BH,-dependent hydroxylase activity of the unactivated enzyme, E, the extent of activation of phenylalanine hydroxylase by phenylalanine has consistently been found to be large, usually around 25- to 30-fold (21, 37). From the preceding discussion, it is clear, however, that there is no consensus on the relative activating effects of phosphorylation and phenylalanine (i.e., the relative activities of E-P and E’, Fig. 7). To summarize the relevant findings on this point, the two most divergent views are those of Shiman et al. (35),who reported that E-P is esentially inactive, and those of Dgskeland et al. (37), who found that the phosphorylated enzyme, E-P, has about the same activity as the phenylalanineactivated species, E’. Our findings fall between these extremes; we reported a 28-fold activation by preincubation of the native enzyme with phenylalanine (21) compared to a 3- to 4-fold activation by phosphorylation (33, 42). According to our values, E’ would have seven to nine times higher activity than E-P. Similarly, these divergent findings complicate any attempt to compare the activities of the remaining two forms of the enzyme, E-P and E’-P. As previously mentioned, Shiman et al. (35) claim that E-P is essentially devoid of activity, whereas Dgskeland et al. (37) reported that the activity of the phosphorylated enzyme can be increased only by about 1.7-fold after activation by phenylalanine (E’-P). ”

8.

AROMATIC AMINO ACID HYDROXYLASES

239

It seems likely that the opposing effects of BH, and phenylalanine on both the direct activation of the enzyme and on the kinase-mediated activation is one of the dominant features of the physiological regulation of hepatic phenylalanine hydroxylase. In the case of the kinase reaction, the effect of BH, could be to limit the extent of phosphorylation, and thus activation, when the levels of hepatic phenylalanine are very low and high hydroxylase activity is not required. This would be true under basal conditions. Higher concentrations of phenylalanine would then be able to overcome the inhibitory effect of BH,, allowing an increase in the extent of phosphorylation and activation when the organism needs this higher hydroxylase activity in order to catabolize the excess phenylalanine. This inhibitor effect of BH, would then serve as a “safety valve” to protect against depletion of the organism’s pool of phenylalanine below essential levels. These effects of phenylalanine and BH, on the kinase reaction are relevant to the interpretation of the report of Shiman et al. (35) that the phosphorylated hydroxylase has little or no activity without activation by phenylalanine. These workers concluded that this restraint on the enzyme’s activity ‘‘makes physiological sense,” implying that if phosphorylation activated the enzyme directly, “the effects could be very serious for the animal,” presumably because the organism’s supply of phenylalanine might be quickly depleted by the hydroxylaseinitiated catabolism of this amino acid. The results of Phillips and Kaufman (60) on the ability of phenylalanine to overcome the inhibition by BH, of the kinase-catalyzed activation and phosphorylation of the hydroxylase indicate that, contrary to the view expressed by Shiman et al. (33,there probably would be no serious consequences to the animal if phosphorylation of the hydroxylase activated it directly since their findings suggest that the enzyme will not be rapidly activated by phosphorylation in vivo in the absence of phenylalanine. The suggestion that direct activation of phenylalanine hydroxylase by phosphorylation would be bad for the organism also overlooks another built-in safety factor, that is, that the rate of the hydroxylation reaction catalyzed by both the phosphorylated and the nonphosphorylated enzyme decreases as a function of the square of the phenylalanine concentration (33).This relationship, together with the facilitating effect of phenylalanine on the phosphorylation reaction, would ensure that the hydroxylase cannot deplete the animal’s supply of phenylalanine. AND DRUGSO N THE F. EFFECTOF OTHER HORMONES PHOSPHORYLATION OF PHENYLALANINE HYDROXYLASE

The administration of epinephrine to rats has been shown to stimulate the hepatic BH,-dependent phenylalanine hydroxylase activity (50). The maximum stimulation that has been observed, about two-fold (at a dose of 0.75 to 1.0

240

SEYMOUR KAUFMAN

-

Epinephrine (SC

Theophyllhe (IP) Methyl lsobutyl Xanthine (IPk

4.0 -

3.0 -

2.01.0

-

~..;-4 I

I

I

-Control

-Epinephrine

-Theophylline

fn

-

-

-

x

-

-

-

t

c 2 IV 4:

I

I

~

I-

t

I m

1

1

1

1

1

,

1

1

1

1

1

1

1

1

1

1

1

1

FIG. 8. The effects of the administration to rats of epinephrine, theophylline, and methyl isobutyl xanthine on hepatic BH4-dependent phenylalanine hydroxylase activity (upper panel) and on the elution pattern of phenylalanine hydroxylase activity from calcium phosphate-cellulose columns (lower panel). SC, subcutaneous injection; IP, intrapentoneal injection.

mg/kg body weight, injected subcutaneously), is clearly less than the typical four-fold glucagon induced activation (42).As can be seen in Fig. 8, activation by epinephrine is associated with a shift in the enzyme’s elution pattern from cellulose-calcium phosphate gels to one that is characteristic of the more highly phosphorylated forms of the enzyme. In accord with the lesser degree of activation that is elicited by epinephrine, however, the shift toward form IV is not as complete as it is when the enzyme is activated by either glucagon injection ( 4 2 ) , aminophylline (44), or theophylline administration. With all of these agents, activation is associated with an almost complete conversion of the various forms of the enzyme to form IV (Figs. 5, 8). The reason for the less complete activation and phosphorylation induced by epinephrine is not known. An increase of the BH,-dependent hydroxylase activity has been observed in rat hepatocytes treated with norepinephrine. The stimulation was about 50% at M norepinephrine but, in contrast to our results with the intact rat, the activation did not appear to be less than, and may have exceeded, that observed with glucagon (about 30% at l O - * O M glucagon) (54). The stimulation by norepinephrine was blocked by the p adrenergic antagonist, propranalol, but not by the ci blocking agent, ergocryptine. Abita et al. (54) concluded, therefore, that this effect of norepinephrine in hepatocytes is not mediated by an ci adre-

24 1

8 . AROMATIC AMINO ACID HYDROXYLASES

nergic mechanism, but instead involves a P adrenergic-mediated process. These workers also observed a small stimulation of phenylalanine hydroxylase activity by l o p 5 M concentrations of the calcium ionophore, A23187 (54). Additional evidence implicating a Ca2 -sensitive protein kinase in the phosphorylation of hepatic phenylalanine hydroxylase has been presented by Garrison and Wagner (56). Using isolated rat hepatocytes, they showed that 10 p l 4 norepinephrine increased the level of phosphorylation of phenylalanine hydroxylase by 1.8-fold. Since this effect was seen in hepatocytes that had been pretreated with propranolol to block any rise in CAMP levels that might have been mediated by the hepatic P-adrenergic receptor, these workers concluded that a CAMP-independent protein kinase was involved. They also showed that vasopressin and angiotensin 11, hormones that are believed to utilize Ca2 -linked pathways, stimulated the phosphorylation of phenylalanine hydroxylase to about the same extent as did norepinephrine. It is of interest that the 1.8-fold stimulation of phosphorylation that was induced by norepinephrine was about half the stimulation elicited by glucagon. In this study, therefore, the relative effects of these two hormones on the level of phosphorylation of phenylalanine hydroxylase in hepatocytes were consistent with their relative effectiveness in stimulating BH,-dependent hydroxylase activity in vivo since, as previously discussed, administration of glucagon to rats has been found to increase phenylalanine hydroxylase about fourfold (42),whereas epinephrine has been found to activate about twofold (see Fig. 8). Not all of these results have been replicated. Fisher et al. (66),for instance, failed to detect any increase in phosphorylation of phenylalanine hydroxylase or any stimulation of hydroxylase activity, measured in situ, in rat hepatocytes that were exposed to vasopressin or to the P agonist, isoprenaline. They were, however, able to confirm the previous observations made in the intact rat (50) (see Fig. 8) and in rat hepatocytes (54, 56) that norepinephrine can stimulate the activity and the state of phosphorylation of phenylalanine hydroxylase. In addition, they were able to reproduce the finding (54) that the Ca2+ ionophore, A23187, can stimulate the activity of phenylalanine hydroxylase and increase its state of phosphorylation (67). Furthermore, in studies carried out with a-and Padrenergic antogonists, they obtained evidence that in hepatocytes from mature rats, the stimulation of the hydroxylase by norepinephrine and epinephrine is mediated by an a-adrenergic mechanism, whereas in hepatocytes from young rats (80 g), the stimulation is mediated by P-adrenergic mechanisms. This change in pattern of the adrenergic response with age is, in general, consistent with the findings of Blair et al. (68)who showed that glucose output in hepatocytes from mature male rats is mainly regulated through a-adrenergic receptors, whereas it is regulated by both a- and P-adrenergic receptors in juvenile male rats. The kinase that is responsible for the Ca2 -mediated phosphorylation of phe+

+

+

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nylalanine hydroxylase in rat hepatocytes has not been identified with certainty. A potential candidate for this role is calmodulin-dependent glycogen synthase kinase, which has been shown to catalyze the phosphorylation of pure rat liver phenylalanine hydroxylase (69). Although this initial preliminary communication did not report whether phosphorylation by this kinase also activated the hydroxylase, it has been shown that incorporation of about 0.5 mol of phosphate/mol of hydroxylase subunit catalyzed by a calmodulin, Ca2 -dependent kinase is associated with a threefold increase in BH,-dependent hydroxylase activity (70). It was also shown that, in contrast to the stimulation by phenylalanine of phosphory lation of the hydroxylase by CAMP-dependent protein kinase (37, 60), previously discussed, phosphorylation by the calmodulin, Ca2 -dependent kinase is inhibited by phenylalanine (about 50% inhibition at 0.1 mM phenylalanine) (70). Doskeland et al. (70) presented strong evidence that the site of phosphorylation of the hydroxylase catalyzed by this kinase is the same as the one catalyzed by the CAMP-dependent kinase. Despite this finding, the physiological significance of the calmodulin-dependent phosphorylation of the hydroxylase is obscure, since this reaction would be inhibited under just those conditions when the phosphorylation-mediated activation of the enzyme would appear to be most useful, that is, when tissue phenylalanine concentrations are elevated. Another potential enzyme that could be involved in the Ca2+-mediated phosphorylation of the hydroxylase is protein kinase C. The finding, however, that phorbol esters, known to activate this enzyme in platelets (71, 72) have no effect on the state of phosphorylation of phenylalanine hydroxylase provides evidence against the participation of this particular Ca2 -dependent kinase in the phosphorylation of the hydroxylase (67). Other kinases that have been shown to be unable to phosphorylate phenylalanine hydroxylase are liver and muscle phosphorylase kinase (70). Although phenylalanine hydroxylase was shown to be a substrate for cGMP-dependent protein kinase, the rate of the phosphorylation reaction was only about 10% that of the CAMP-dependent protein kinase, making it unlikely that the cGMPdependent enzyme is involved in the in vivo regulation of the hydroxylase (70). +

+

+

G. PHYSIOLOGICAL SIGNIFICANCE OF THE REGULATION OF PHENYLALANINE HYDROXYLASE ACTIVITY BY PHOSPHORYLATION-DEPHOSPHORYLATION There are at least two mechanisms for the acute regulation of the activity of hepatic phenylalanine hydroxylase that appear to operate in vivo, activation by phosphorylation and activation by phenylalanine. As has been discussed briefly, the evidence indicates that these are probably not independent modes of regulation of the enzyme but rather that they act synergistically. Thus, phosphorylation

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(and activation) of the enzyme by CAMP-dependent protein kinase is stimulated by phenylalanine (37, 60) whereas phosphorylation sensitizes the enzyme to activation by phenylalanine (27, 37, 55). One of the consequences of these interlocking control mechanisms is to enhance the responsiveness of the activity of the hydroxylase to alterations in tissue levels of phenylalanine. Since phenylalanine is believed to be a glycogenic (as well as a ketogenic) amino acid (73), it seems likely that activation of phenylalanine hydroxylase might be geared to the needs of the organism for increased gluconeogenesis. Furthermore, in view of the fact that phenylalanine hydroxylase catalyzes the rate-limiting step in the conversion of phenylalanine to gluconeogenic metabolites (74), it was not unexpected that its activity would be tightly regulated. In addition, as has been indicated previously (46, 7 3 , the finding that the enzyme in rat liver can be activated by glucagon-mediated phosphorylation (42)strengthens the likelihood that activation of the hydroxylase might be related to gluconeogenesis. The glucagon activation of phenylalanine hydroxylase suggests that one of the variables that can trigger this regulatory mechanism would involve changes in blood sugar levels; that is, a fall in the concentration of blood sugar would lead to an increased release of glucagon (and decreased release of insulin) from the pancreas that would activate hepatic adenylate cyclase, leading to an increase in hepatic CAMP concentrations, that, in turn, would activate CAMP-dependent protein kinase resulting in the phosphorylation-mediated activation of phenylalanine hydroxylase. This glucagon-mediated activation of the enzyme would probably also involve activation by phenylalanine since, under these conditions, increased amounts of phenylalanine (as well as other gluconeogenic amino acids) would be released into the blood from muscle proteins and taken up by the liver where the phenylalanine would enhance the rate of phosphorylation-mediated activation of the hydroxylase. Another physiological rationale for activating phenylalanine hydroxylase is one that links metabolic events in the liver to those occurring in other organs. It is known, for instance, that when blood phenylalanine levels are elevated 15- to 20fold, as they are in patients with classical phenylketonuria, the disease caused by a genetically determined lack of hepatic phenylalanine hydroxylase, the brain does not develop normally [see Ref. (22) for a review]. The neonatal brain, therefore, is very sensitive to damage by excess phenylalanine. It is also known that sustained hyperphenylalanemia during pregnancy can damage the fetus in utero (76).In view of this sensitivity, it would be beneficial, at least during the fetal and neonatal period, for the organism to have mechanisms that would minimize elevations in blood phenylalanine levels. It seems reasonable to expect that the synergistic effects on phenylalanine hydroxylase of the interaction of phenylalanine-mediated and phosphorylation-mediated activation of the hydroxylase would achieve this result.

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This regulatory process would be triggered by the ingestion of a proteincontaining meal. It is known that following such a meal, pancreatic glucagon secretion is stimulated, resulting in a large rise in blood glucagon levels (77). This increase is believed to be caused by the consequent elevation of the blood concentrations of several amino acids, including phenylalanine, that are potent stimulators of pancreatic glucagon release (78, 79).The elevated blood glucagon levels would, by the same sequence of reactions outlined above, mediate activation of the hydroxylase. One of the consequences of the operation of this regulatory system, which would be set in motion by the postprandial elevations of blood phenylalanine concentrations, would be the accelerated catabolism of the increased amounts of phenylalanine and a more rapid return of blood and tissue phenylalanine levels to the basal range. Each step in the sequence of reactions is supported by experimental data. As previously discussed, it has been shown that the administration of glucagon to rats increases the rate of metabolism of elevated levels of phenylalanine (Fig. 6 ) . It has also been shown that feeding rats a high protein-low carbohydrate diet for a week leads to activation of hepatic phenylalanine hydroxylase and that this activation is probably due to increased phosphorylation of the enzyme (58). Interpretation of this last result is somewhat uncertain, however, because the effects of the high protein diet were not studied as a single variable but only in conjunction with a low carbohydrate diet. It is, therefore, possible that it was the low carbohydrate diet that led to activation of the hydroxylase. It should also be noted that the sequence of reactions describes a possible mechansim for the acute in vivo activation of the hydroxylase following ingestion of a protein meal, whereas the activation observed by Donlon and Beirne (58) was seen after feeding rats the high protein diet for a week. In this regard, there is evidence that prolonged protein feeding (for 9 to 10 days) stimulates glucagon secretion by mechanisms that may be unrelated to the rise of plasma amino acids that follow protein ingestion (80). The ability of phenylalanine to stimulate the release of glucagon from the pancreas may have complicated the interpretation of data published by Shiman et al. (35).These workers reported that the administration to rats of a large dose of phenylalanine led to an 18- to a 28-fold activation of phenylalanine hydroxylase in liver extracts from the treated animals compared to the activity in salinetreated controls. They assumed that this dramatic activation reflected the direct in vivo activation of the enzyme by the injected phenylalanine. They did not rule out the possibility that part of this activation may have been due to phenylalanine-mediated glucagon release and the consequent activation of the enzyme by phosphorylation. It is possible, in fact, that the phenylalanine-stimulated release of pancreatic glucagon would be much more effective in mediating the phosphorylation of the hydroxylase than would the intraperitoneal injection of glucagon, itself. We have found, for instance, that the amount of glucagon

8.

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245

FIG. 9. Scheme showing the effects of glucagon, BH4, and phenylalanine on the interconversion of the various forms of phenylalanine hydroxylase.

needed to elicit half-maximal activation of the hydroxylase when the glucagon is injected intraperitoneally, as it was in the experiments reported by Shiman et al. (35),is more than 10 times greater than when it is administered intravenously (50). It is noteworthy that Shiman et al. (35)showed that valine, an amino acid that does not activate phenylalanine hydroxylase in vitro, did not activate the enzyme when it was injected into rats. Although they cited this negative result with valine to support their conclusion that the administered phenylalanine had activated the hydroxylase directly, the result with valine cannot discriminate between a direct and an indirect, glucagon-mediated activation because valine, in contrast to phenylalanine, does not stimulate the pancreatic release of glucagon (78). The interaction between phenylalanine-mediated and phosphorylation-mediated activation of hepatic phenylalanine hydroxylase is summarized in Fig. 9 which is an elaboration of Fig. 7. One of the salient features of the regulation of this enzyme’s activity is the multiple roles that phenylalanine appears to play. 1. Phenylalanine can activate the hydroxylase directly, in part by overcoming

inhibition by BH,. 2. Phenylalanine can stimulate the phosphorylation and activation of the hydroxylase by CAMP-dependent protein kinase, also in part by overcoming the BH,-mediated inhibition of the reaction. 3. Phenylalanine can elicit the release of glucagon from the pancreas, setting in motion activation of the hydroxylase by CAMP-dependent protein kinase.

246

SEYMOUR KAUFMAN TABLE IV PROBABLE EFFECTOF DIFFERENT TYPESOF DIETSON THE MODEOF ACTIVATION OF HEPATICPHENYLALANINE HYDROXYLASE

Type of meal consumed

Expected effect on blood glucagon levels

Carbohydrate, protein Protein without carbohydrate

Little or no change Increase

Fasting or starvation

Increase

Expected predominant mechanism of phenylalanine hydroxylase activation Phenylalanine-mediated Phenylalanine-mediated and glucagon-mediated Glucagon-mediated

From the points that have been previously discussed it seems likely that the composition of the diet consumed by the organism determines the mechanism of activation of hepatic phenylalanine hydroxylase. Table IV outlines the results expected from the consumption of several different types of diet. The table also serves to underline the fact that even though the last two dietary situations (high protein without carbohydrate, on the one hand, and fasting on the other) would both be expected to lead to increased secretion of glucagon, activation of the hydroxylase would probably fulfill different needs of the organism in these two situations (i.e., in the former one, it would be useful for the disposal of excess phenylalanine as well as for increased gluconeogenesis, whereas in the latter it would be useful mainly for increased gluconeogenesis). A final point to be made about the summary shown in Table IV is that there appear to be dietary situations where the acute activation of phenylalanine hydroxylase is mediated by glucagon-stimulated phosphorylation. In this regard, it may be recalled that Shiman et al. (35)reported that the phosphorylated form of the enzyme is essentially inactive unless it is first activated by phenylalanine, implying that the sole mechanism by which phosphorylation can increase the activity of the enzyme is by facilitating its activation by phenylalanine. It has also been pointed out that although there is agreement that phosphorylation does decrease the amount of phenylalanine needed to activate, there is sharp disagreement about the activity of the phosphorylated enzyme. Shiman et al. (35)have also implied that the extent of activation of the enzyme by glucagon-mediated phosphorylation is too small to be of physiological importance. This view seemed to be supported by their finding that after glucagon administration to rats, the phosphorylated enzyme is at most 5-7% activated, whereas after a very large load of phenylalanine (about 540 mg/kg) the percent of activated enzyme was increased to 40% of maximum. It seems likely, however, that the comparison of the activation of the hydroxylase that is elicited by glucagon to the maximum activation that can be achieved

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by a large dose of phenylalanine, (a part of which may actually be, as already pointed out, due to phenylalanine-stimulated glucagon release) is probably of little physiological relevance. There is no evidence to suggest that the physiological regulation of the hydroxylase ever involves anything more than a small fraction of its maximum activity. This limit probably reflects the fact that blood phenylalanine levels normally fluctuate within a relatively narrow range. In man, for example, even after phenylalanine intake in the form of protein, has been doubled (to 80 mg/kg/day) serum phenylalanine levels barely rise (81). Under these circumstances, it can be anticipated that activation of the hydroxylase will be small compared to that seen after a huge pharmacological dose of phenylalanine. The glucagon activation must be compared to this small activation by phenylalanine that would likely follow the ingestion of a normal protein-containing meal in order to assess the physiological significance of this hormone’s ability to activate phenylalanine hydroxylase. Is Human Liver Phenylalanine Hydroxylase Regulated by Phosphorylation-Dephosphorylation ? It has been reported that, in sharp contrast with phenylalanine hydroxylase from rat liver, the pure enzyme from fresh human liver cannot be activated by CAMP-dependent protein kinase (82).Furthermore, in the presence of [32P]ATP and the kinase, there was no incorporation of 32Piinto the enzyme. The failure to phosphorylate the hydroxylase could not be explained by the fact that it was already fully phosphorylated, since direct analysis showed that the enzyme contained no detectable protein-bound phosphate. Smith et al. (83) were unable to confirm the finding that human liver phenylalanine hydroxylase cannot be phosphorylated by CAMP-dependent protein kinase. Using the catalytic subunit of the bovine heart kinase, they observed the incorporation of about 0.25 rnol of 32Pi into the enzyme from [32P]ATP. If the enzyme was first treated with alkaline phosphatase, the amount of 32Piincorporated into the hydroxylase was increased to 0.67 mol per mol of hydroxylase subunit, a finding that suggests that their preparation of the human liver enzyme was even more highly phosphorylated than the rat liver enzyme. Smith et al. (83) did, however, confirm the finding of Abita et al. (82) that the BH,-dependent hydroxylase activity was not stimulated by phosphorylation. The explanation for the discrepancies between the results of Abita et al. (82)and those of Smith et al. (83) are not obvious. However, it is possible to explain the failure of Smith et al. (83)to detect any activation of the enzyme resulting from its phosphorylation. The failure may be due to the high state of phosphorylation of their enzyme preparation. As previously mentioned, their finding that an additional 0.42 mol of 32P/mol of hydroxylase could be incorporated by the prior treatment of the enzyme with alkaline phosphatase indicates that their isolated enzyme contained a minimum

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of 0.42 mol of P/mol of hydroxylase. Since they could only incorporate about 0.25 mol 32P/mol of hydroxylase prior to phosphatase treatment, it is even possible that their isolated enzyme contained about 0.75 mol of P/mol of hydroxylase. With fully phosphorylated phenylalanine hydroxylase (i.e., enzyme containing 1 mol P/mol hydroxylase subunit), we have found that there is only a small decrease in BH,-dependent hydroxylase activity on removing 25% of the protein-bound phosphate through the action of rat liver phosphatase. Even removal of 50% of the protein-bound phosphate from the fully phosphorylated enzyme leads to a loss of only 30% of the activity (44) (see Fig. 4). If the human liver enzyme behaves like the rat liver enzyme, therefore, starting with an enzyme that contains between 0.50 and 0.75 mol P/mol of hydroxylase would make it more difficult to detect any increase in activity resulting from the enzyme’s further phosphorylation. The question of whether the activity of human liver phenylalanine hydroxylase is regulated by phosphorylation-dephosphorylation is clearly one of great interest. Despite the findings that suggest that its activity cannot be increased by the action of CAMP-dependent protein kinase, too many uncertainties remain about the interpretation of these findings to regard this question as one that has been settled. We have found that rat kidney phenylalanine hydroxylase resembles the human liver enzyme in that it cannot be activated by the action of CAMPdependent protein kinase (84). It is not known, however, whether this kinase can catalyze the phosphorylation of the kidney enzyme.

111. Tyrosine Hydroxylase A.

INTRODUCTION

The discovery of the essential role of unconjugated pterins in the enzymic hydroxylation of phenylalanine (2, 3 ) and the identification of the naturally occurring hydroxylation cofactor in liver as tetrahydrobiopterin (BH,) ( 4 ) facilitated the subsequent characterization of both tyrosine and tryptophan hydroxylases. With respect to tyrosine hydroxylase, for example, it had been postulated by Blaschko as long ago as 1939 (85) that the first step in the biosynthesis of norepinephrine and epinephrine involved the conversion of tyrosine to 3, 4dihydroxyphenylalanine (dopa), followed by decarboxylation of the dopa to dopamine, which was then hydroxylated in the f3 position of the side chain to form norepinephrine and methylated on the amino group to form epinephrine. Although the evidence in favor of the pathway proposed by Blaschko continued

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to mount, (86-89), and is widely accepted as the only quantitatively important one, the enzyme responsible for the ring hydroxylation reaction has been elusive. In 1964, the enzymic conversion of L-tyrosine to L-dopa was demonstrated in particles isolated from adrenal medulla (13, 90, 91), brain, and other sympathetically innervated tissues (91). The properties of this enzyme clearly distinguished it from tyrosinase, another enzyme that is capable of oxidizing tyrosine to dopa. With the latter enzyme, however, dopa is further oxidized to melanin. An even sharper distinction between these two enzymes is that tyrosine hydroxylase shows an absolute and specific requirement for a tetrahydropterin (13, 90, 91). Dopa, for example, which is capable of stimulating the activity of tyrosinase, is unable to substitute for a tetrahydropterin (13, 90) and, in fact, is a potent inhibitor of tyrosine hydroxylase (91). As far as the pterins are concerned, not only was the naturally occurring pterin cofactor, BH,, shown to be highly active, but significantly, it was demonstrated that BH, could function catalytically (13, 90). Furthermore, the marked stimulation of the conversion of tyrosine to dopa by NADPH and dihydropteridine reductase showed that the tetrahydropterin was utilized during the hydroxylation reaction (13, 90). All of these general characteristics of the tyrosine hydroxylase-catalyzed reaction indicated that the role of the tetrahydropterin in this hydroxylation system would prove to be identical to the one previously established in the phenylalanine hydroxylating system (12, 16), an expectation that was fulfilled by subsequent work. It was not until 1971 that the precise nature of the reaction catalyzed by tyrosine hydroxylase from bovine-adrenal medulla was determined. Studies of the stoichiometry of the reaction in the presence of BH, showed that the reaction proceeded according to Eq. (2). (14). L-Tyrosine + BH4

+ 0 2 + L-dopa + quinonoid-BH2 + H2O

(2)

B. PHYSICAL PROPERTIES Since it is not known whether tyrosine hydroxylase from adrenal medulla has the same structure as the enzyme from brain, the properties of the enzyme from these two tissues must be considered separately. Native rat adrenal tyrosine hydroxylase is a tetramer, M, = 260,000, that is believed to be composed of four identical subunits, M, = 59,000 (92). The native hydroxylase from bovine adrenal medulla has been obtained in pure form. The enzyme from this source is also a tetramer, M , = 280,000, composed of four identical subunits, M, = 60,000 (93). The bovine adrenal enzyme, which shows a great tendency to aggregate, can be partially proteolyzed with either chymotrypsin (14) or trypsin (94) to an active species that no longer shows this tendency. The molecular weight of the trypsin-

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treated enzyme was reported to be 34,000 (95).Interestingly, the essentially pure chymotrypsin-treated enzyme was found to have about the same molecular weight, 33,500-34,700 (96). Because the trypsin-treated bovine adrenal enzyme appeared to have a much smaller molecular weight than that of the native enzyme, Musacchio et al. (95) concluded that the proteolyzed hydroxylase represented only a small fragment of the native enzyme. It seems more likely that the large difference in size between the native and the trypsin-treated enzyme reflects not only a decrease in size of the subunit due to partial proteolysis, but also a change in the oligomeric structure of the proteolyzed enzyme so that it can no longer form dimers and tetramers. In this respect, the behavior of tyrosine hydroxylase resembles that of rat liver phenylalanine hydroxylase. With that enzyme it has been shown that in its native form it exists mainly as tetramers ( M , = 200,000). By contrast, the highly active product of limited chymotrypsin treatment ( M , = 35,000) has lost the ability to form tetramers, although it can still form dimers (29). As is true of most of the other properties of tyrosine hydroxylase, the situation with respect to the size of the brain enzyme appears to be more complicated than that of the adrenal enzyme. In this context, it has been reported (97) that the locus coeruleus of rat brain, a region enriched in cell bodies of noradrenergic neurons, contains a high-molecular-weight ( M , = 200,000) form of the enzyme, whereas in the substantia nigra and the caudate nucleus, the areas enriched in cell bodies and nerve endings, respectively, of dopaminergic neurons, a low-molecular-weight form of the enzyme ( M , = 65,000) is present and is the major form in the latter brain region. Peripheral noradrenergic neurons in the superior cervical gangion contain a form of the enzyme with an intermediate molecular weight ( M , = 130,000). Further complicating the picture is the finding that incubation of homogenates of locus coeruleus with RNase reduced the molecular weight of the hydroxylase from about 200,000 to 150,000 (97). These results indicate that at least part of the apparently high molecular weight of the enzyme in this region is due to its association with RNA, a finding that makes it difficult to decide whether tyrosine hydroxylase molecules of different structure are present in these different regions of the brain. Highly purified tyrosine hydroxylase from bovine striatium has been reported to be composed of two different types of subunits, M , = 60,000 and 62,000 (98). Since Lazar et al. (98) observed a decline in molecular weight from M , = 61,000-62,000 to M , = 50,000 when the purified enzyme was stored at 4°C for 22 h, it is possible that even their observation of two different types of subunit is due to breakdown of the larger form ( M , = 62,000) into the smaller one ( M , = 60,000). With respect to the question of whether tyrosine hydroxylase is composed of different subunits, it should also be mentioned that the enzyme purified from rat

8. AROMATIC AMINO ACID HYDROXYLASES

25 1

pheochromocytoma was found to be a tetramer ( M , = 210,000-222,000) which appeared to be composed of a single subunit, M , = 62,000 (99).

C. ACTIVATIONBY PHOSPHORYLATION As previously mentioned, it is useful to discuss the phosphorylation-mediated activation of the three aromatic amino acid hydroxylases within the context of the roles that these enzymes play in the organism. It may be recalled that in contrast to phenylalanine hydroxylase, which catalyzes the rate-limiting step in the pathway by which phenylalanine is catabolized, neither tyrosine no tryptophan hydroxylases are involved in the catabolism of their respective amino acid substrates. Rather, the reactions that these enzymes catalyze are essential for the synthesis of neurotransmitters. It would be expected, therfore, that the regulation of their activities would be related to neuronal function. The first type of regulation of tyrosine hydroxylase activity that was described was the activation of the enzyme from rat brain by heparin (100). Although this effect was originally believed to be a specific one, subsequent work indicated that it was probably an example of a more general activation of the enzyme by polyanions. It was found, for example, that certain phospholipids, such as phosphatidyl-L-serine (101), as well as other polyanions, including polyacrylic acid (102), polyvinyl sulfuric acid (102) and polyglutamic acid (101, 102) can activate brain tyrosine hydroxylase. The hydroxylase from adrenal tissue can also be activated by heparin (103), phospholipids, (104) and other polyanions (105). Under the assay conditions that were used in these early studies, activation of the hydroxylase was expressed mainly as a decrease in the K , for the pterin cofactor. In the presence of heparin, for example, the K , for DMPH, was decreased from 0.79 mM to 0.14 mM with a small increase in V,,, (100). Activation of the enzyme by phosphatidyl-L-serine was also shown to be characterized by a decrease in the K , for the cofactor, and, in this case, the decrease was demonstrated with the natural cofactor, BH,, whose K , was decreased from 0.22 mM to 0.067 mM. The maximum velocity, however, was not altered (101). The finding that activated tyrosine hydroxylase has a markedly lower K , for BH, was noteworthy because it provided at least a partial solution to a puzzle; that is, how could tyrosine hydroxylase function with a K , for BH, that appeared to be much higher than brain levels of BH,? The partial solution was that the discrepancy between these two values was not nearly as great for the activated as for the unactivated enzyme. Other factors that can narrow this difference still further are discussed later in this section. Although there is little evidence to suggest that activation of tyrosine hydroxylase by polyanions such as heparin and certain phospholipids is of physiological significance, these early studies were nevertheless important because they fore-

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shadowed the changes in properties of the enzyme that probably are involved in its physiological regulation. Specifically, results of these studies made it seem likely that regulation of the activity of this enzyme in vivo would be manifested by an increase in velocity, especially, it should be emphasized, at physiological pH values (101);an increase in the affinity of the enzyme for its pterin cofactor but not for its substrate, tyrosine; and with a concomitant decrease in affinity for the end products in the pathway for catecholamine biosynthesis, dopamine, and norepinephrine. That tyrosine hydroxylase activity is capable of being acutely regulated in vivo has been recognized for 25 to 30 years. In that period, numerous reports were published describing the acceleration of catecholamine synthesis that occurs in adrenal and nerve tissue in response to increased nerve activity (106-113). Since activation of dopamine and norepinephrine-containing tissues usually leads to a parallel increase in both synthesis and release of catecholamines, the tight coupling between these two events permits these tissues to maintain steady-state levels of catecholamines. Furthermore, since, catecholamines are potent inhibitors of tyrosine hydroxylase in vitro (91, 114), the first mechanism proposed to explain this acute activation of tyrosine hydroxylase assumed that activation of the enzyme was due to a decrease in a small pool of catecholamines which normally inhibit tyrosine hydroxylase (110, 115, 116). This kind of mechanism would not necessarily demand that the tyrosine hydroxylase isolated from the stimulated tissue be in an activated state, since in this hypothesis, it is the enzyme’s intracellular environment, rather than the enzyme, itself, that would be altered by the activating stimulus. Not only did subsequent studies fail to provide support for this hypothesis (113, but evidence began to accumulate that indicated that the hydroxylase itself might be altered in stimulated tissues. Furthermore, the evidence raised the possibility that this alteration might involve phosphorylation of the hydroxylase. As will become evident, there is little doubt that the enzyme can be activated when it is exposed to protein-phosphorylating systems. Beyond that point, three related questions need to be examined: 1. Does activation of tyrosine hydroxylase by a phosphorylating system involve the direct phosphorylation of the tyrosine hydroxylase molecule and if so, does it involve a stoichiometric amount of phosphorylation? 2. How is the activated state of the enzyme expressed? 3. Is phosphorylation-dephosphorylation of the hydroxylase involved in the acute physiological regulation of the activity of the enzyme? In 1974, Morgenroth et al. reported that tyrosine hydroxylase in guinea pig vas deferentia was activated after electrical stimulation or potassium depolarization of the sympathetic nerves in this tissue (118).Furthermore, they were able to show that the enzyme in the high-speed supernatant fraction obtained from the

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253

stimulated tissue was still in an activated state (5- to 6-fold decrease in K,,, for DMPH, and for tyrosine, decreased affinity for end product inhibitors, and a 50% increase in V,J, an indication that in response to stimulation the hydroxylase had undergone a rather stable modification that altered its catalytic properties. Since it was also known that both electrical stimulation and depolarizing agents can increase the concentrations of cAMP in brain slices (119, 120), it seemed likely that activation of tyrosine hydroxylase in response to nerve stimulation might be mediated by a CAMP-dependent process. Evidence suggesting that tyrosine hydroxylase could be activated by cAMP was obtained when it was shown that dibutyryl-CAMP added to rat striatal slices stimulated the conversion of [ 14C]tyrosine to [ 14C]dopamine(121). This point was proven directly in experiments with both homogenates and a soluble fraction prepared from rat striatum in which it was shown that added dibutyryl-CAMPstimulated tyrosine hydroxylase and that the stimulation was associated with a decrease in K,,, values for both tyrosine and DMPH, and an increase in Ki for dopamine (122) (i.e., added cAMP produced changes in tyrosine hydroxylase that were similar to those seen after nerve stimulation). That activation of tyrosine hydroxylase by cAMP probably involves phosphorylation of some kind of protein was strongly indicated by the finding that partially purified CAMP-dependent protein kinase added to high-speed supematant fractions from rat hippocampi caused a severalfold increase in tyrosine hydroxylase activity (123). Lloyd and Kaufman, using a highly purified preparation of tyrosine hydroxylase from bovine caudate, independently demonstrated that this hydroxylase could be activated by cAMP and CAMP-dependent protein kinase (124) and showed that the activation was due to a fourfold decrease in the K,,, for the pterin cofactor (either BH, or 6MPH,). These results with BH, demonstrated for the first time that activation of tyrosine hydroxylase by phosphorylation was associated with a decrease in the K,,, of the enzyme for the naturally occurring pterin cofactor. In contrast to the results obtained by Harris et al. (122) with crude rat striatal tyrosine hydroxylase, however, activation of the purified bovine brain enzyme did not lead to a decrease in the K,n for tyrosine (124). Significantly, activation of the purified bovine brain enzyme led to a marked shift in the pH optimum from pH 6.0 to pH 7.4. It was pointed out (124) that all of the changes in kinetic properties of tyrosine hydroxylase after activation by exposure to phosphorylating conditions were essentially the same as those seen when the enzyme was activated by phosphatidyl-L-serine (101). Another indication that these diverse processes lead to similar changes in the enzyme’s kinetic properties was the finding that the two types of activation were not additive (124). Crude bovine adrenal tyrosine hydroxylase was also shown to be activated by exposure to ATP and CAMP-dependent protein kinase, with the activation being

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expressed as a decrease in the apparent K , for 6MPH, and an increase in V,,, (96). In contrast to the results obtained with purified bovine brain tyrosine hydroxylase (124), however, activation of the bovine adrenal enzyme led to only a slight shift in the pH optimum (from pH 6.2 to pH 6.4). The increase in V,,, observed with the adrenal enzyme was also different from results with the rat brain enzyme, where exposure to phosphorylating conditions did not lead to a change in V,,, (122). With respect to differences between adrenal and brain tyrosine hydroxylase, it was found that activation of the adrenal enzyme by phosphatidyl serine resembles activation by phosphorylation (i.e., the K,,, for the pterin cofactor is decreased and V,,, increased) (96). These results provide further evidence for the marked similarity between the effects of phosphorylation and phosphatidyl serine on tyrosine hydroxylase; with brain tyrosine hydroxylase, both modes of activation are expressed by decreases in the K , for the pterin cofactor without an increase in Vmax, whereas with adrenal tyrosine hydroxylase, both of these activations lead to decreases in the K, for the pterin cofactor and increases in V,,, (96, 124). Another similarity between these two types of activation is that they are both lost when the bovine adrenal tyrosine hydroxylase is subjected to limited proteolysis (96). The observation that purified tyrosine hydroxylase could be activated by a CAMP-dependent protein kinase made it seem likely that this activation was the result of a direct phosphorylation of tyrosine hydroxylase. Despite these expectations, the first attempts to show direct phosphorylation of tyrosine hydroxylase from bovine brain (124) or rat brain (125) were unsuccessful. The question of whether tyrosine hydroxylase can be activated by direct phosphorylation remained unanswered even after data were presented indicating that tyrosine hydroxylase is a phosphoprotein. Indeed, the demonstration that tyrosine hydroxylase molecules that are synthesized in organ cultures of adrenal medulla and superior cervical ganglia contain protein-bound phosphate (126, 127), ironically, raised serious doubts about the relationship between phosphorylation of tyrosine hydroxylase and regulation of its activity. These doubts arose because it was estimated that the amount of 32Pithat was incorporated into the hydroxylase in these experiments was about 1 mol of P,/mol enzyme. Based on this finding, the authors concluded that inorganic phosphate is a constitutive part of the enzyme and not a participant in any meaningful regulatory phenomena “and that Pi is a normal part of the enzyme and not one whose addition and removal plays a regulatory role in the action of tyrosine hydroxylase’’ (127). This view was also in accord with their findings that cycloheximide, an inhibitor of protein synthesis, not only decreased the incorporation of [3H]leucine into tyrosine hydroxylase, but it also led to a parallel decrease in incorporation of 32P into the enzyme. These data indicated that 32P was selectively incorporated into newly synthesized tyrosine hydroxylase molecules.

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255

Yamauchi and Fujisawa reported in 1979 (128) that partially purified bovine adrenal tyrosine hydroxylase can be activated in vitro by CAMP-dependent protein kinase. They demonstrated that the activation was accompanied by the incorporation of 32P from [32P]ATPinto the tyrosine hydroxylase molecule. The activation, about 3-fold at pH 6.8, was associated with a shift in the pH optimum of the enzyme from 6.0 to 6.8, confirming the original observation of Lloyd and Kaufman (124). Although the authors stated that there was a good correlation between protein phosphorylation and tyrosine hydroxylase activation, their results showed that the correlation was not always a close one. Thus, activation of the enzyme was considerably more sensitive to the omission of CAMPthan was incorporation of 32P [see Table I1 in Ref. (128)], an indication that each increment in protein-bound phosphate was not associated with an equal increment in hydroxylase activity. As is true of most of the studies on phosphorylation of tyrosine hydroxylase, neither the amount of [32P]phosphateincorporated into the enzyme in vitro, nor the amount (if any) of protein-bound phosphate already present in the isolated enzyme was determined. From the data in Table I1 of their paper, it can be estimated that the amount of 32P incorporated into tyrosine hydroxylase in vitro was much less than stoichiometric, about 0.25 mol/mol of M,.= 60,000-subunit. The effect of phosphorylation on the kinetic constants of the enzyme were not determined. Yamauchi and Fujisawa (129) also showed that tyrosine hydroxylase in bovine adrenal medullary extracts can be activated by an endogenous CAMP-dependent protein kinase and that this activation can be reversed by the action of an endogenous phosphatase. These results showed that the two enzymes needed for phosphorylation-dephosphorylation of tyrosine hydroxylase-a protein kinase and a phosphatase-are present in bovine adrenal medulla. Vulliet et al. (130)carried out similar studies on tyrosine hydroxylase purified from rat pheochromocytoma. They found that highly purified hydroxylase is a substrate for bovine heart CAMP-dependent protein kinase. Activation of the enzyme, which was expressed as a substantial decrease in the K , for 6MPH, (from 0.480 mM to 0.120 mM at pH 6.2) with no change in the K , for tyrosine or increase in Vmax, was accompanied by the incorporation of about 0.7 mol of Pi/60,000 M,-subunit of tyrosine hydroxylase. The amount of protein-bound Pi, if any, present in the nonactivated enzyme was not determined. In contrast to the results of Yamauchi and Fujisawa (128), there was a good correlation between the amount of 32Pi incorporated and the degree of activation during the first 15 min of incubation with the phosphorylating system. Beyond that period, tyrosine hydroxylase activity fell and 32Piincorporation continued, as if the enzyme was being inactivated by further phosphorylation. Vrana et al. (131) reported similar observations that indicated a phosphorylation-mediated loss of hydroxylase activity. Using adrenal chromaffin cells, Meligeni et al. (132) attempted to correlate

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the CAMP-induced stimulation of catecholamine synthesis in situ with activation of tyrosine hydroxylase. One of the burdens that this attempted correlation must bear is the observation that exogenous 8-bromo-CAMP did not stimulate catecholamine synthesis at physiological pH values (pH 7.2-7.4). This observation suggests that the relevance of these results to the physiological regulation of tyrosine hydroxylase may be limited, since it indicates that in adrenal chromaffin cells a CAMP-mediated increase in tyrosine hydroxylase activity, even if it were to occur, would not be expressed. At pH 6.0, exogenous cAMP derivatives stimulated catecholamine synthesis. The stimulation appeared to be largely independent of added tyrosine and DMPH,; that is, although catecholamine synthesis was accelerated by the addition of the cAMP derivatives, the acceleration was somewhat higher at elevated extracellular tyrosine concentrations and somewhat lower at elevated extracellular DMPH, concentrations. These results make it seem unlikely that the observed increase in catecholamine synthesis was being mediated by changes in the K , of tyrosine hydroxylase for either tyrosine or the pterin cofactor. Tyrosine hydroxylase isolated from cells that had been treated with 8-bromo-CAMP had lower K , values for 6MPH,, and higher V,,,, these constants being determined at pH 6.2. These changes in kinetic properties are similar to those found for tyrosine hydroxylase that was activated by exposure to cAMP in extracts of chromaffin cells. When tyrosine hydroxylase in chromaffin cells or in extracts of these cells was activated by addition of 8-bromo-CAMPor CAMP, respectively, in the presence of [32P]ATP, 32P was found to be incorporated into tyrosine hydroxylase, but in neither situation was the stoichiometry of 32P incorporation determined (132). In the experiments carried out with extracts it was shown that incorporation of 32P was associated with activation of tyrosine hydroxylase. The authors concluded that these two changes exhibit “a similar dependence upon cAMP concentrations.” The published data, however, do not support this conclusion. The data show that on going from 1.O p i 4 to 10 pA4 CAMP, there was only a slight (about 20%)increase in the amount of 32P incorporated, but this small increment in 32P incorporation accounted for more than half of the increase in tyrosine hydroxylase activity [see Fig. 9 in Ref. (132)].This discrepancy between the extent of phosphorylation of tyrosine hydroxylase and its activation is similar to the one previously discussed that is apparent in the data reported by Yamauchi and Fujisawa (128). Unfortunately, the attempts to evaluate the kinetic constants for tyrosine and pterin of the activated hydroxylase in chromaffin cells were not successful. An observation that promises to provide further insight into the relationship between phosphorylation of tyrosine hydroxylase and the physiological control of this enzyme was reported by Haycock et al. (136). They found that 32Plabeled tyrosine hydroxylase isolated from chromaffin cells, when subjected to

8. AROMATIC AMINO ACID HYDROXYLASES

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limited proteolysis with trypsin, yielded two different 32P-labeled peptides that were separated by two-dimensional electrophoresis-chromatography. One of the peptides (designated “ C ” ) contained about twice as much 32P as the other one (designated “E”). Since further digestion with trypsin did not alter this ratio (and presumably did not convert one peptide into the other), the authors concluded that tyrosine hydroxylase is phorphorylated in situ at two different sites. Exogenous 8-bromo-CAMP when added to the cells at pH 6.0 was found to increase the amount of 32P incorporation into the E peptide whereas it had no effect on the phosphorylation of the C peptide. In contrast to these results, exogenous acetylcholine increased to about the same extent the 32Pincorporation into both peptides. These results indicate that the effects of exogenous 8-bromo-CAMP and acetylcholine on tyrosine hydroxylase phosphorylation are mediated by different protein kinases. Two potential candidates for the kinase that might mediate the acetylcholine effect in chromaffin cells are the calcium-activated, phospholipiddependent kinase (kinase C) first described by Nishizuka and his colleagues (137) and the calcium- and calmodulin-dependent protein kinase which Yamauchi and Fujisawa (138) showed was able to phosphorylate and in the presence of an activator protein (139) to activate brain stem tyrosine hydroxylase. Although kinase C is present in adrenal tissue (140) and has been reported to be capable of activating and phosphorylating tyrosine hydroxylase (141), the calcium- and calmodulin-dependent kinase has been reported to be absent from adrenal tissue (139).These results indicate, therefore, that of the two kinases that could potentially mediate the acetylcholine activation of tyrosine hydroxylase in chromaffin cells, protein kinase C is the more likely one. Whichever is the responsible kinase, it is not yet known whether the phosphorylation of peptide C or peptide E correlates better with activation of tyrosine hydroxylase. The finding that tyrosine hydroxylase in chromaffin cells can be phosphorylated on two distinct sites may have important implications for our ultimate understanding of the relationship between phosphorylation of tyrosine hydroxylase and its activation. These results indicate that fully phosphorylated tyrosine hydroxylase may contain at least 2 rnol Pi per mol M,60,000 subunit of tyrosine hydroxylase. In fact, Haycock et al. (136) based on their finding that C peptide incorporates twice as much phosphate as E peptide (C/Eratio equals 2) have predicted that it should be possible to incorporate 3 rnol Pi per mol of tyrosine hydroxylase subunit. On this last point, however, it should be noted that the C/E ratio of 2 may merely reflect phosphorylation of C peptide at twice the rate of E peptide rather than reflect two phosphorylation sites on C peptide and one site on E peptide. In either case, it is clear that the possibility raised by these studies that fully phosphorylated tyrosine hydroxylase may contain 2 or even 3 mol Pi per mol tyrosine hydroxylase subunit, could significantly alter and perhaps confound the interpretation of findings of less than stoichiometric amounts of Pi incorporat-

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ed into tyrosine hydroxylase. The previously discussed finding of Yamauchi and Fujisawa (128), that activation of adrenal tyrosine hydroxylase is accompanied by incorporation of about 0.25 mol Pi per mol of M, = 60,000 tyrosine hydroxylase subunit could mean that incorporation of only 10% of the maximum amount of Pi is associated with full activation of the enzyme. Studies on the activation of brain tyrosine hydroxylase by phosphorylation have lagged behind those on tyrosine hydroxylase from other tissues such as adrenal medulla or pheochromocytoma. Incorporation of unknown amounts of 32Piinto tyrosine hydroxylase purified from rat caudate nucleus was reported to be associated with activation of the enzyme (133). It is difficult to relate these findings to others that deal with phosphorylation of the enzyme, however, because the activated enzyme in this study showed no change in K , for either 6MPH, of tyrosine, but, instead, showed a marked increase in V,,,. The first study that reported the amount of 32Pithat could be incorporated into purified bovine striatal tyrosine hydroxylase was published in 1981 (142). In this important work it was shown that the catalytic subunit of CAMP-dependent protein kinase catalyzed the incorporation of 0.7 to 0.9 mol of 32Pi per mol of tyrosine hydroxylase subunit (M,60,000). Although these results leave no doubt that brain tyrosine hydroxylase is a substrate for CAMP-dependent kinase, no attempt was made in this study to correlate the extent of phosphorylation of the enzyme with the extent of activation. It is not possible to guess at what the correlation might be because in the experiments in which 32Piincorporation was determined, incubations for up to 240 min were carried out (142), whereas in other studies where the extent of activation of tyrosine hydroxylase was investigated, incubations were carried out for only 10 min (98). It is essential to know how much Pi was incorporated into the enzyme at the time when full activation was achieved, rather than the amount incorporated after an additional 4-h incubation with the kinase. As one may judge from the foregoing discussion, there is general agreement that exposure of tyrosine hydroxylase of various stages of purity and from different tissues to protein-phosphorylating conditions leads to activation of the enzyme. It may be clear, however, that a considerable amount of disagreement remains about the kinetic expression of this activation. Whereas most results agree with those first reported by Lloyd and Kaufman (124) which showed that activation of tyrosine hydroxylase by CAMP-dependent protein kinase is expressed as a decrease in the K,,, for BH, (or 6MPH, or DMPH,), with both the K,,, for tyrosine and the V,, essentially unchanged, others have reported that the activation is expressed exclusively as a change in V,, at pH 5.5 (133). Still others claim that the enzyme can be activated only to the most modest extent (12 to 25% decrease in K,,, for 6MPH,, with little change in V,,,) in gel-filtered rat striatal extracts (143). A clue that may explain some of the divergent results in this area was provided

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- 800

> 0

6

I

1

6.5

7 PH

7.5

0

6

I

I

6.5

7

>

PH

FIG. 10. V,,, and K,, for 6MPH4 of activated and control tyrosine hydroxylase at different pH values. Partially purified bovine-striatal tyrosine hydroxylase was incubated at pH 7.0 with beef brain protein kinase without ATP (control) or with ATP (activated). Aliquots from each incubation were then further incubated at the indicated pH with the components of the tyrosine hydroxylase assay plus EDTA to stop further protein kinase activity. Six different concentrations of 6MPH4 were used. Data from Ref. (144).

by a study of how the K , for the pterin cofactor and V,, vary with pH for both the control and the activated hydroxylase. Pollack er al. (144) found that, for partially purified bovine striatal tyrosine hydroxylase, the way in which activation by phosphorylation is expressed is critically dependent on the pH at which the hydroxylase activity is measured. As can be seen in Fig. 10, at pH 6.0, activation is expressed as a decrease in the K , for the pterin cofactor with little change in V,,,, whereas at pH 7.0-7.2, activation is expressed at a large increase in V,,, and a much smaller decrease in K,. Similar results were found with gel-filtered rat striatal extracts (144). As can be seen by comparison of the data in Fig. 10 and those in Table V, one difference between the partially purified bovine enzyme and this crude rat enzyme is that with the latter preparation, the velocity (V,,) of the activated enzyme is still higher at pH 6.2 than it is at pH 7.2. It should be noted, however, that even with the rat enzyme the shape of the pH-V,,, curve has been dramatically altered by declines by a factor of 50 on activation so that for the control enzyme, V,, going from pH 6.2 to pH 7.2, whereas for the activated enzyme, V,, declines by a factor of only 3.3 for the same shift in pH. One of the questions that these results have helped to clarify is whether activation of the enzyme by exposure to protein-phosphorylating conditions leads to a shift in the pH optimum of the enzyme. Lloyd and Kaufman (124) first

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TABLE V EFFECTOF PH ON KINETICPARAMETERS OF RAT-STRIATAL TYROSINE HYDROXYLASE~’ State of the enzyme Control

Activated

PH

K,, for 6MPH4 (W)

(nmollmin)

6.2 6.7 7.2 6.2 7.2

2200 1100 220 840 93

0.100 0.017 0.002 0.073 0.022

Vmax

aData are from Ref. (144).

reported such an alkaline shift for the bovine-striatal enzyme, the pH optimum increasing from 6.0 for the control to pH 7.4 for the activated enzyme. Although these results were qualitatively confirmed by Lazar et al. (98) (optimum pH shifted from 6.0-6.2 to pH 6.8) for bovine striatal tyrosine hydroxylase, an alkaline shift in pH optimum was not reported by Hegstrand et al. (145) or Goldstein et al. (146) for the rat striatal enzyme. The demonstration (144) that tyrosine hydroxylase in gel-filtered rat striatal extracts can be activated by a CAMP-dependent protein kinase and that the activation at pH 6.2 is expressed as a decrease in the K,,, for 6MPH4 is not in accord with the results reported by Ames et al. (143) who were unable to demonstrate activation of tyrosine hydroxylase (neither a significant decrease in K,,, for 6MPH, nor an increase in V,,,) in gel-filtered rat striatal extracts. Since their assays were carried out at pH 6, a large decrease in the K,,, for the pterin cofactor would have been expected from the results of Pollock et al. (144) and Lazar et al. (98). In agreement with the results of Pollock et al. ( 1 4 4 , but in contrast to those of Ames et al. (143), Vrana et al. (131) were also able to demonstrate activation of tyrosine hydroxylase by exposure of gel-filtered rat striatal extracts to phosphorylation conditions. They were, however, unable to replicate the finding of Ames et al. (143) that gel-filtration of striatal extracts by itself led to a decrease in the K,,, for 6MPH4. The explanation for the failure of Arnes et al. (143) to observe significant activation of tyrosine hydroxylase in gelfiltered extracts of rat striatum is obscure. But whatever the explanation might be, the conclusion that these authors drew from their results, namely, that activation of tyrosine hydroxylase by phosphorylation is “primarily dependent upon the presence of catecholamines” cannot be correct since this conclusion ignores the numerous reports, some of which have just been discussed, that tyrosine hydroxylase at various stages of purity can be activated after exposure to a protein phosphorylating system. In none of these studies with partially purified

8.

AROMATIC AMINO ACID HYDROXYLASES

26 1

tyrosine hydroxylase preparations can the activation have been dependent on the presence of catecholamines. The data shown in Fig. 10 and Table V clarify several other confusing issues about regulation of tyrosine hydroxylase activity. First, they indicate that the K , values for the pterin cofactor that have been determined at the pH optimum of the nonactivated enzyme (around pH 6) probably have little, if any, relevance to the physiological regulation of tyrosine hydroxylase. These values, as already discussed, appeared to be very high when compared to tissue levels of BH,. The finding that the K,,, for the pterin cofactor is at least an order of magnitude less at pH 7.2-7.4 than it is at pH 6 (Fig. 10, Table V), indicates that the problem of cofactor availability has probably been grossly overstated. Coherent with this conclusion that tyrosine hydroxylase does not normally operate under such a bizarre handicap, it has been found that in rat striatal synaptosomes, at physiological pH, the enzyme appears to operate with endogenous pterin cofactor concentrations that are much closer to saturation than would have been predicted from results of cell-free assays carried out at pH 6.0-6.2 (144). Thus, it was found that the addition of saturating amounts of BH, to synaptosomes increased the in situ activity of tyrosine hydroxylase by only 50%, a result that would be difficult to reconcile with the very high K , values for BH, that are found at pH 6.0-6.2, but which is readily understood by the lower K,,, value. The results of Pollock et af. (144) (Fig. 10) also predict that in the physiological pH range, activation of tyrosine hydroxylase by CAMP-dependent protein kinase would be expressed primarily as an increase in V,,,. Based on these in vitro results, it can be anticipated that in an intact tissue preparation maintained at physiological pH, the extent of activation of tyrosine hydroxylase by CAMPdependent kinase would be relatively independent of the BH, concentration. To test this prediction, the degree of activation of tyrosine hydroxylase in rat striatal synaptosomes by exogenous 8-bromo-CAMPwas studied as a function of varying concentrations of BH, added to the medium. In accord with the above expectations, it was found that the percentage of stimulation of in situ tyrosine hydroxylase activity by the cAMP derivative was relatively independent of added BH, concentrations. Thus, 8-bromo-CAMP stimulated to about the same extent in the absence (72%stimulation) and in the presence (79%stimulation) of 1 mM BH,, a result that is consistent with the conclusion that the cAMP derivative stimulated tyrosine hydroxylase mainly by an increase in V,,,. Also supporting this conclusion was the finding that the added cyclic nucleotide led to only a slight decrease in the apparent K,, for the exogenous BH, (0.34 mM versus 0.27 mM) (144). It should be emphasized that the data shown in Fig. 10 and Table V cannot explain the unique finding of Joh et af. (133) that activation of tyrosine hydroxylase from rat caudate nuclei by CAMP-dependent protein kinase is due to an increase in V,,, with no change in K,,, for the pterin cofactor even though the enzyme was presumably assayed at pH 5.5. Not only is this finding at variance

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with most other descriptions of how the kinetic properties of activated tyrosine hydroxylase are changed, but the explanation put forth by the authors does not adequately reconcile their results with those of others. They suggested, for example, that all of the earlier reports of a change in K,, for the pterin cofactor on activation of tyrosine hydroxylase were due to the presence of inhibitors (presumably catechols). This suggestion, however, ignored the finding that activation of extensively purified bovine striatal tyrosine hydroxylase by CAMP-dependent protein kinase was due to a marked decrease in the K,,, for the pterin cofactor (124). There is no more reason to believe that the hydroxylase preparation used by the latter workers was contaminated by inhibitors than there is to believe that the enzyme preparation used by Joh et al. (133) was so contaminated. In any case, many subsequent studies with highly purified tyrosine hydroxylase have confirmed the original finding of Lloyd and Kaufman (124) and have therefore shown that the explanation offered by Joh et al. (133) is incorrect. Subsequent results have dispelled any doubt that tyrosine hydroxylase from both adrenal medulla and from certain brain areas such as striatum can be phosphorylated by CAMP-dependent protein kinase. Despite these advances, however, it is still not possible to draw a detailed picture of the relationship of hydroxylase activity to the state of phosphorylation of the enzyme. What is lacking from the picture is any knowledge of how much (if any) protein-bound phosphate there might be in the tyrosine hydroxylase molecule before it is exposed to the protein kinase. Thus, even in the study of Vulliet et al. (130), which probably comes closest to describing the relationship between the amount of Pi incorporated and the extent of activation, there remain important gaps. As already discussed, for example, it was only during the first 15 min of incubation that tyrosine hydroxylase activity and extent of 32Piincorporation increased in a parallel fashion; beyond that period, 32Piincorporation continued to increase but tyrosine hydroxylase activity actually declined toward basal levels, introducing an element of arbitrariness into any attempt to correlate activation and 32Pi incorporation. But the biggest unknown in this as in all other investigations of this relationship is the total amount of protein-bound Pi before and after activation. Clearly, the interpretation of the finding of Vulliet et al. (130)that 0.7 mol of phosphate was incorporated/mol of 60,000 M,-subunit might be different if that 0.7 mol increased the total amount of protein-bound Pi up to 1 .O mol Pi/mol of 60,000 M,-subunit compared to the interpretation if that 0.7 mol brought the total amount up to 3.0 mol Pi/mol of subunit. In this regard, it should be noted that with rat-liver phenylalanine hydroxylase there does not appear to be a strict parallelism between the extent of activation and extent of phosphorylation. Rather, as previously discussed, most of the change in activity of the enzyme is associated with the transition from the quarter-phosphorylated tetramer ( 1 mol PJtetramer) to the half-phosphorylated tetramer ( 2 mol PJtetramer) (44). Although activation of tyrosine hydroxylase by CAMP-dependent protein

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kinase has been clearly established, there is also evidence that one or more CAMP-independent, Ca2 -dependent kinases can also activate the enzyme. Indeed, the first report that nerve stimulation leads to a persistent activation of tyrosine hydroxylase also demonstrated that activation of the enzyme in this system was strictly dependent on the presence of Ca2+ in the medium (128). Since Ca2+ was not known to affect the activity of CAMP-dependent protein kinase, this Ca2 requirement provided an early hint that another protein kinase might be involved in activating the hydroxylase in situ, although the possibility remained that Ca2 exerted some indirect effect on CAMP-dependent protein kinase . The first conclusive evidence that a Ca2 -dependent protein kinase could activate tyrosine hydroxylase was reported by Yamauchi and Fujisawa (138) who found that tyrosine hydroxylase in rat-brainstem cytosol can, in the presence of ATP and Mg2+, be activated not only by cAMP but also by Ca2+, the Ca2+ activation showing an absolute requirement for calmodulin. The additional finding that the cAMP activation and the Ca2 -calmodulin activation were additive suggested different mechanisms for these two modes of activation. When the Ca2 -calmodulin activation was studied with purified bovineadrenal tyrosine hydroxylase, it was found that the enzyme could be phosphorylated by a Ca2 -calmodulin-dependent kinase, but that the phosphorylated enzyme had not been activated (239). Addition of another component, namely, “activator protein,” was required to achieve activation of the phosphorylated enzyme. The maximum activation was about twofold and, in contrast to activation by CAMP-dependent protein kinase, was expressed exclusively as an increase in V,,, (147). These findings indicated that activation and phosphorylation by Ca2 -calmodulin-dependent protein kinase are two distinct reactions, the first one involving phosphorylation of the hydroxylase by the kinase and the second involving activation of the phosphorylated hydroxylase by the activator protein by a mechanism that has yet to be explored. The activator protein was not required for the Ca2 -calmodulin-dependent phosphorylation of tyrosine hydroxylase, nor did it have any effect on the activity of the unphosphorylated enzyme or, it should be emphasized, on the activity of the enzyme phosphorylated and activated by the CAMP-dependent kinase. The purified activator protein has a molecular weight of 70,000 (148). A single band on SDS-polyacrylamide gel, with an apparent molecular weight of 35,000 suggested that the activator protein is a dimer of identical subunits. The activator protein is present in all of the subcellular fractions, although the highest relative specific activity was found in the cytosol. In addition to brain, it was found in a number of peripheral tissues, such as adrenal glands, liver, heart, and skeletal muscle; however, its concentration in these tissues is lower than that in brain. The presence of activator protein in tissues that do not contain either tyrosine hydroxylase or tryptophan hydroxylase, the Ca2 -dependent phos+

+

+

+

+

+

+

+

+

+

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SEYMOUR KAUFMAN

phorylated form of which can also be activated by this protein, suggests other roles for this activator protein. The calmodulin-Ca2 -dependent protein kinase that catalyzes the phosphorylation of tyrosine hydroxylase and, as discussed in the next section, tryphophan hydroxylase, has been designated protein kinase I1 (147). It was reported to occur only in brain tissues, has a molecular weight of 540,000, and is composed of subunits, M, = 55,000 (147). It is of interest that Fujisawa and his co-workers have also demonstrated that rat brain cytosol contains two other calmodulin-Ca2 -dependent protein kinases with substrate specificities that are distinct from that of protein kinase 11. They have designated these other enzymes as kinase I, molecular weight about 1,000,000, which appears to be similar to brain phosphorylase kinase, and kinase 111, molecular weight about 100,000, which appears to be similar to myosin light chain kinase (147). As mentioned previously, it has been demonstrated that tyrosine hydroxylase in the cytosol from adrenal medulla can be activated by a CAMP-dependent phosphorylation and moreover, that 32P from [32P]ATPis incorporated into the enzyme (128). However, in contrast to the brain stem preparations, addition of Ca2+ to the cytosol from adrenal medulla did not lead to either activation or to the incorporation of 32P (129). It has been shown (148) that adrenal medulla has both calmodulin and activator protein but lacks the Ca2 -calmodulin-dependent protein kinase 11. Thus, although purified tyrosine hydroxylase from adrenal medulla can be activated by incubation with Ca2 , calmodulin, Ca2 -calmodulin-dependent kinase, activator protein, ATP, and Mg2+, the failure to detect kinase I1 in adrenal tissue would seem to eliminate the possibility that this particular Ca2 -calmodulin-dependent process is of any physiological significance for the activation of the hydroxylase in adrenal medulla. This last conclusion, however, must be regarded as a tentative one, since evidence has been presented that the original report that calmodulin-dependent protein kinase I1 is restricted to brain tissue (149) may have to be modified. In this regard, it has been found that a very similar enzyme, referred to as the “calmodulin-dependent multiprotein kinase,” is present in skeletal muscle (150) and liver (151, 152). Of particular interest, this calmodulin-dependent kinase appears to be present in pheochromocytoma cells, raising the possibility that it is also present in adrenal medulla (153). The kinase from skeletal muscle has been shown to be capable of catalyzing the phosphorylation of tyrosine hydroxylase purified from pheochromocytoma at a different site from that phosphorylated in the presence of CAMP-dependent protein kinase. Phosphorylation of the hydroxylase by the calmodulin-dependent multiprotein kinase did not activate the enzyme (153), a finding that is not surprising in view of the report that the hydroxylase that has been phosphorylated by calmodulin-dependent protein kinase (protein kinase 11) requires activator protein in order to express its activated state (136). Unfortunately, Vulliet et af. (153) did not test the effect of activator +

+

+

+

+

+

265

8. AROMATIC AMINO ACID HYDROXYLASES

protein on the hydroxylase that had been phosphorylated by calmodulin-dependent multiprotein kinase. Another Ca2+-dependent protein kinase that can catalyze the phosphorylation of tyrosine hydroxylase is protein kinase C, the Ca2 -activated, phospholipiddependent enzyme that was isolated and characterized by Nishizuka and his coworkers (137). The first indication that tyrosine hydroxylase might be a substrate for this kinase came from a study by Raese et al. (154). These workers reported the isolation from rat brain of a CAMP-independent protein kinase that can phosphorylate brain striatal tyrosine hydroxylase. Although the kinase was isolated by a minor modification of the procedures used by Nishizuka et al. (153a,b)for the purification of kinase C, the kinase used by Raese et al. (154) was not further characterized with respect to its catalytic properties. The incorporation into the hydroxylase of an undetermined amount of 32Pi from [32P]ATPin the presence of this kinase led to activation of the enzyme that was expressed exclusively as a decrease in the K , for BH, (from I .O mM to 0.22 mM), the same change as that seen when the hydroxylase is activated by the catalytic subunit of CAMP-dependent protein kinase at pH 6 (144). Subsequently, it was shown that pure protein kinase C from bovine cerebral cortex can, in fact, phosphorylate and activate tyrosine hydroxylase that has been purified from the cytosol of PC 12 pheochromocytoma cells (155). Activation, which led to the same changes in the kinetic properties of the enzyme as that seen after activation by CAMP-dependent kinase, namely, a decrease in K , for 6MPH,, no change in K , for tyrosine or in V,,,, all measured at pH 7.0, was accompanied by incorporation of about 0.5 mol 32Pi/molof M,62,000-60,000 doublet. Consonant with the observation that activation by protein kinase C and by CAMP-dependent protein kinase produced similar changes in the catalytic properties of the hydroxylase, the same peptide appeared to be phosphorylated by the two kinases. This last finding, incidentally, indicates that protein kinase C cannot be the enzyme responsible for the acetylcholine-mediated activation of tyrosine hydroxylase in chromaffin cells. It may be recalled that the acetycholine-mediated phosphorylation of the hydroxylase led to a different pattern of phosphorylation of the enzyme than did that produced by the addition of 8bromo-CAMP (133). In addition to the ability of CAMP-dependent protein kinase, Ca2 -calmodulin dependent protein kinase (protein kinase 11), and Ca2 -phospholipid-dependent protein kinase (protein kinase C) to catalyze the phosphorylation of tyrosine hydroxylase, there may be still another one that can carry out this reaction. Andrews et al. (156) reported that an unfractionated extract of rat striatal tissue contains an enzyme that, in the presence of ATP and Mg2+, can activate tyrosine hydroxylase in these tissue extracts, presumably by a process that involves phosphorylation of the enzyme. That the activation is not mediated by +

+

+

266

SEYMOUR KAUFMAN

CAMP-dependent protein kinase was supported by the finding that this activity was insensitive to protein kinase inhibitor. Furthermore, activation of the hydroxylase, which was expressed as a twofold increase in V,,, with no change in K , for either 6MPH, or tyrosine, was not affected by the addition of either Ca2+ or EGTA, suggesting that the responsible enzyme may not be dependent on Ca2 . It is noteworthy, however, that the change in the catalytic behavior of the enzyme is similar to that produced by the action of brain Ca2 -calmodulindependent protein kinase. Protein kinases that have been tested with highly purified tyrosine hydroxylase (from rat pheochromocytoma) and found to be essentially inactive are phosphorylase kinase, glycogen-synthase kinases 3 and 4, and casein kinases I and 11 +

+

(153).

Before considering the protein kinase(s) that might be involved in the acute regulation of tyrosine hydroxylase in response to nerve stimulation, it might be useful to summarize the manner in which activation of the hydroxylase by the action of different kinases is expressed. The results summarized in Table VI indicate that the effects of the different kinases on the properties of the hydroxylase are distinct, with activation by protein kinase I1 leading to an increase in V,,, with no change in K , for the pterin cofactor, activation by protein kinase C leading to the opposite change, and the effects of CAMP-dependent protein kinase depending sharply on the pH of the hydroxylase assay. Although it is clear from the data in Table VI that different kinases can produce different changes in the properties of tyrosine hydroxylase, the effects of nerve stimulation (or K depolarization) on the properties of the hydroxylase in neuronal tissues are less clear; in part, because these properties have been studied mainly in crude tissue extracts. Because of this limitation, attempts have also been made to answer the related, but more restricted, question of whether the change in the hydroxylase seen after nerve stimulation are consistent with those produced by the action of CAMP-dependent protein kinase. As for changes in the hydroxylase seen after nerve stimulation, as discussed earlier, stimulation of the guinea pig sympathetic nerve-vas deferens preparation has been reported to activate tyrosine hydroxylase, with the activation being expressed as a decrease in the K , for both the pterin cofactor (DMPH,) and tyrosine, with a concomitant 50% increase in V,,, (118), a pattern which differs from that mediated by any of the protein kinases listed in Table VI. Except for the changes in K, for tyrosine, however, which has not been reported for any of these kinases, it should be noted that these changes are similar to those reported after the hydroxylase has been activated by CAMP-dependent protein kinase at neutral pH. Some, but not all, of the results of Morgenroth et al. (118) were replicated by Weiner et al. (157). Using the same tissue (i.e., the isolated guinea pig vas deferens-hypogastric nerve preparation), they showed that stimulation of the +

8.

267

AROMATIC AMINO ACID HYDROXYLASES TABLE VI CHANCESIN THE CATALYTIC PROPERTIES OF TYROSINE HYDROXYLASE AFTER ACTIVATION^,^ Protein kinase

K , for pterin cofactor

CAMP-dependent (pH 6.0) CAMP-dependent (pH 7.0-7.4) Ca* -calmodulin-dependent (kinase 11) CaZ+ -calmodulin-dependent multiprotein Ca2+ -phospholipid-dependent (kinase C)(pH 7.0) Non-CAMP-dependent (unstimulated by Ca2 )

J.1 .1

+

+

Vmax

0

t t f

0 ND

ND

J.1

0

0

t

aThe data for CAMP-dependent protein kinase from Pollock et al. (144). The non-CAMPdependent kinase that is not stimulated by added Ca2+ is that described by Andrews et al. (156). bND, not determined; 0, little or no change.

nerve for 40 min increased the in situ activity of tyrosine hydroxylase 2- to 3fold. Since this stimulation was still evident in the presence of what seemed to be an excess of exogenous DMPH,, the authors concluded that the “major effect of nerve stimulation on tyrosine hydroxylase in intact tissue appears to involve an increase in the maximum velocity of the reaction. . . .” When the hydroxylase was assayed in extracts prepared from control and stimulated tissue, however, it was found that nerve stimulation had resulted in about a twofold increase in V,, and a 50% decrease in K,,,for 6MPH, (157). As can be seen in Table VI, these changes in the properties of the enzyme after nerve stimulation are similar to those that would be expected for the hydroxylase activated by CAMP-dependent protein kinase, if the hydroxylase assays had been carried out between pH 6 and pH 7 (144). Since the assays were reported to have been carried out at pH 6.2 ( 1 5 3 , the changes conform closely to those produced by the action on CAMP-dependent protein kinase. Cyclic-AMP was implicated in the mechanism of activation of tyrosine hydroxylase by nerve stimulation by Weiner et al. (157) who showed that the addition of 8-methylthio-CAMPto unstimulated hypogastric nerve-vas deferens preparations enhanced the in situ hydroxylase activity, whereas no significant enhancement occurred in the electrically stimulated organ. Furthermore, when sufficiently large amounts of the CAMP derivative were added to the tissue, the hydroxylase was activated to about the same extent as that seen after nerve stimulation. These results indicated that the cyclic nucleotide activated the hydroxylase in a manner that was similar to that of nerve stimulation and suggested that activation of tyrosine hydroxylase by nerve stimulation was mediated by a CAMP-dependent protein kinase. A similar suggestion was put forth by Roth et

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al. (158). Consistent with this view, the levels of cAMP in the stimulated vas deferens were shown to be increased, but the increase was only about 60%. Further support for this conclusion was provided by the observation that the hydroxylase that was isolated from tissue that had been exposed to the cyclic nucleotide showed changes in kinetic properties that were similar to those seen after nerve stimulation (i.e., a decrease in K,,, for 6MPH,, an increase in V,,,, with no change in the K, for tyrosine. Although these results provide support for the conclusion that CAMP-dependent protein kinase mediates at least part of the activation of tyrosine hydroxylase that is seen after nerve stimulation, they do not explain all of the findings. If CAMP-dependent protein kinase were the only kinase involved, for example, it would be difficult to explain the observation that the nerve-stimulation-mediated activation of the hydroxylase measured in situ appears to be expressed exclusively as an increase in V,, (157). There are also other indications that activation of the hydroxylase by a CAMPdependent protein kinase cannot account completely for the activation elicited by depolarization of neuronal tissues. One of the first bits of evidence suggesting that other mechanisms might be involved, in addition to CAMP-dependent activation, came from studies of the effect of depolarization on the activity of synaptosomal tyrosine hydroxylase. It was found that depolarization induced by the drug, veratridine, activated the enzyme but further activation could be achieved by dibutyryl-CAMP (146), suggesting that the depolarization-induced activation was additive with that mediated by the cAMP derivative. Another study of the possible involvement of a CAMP-dependent protein kinase in the depolarization-induced activation of tyrosine hydroxylase in rat striatal slices by Simon and Roth also led to a different conclusion from the one reached by Weiner et al., namely, that activation of the enzyme produced by depolarization and that produced by cAMP probably occur by different processes (159). The main difference between the results obtained by Weiner et al. (157, 160) with the electrically stimulated hypogastric nerve-vas deferens preparation and those obtained by Simon and Roth (159) with K+-depolarized striatal slices is that with the latter system, activation by depolarization was found to be additive with activation produced by exogenous dibutyryl-CAMP. Thus, despite the finding that both modes of activation produced similar changes in the kinetic properties of the hydroxylase, the observation that the cyclic nucleotide stimulated to about the same extent in the presence (3-fold) or absence (3.6-fold) of depolarizing concentrations of K f (which by itself stimulated 1.9-fold) indicates that activation by depolarization cannot be mediated entirely via a CAMP-dependent protein kinase. It would have been of considerable interest to determine the kinetic characteristics of the hydroxylase that had been activated by the simul-

269

8 . AROMATIC AMINO ACID HYDROXYLASES

taneous effects of the cyclic nucleotide and depolarization but neither K,,, nor V,,, values were reported. Another indication that the two modes of activation have distinct effects on tyrosine hydroxylase comes from the finding that, in confirmation of the report of Lloyd and Kaufman (124), the pH optimum of the enzyme that had been activated by dibutyryl cAMP was shifted from 6.0 to 7.5, whereas the pH optimum of the enzyme that was activated by depolarizing concentrations of K + was only slightly shifted. Furthermore, activation by depolarization, but not by the cyclic nucleotide, was dependent on the presence of Ca2+ in the medium (159). As an added argument against the notion that depolarization-induced activation of tyrosine hydroxylase might be mediated by any CAMP-dependent process, Bustos and Roth (161) pointed out that although chemical or electrical depolarization of brain cortical slices are known to increase the levels of cAMP in the tissue, this increase probably occurs in the postsynaptic cells, whereas tyrosine hydroxylase is located mainly presynaptically . Similar results were reported by El Mestikawy et al. (162). Using striatal slices, they confirmed the finding that Ca2+ is required for the activation induced by high potassium but not for that mediated by dibutyryl-CAMP. They also showed that when the hydroxylase was first activated by the CAMP-dependent process, it could still be further activated by ATP, Mg2+, and Ca2+. The converse was also true. Kinetic studies of the hydroxylase in striatal extracts indicated that the activation caused by high-potassium depolarization was characterized by a 1.6-fold increase in V,,, with no change in the apparent K,, values for 6MPH4 or tyrosine, whereas activation mediated by dibutyryl-CAMP was characterized by a decrease in K,,, for 6MPH4 (but not for tyrosine) with little or no increase in V,,,. Activation of the enzyme in striatal extracts by Ca2 in the presence of ATP and Mg2 could also be demonstrated. This effect of Ca2 in extracts was blocked by trifluoperazine, a calmodulin antagonist, indicating that the Ca2 -dependent activation of the hydroxylase in extracts might be catalyzed by a calmodulin-dependent protein kinase. As can be seen from the data in Table VI, the changes in the hydroxylase elicited by high K + depolarization (162) are those expected from the action of Ca2 -calmodulin-dependent protein kinase on the hydroxylase. In summary, although some results, especially those obtained with the guinea pig sympathetic nerve-vas deferens preparation, are consistent with the conclusion that the acute activation of tyrosine hydroxylase caused by nerve stimulation is due to the action of CAMP-dependent protein kinase (157, 158),there are also convincing data, especially from studies with brain tissue, that a CAMP-independent protein kinase (159, 161), perhaps a Ca2+-calmodulin-dependent kinase may be involved (162). At present, there are no data that indicate that protein kinase C plays a role in the acute activation of the hydroxylase that is elicited by nerve stimulation or depolarization. +

+

+

+

+

270

SEYMOUR KAUFMAN

D. THE STABILITY OF THE PHOSPHORYLATED FORMOF TYROSINE HYDROXYLASE An early indication that phosphorylation of tyrosine hydroxylase might decrease the enzyme's stability was reported by Lazar er al. (163) who found that the activation of highly purified brain (striatal) tyrosine hydroxylase by CAMPdependent protein kinase was biphasic. At 30°C they found that activation reached a peak within several minutes of incubation with the kinase system, but this increase in activity was followed by a rapid decrease to the preincubation levels, but not below. From these results, they concluded that the phosphorylated hydroxylase was less stable than the nonphosphorylated form, a conclusion that was supported by the finding that at 50°C, the half-life of the activated enzyme was 5 ? 2 min compared to a half-life of 15 3 min for the control, nonphosphorylated enzyme. Since there is evidence that indicates that the in vitro thermal stability of proteins may correlate with their in vivo half-lifes (164, Lazar ef al. suggested that phosphorylation of tyrosine hydroxylase could increase its rate of degradation in vivo. In 1983, Vrana and Roskoski (165) also presented evidence that phosphorylated tyrosine hydroxylase is less stable than the nonphosphorylated form. They found that activation of the hydroxylase by CAMP-dependent protein kinase in rat striatal homogenates also followed a biphasic course with the enhanced activity decreasing rapidly to levels seen in the absence of ATP, Mg2 , and CAMP. That at least part of this decline was due to dephosphorylation of the hydroxylase (subsequent to the hydrolysis of cAMP by endogenous phosphodiesterase) was indicated by the finding that a second addition of cAMP led to another increase in hydroxylase activity. Significantly, the level of activity reached after this second addition of cAMP was only about 50% of the peak reached after the initial addition of CAMP, an indication that another inactivation process, not reversed by phosphorylation, was taking place. Evidence for another, irreversible inactivation was obtained when the hydroxylase was activated under more constant phosphorylating conditions (incubation with the catalytic subunit of CAMP-dependent protein kinase). Under these conditions, the activation of the hydroxylase was also followed by a rapid loss of activity to a level that was much lower than that of the control enzyme. This loss of activity, interestingly enough, was due to a decrease in V,,,, with the enzyme retaining the lower K , for the pterin cofactor that is characteristic of the hydroxylase that had been activated by CAMP-dependent protein kinase. This cycle of activation followed by inactivation, therefore, appeared to lead to the formation of an unusual species of tyrosine hydroxylase, one with a low K,,, for the pterin cofactor and a very low V,,,. The mechanism of this irreversible inactivation of previously activated (and presumably phosphorylated) enzyme has not been elucidated, although it appears to involve the participation of a small molecule that

*

+

8. AROMATIC AMINO ACID HYDROXYLASES

27 1

can be removed from the extracts by gel filtration on a Sephadex G-15 column

(165). The reversible inactivation of activated tyrosine hydroxylase in striatal extracts indicated, as just mentioned, that a phosphatase that can dephosphorylate the phosphorylated enzyme is, in all likelihood, present in these unfractionated striatal preparations. Surprisingly, other than a related finding reported in preliminary form by Lazar et al. (166), and a report of a similar dephosphorylating activity present in extracts of bovine-adrenal medulla (126), the dephosphorylation of phosphorylated tyrosine hydroxylase has not been studied in any detail.

IV. Tryptophan Hydroxylase A.

INTRODUCTION

Considerably less is known about the third pterin-dependent aromatic amino acid hydroxylase, tryptophan hydroxylase, than is known about either phenylalanine or tyrosine hydroxylases. The enzyme is believed to catalyze the rate-limiting step in the biosynthesis of the putative neurotransmitter, 5-hydroxytryptamine (serotonin) (167, 168). In view of the fact that the pathway involved in serotonin synthesis consists of only two consecutive steps, hydroxylation of tryptophan to 5-hydroxytryptophan, followed by decarboxylation of the hydroxylated amino acid to serotonin, and the fact that the decarboxylase has much higher in vitro activity than does the hydroxylase (169), the conclusion that the hydroxylation step is limiting is reasonable, if largely untested. The first reports of the in vitro hydroxylation of tryptophan catalyzed by mammalian tissue preparations were published in 1961. In that year, Cooper and Melcer (170) reported the presence in intestinal mucosa of a particulate, ascorbate-dependent enzyme that could catalyze the anaerobic hydroxylation of tryptophan. In the same year Freedland et al. reported the presence of a soluble tryptophan hydroxylase in a rat liver supernatant fraction that required oxygen and reduced pyridine nucleotide (171). The claim of Cooper and Melcer has never been confirmed, whereas the tryptophan hydroxylating activity in rat liver was traced to phenylalanine hydroxylase (172). In 1964, Grahame-Smith (167) reported evidence for the presence of a specific tryptophan hydroxylase in a normal mammalian tissue. The enzyme was detected in whole homogenates, but not in the high-speed supernatant fraction, of brain stem from dogs and rabbits. In dog brain, the activity was found in the hypothalamus-thalamus and the midbrain-medulla regions; no activity was detected in the cerebellum or cortex. The anatomical distribution of the enzyme closely paralleled the serotonin content of these areas.

272

SEYMOUR KAUFMAN

After Grahame-Smith’s demonstration that a specific tryptophan hydroxylase exists in the brain, the enzyme was partially purified from this tissue and from pineal glands, the latter tissue being an especially rich source of the enzyme.

B. CATALYTIC PROPERTIES With the soluble enzyme from both rabbit brain and bovine pineal glands, an absolute requirement for a tetrahydropterin (DMPH,) was demonstrated (I73). Subsequently, Lovenberg and his co-workers also reported (174)that the naturally occurring cofactor, BH,, could stimulate tryptophan hydroxylation with the crude soluble enzyme from rat brain stem. The K,,, value that they obtained for BH,, however [i.e., 5 pM), was so much smaller than values that were subsequently reported with authentic BH, (i.e., 31 pM for the enzyme from rabbit brain (15) and 57 pM for the enzyme from rat brain stem (175)] that there is some uncertainty about the identity of the pterin that was actually used in those early experiments. In contrast to unactivated rat liver phenylalanine hydroxylase, where V,,, with BH, is much lower than it is with synthetic tetrahydropterins such as DMPH, and 6MPH, (31, 33), tryptophan hydroxylase resembles much more closely tyrosine hydroxylase where the activity of the unactivated enzyme with BH, equals or exceeds the activity with these synthetic pterins. Thus, Friedman er al. (15) reported that the maximum velocity for rabbit-brain tryptophan hydroxylase in the presence of BH, is slightly higher than it is with DMPH,. Similarly, with the hydroxylase from rat brain it has been found that V,,, in the presence of BH, is about 10% higher than it is with 6MPH, (175). Although it was reasonable to assume that tetrahydropterins function with tryptophan hydroxylase in the same way that they were first shown to function with phenylalanine hydroxylase (16), this assumption could not be tested until the enzyme had been partially purified. Indeed, it had not been shown that the pterin functions catalytically during tryptophan hydroxylation. In 1972, Friedman et al. (IS),using a partially purified preparation of the hydroxylase from rabbit brain, proved this point for the first time when they showed that with 8.5 nmol of tetrahydrobiopterin, 40 nmol of 5-hydroxytryptophan were formed. The demonstration of the catalytic role for the pterin implies that the tetrahydropterin must be capable of being regenerated during the hydroxylation reaction. Unfortunately, the use of 2-mercaptoethanol in the assay employed by some workers (176) has made it difficult to demonstrate the need for a tetrahydropterin-generating system. Sulfhydryl compounds can stimulate pterin-dependent hydroxylases not only by reducing quinonoid dihydropterins (6, 177) but perhaps also by stabilizing one of the enzymes in the system. Thus, Lovenberg et a1. (173), with a soluble enzyme preparation that showed

8.

AROMATIC AMINO ACID HYDROXYLASES

273

a sharp requirement for DMPH,, were unable to demonstrate a reduced pyridine nucleotide requirement, probably because 2-mercaptoethanol was functioning to keep the pterin reduced. However, Gal el af. (178),with a particulate preparation of the enzyme that showed only a minimal stimulation by a tetrahydropterin, were able to show a twofold stimulation of the rate of tryptophan hydroxylation with NADPH. Ichiyama et af. (179) were the first to show a clear-cut stimulation of the reaction by the combination of DMPH, and NADH or NADPH. The first demonstration that tryptophan hydroxylation can be stimulated by highly purified dihydropteridine reductase, in addition to NADPH and tetrahydropterin, was reported by Friedman et af. (15). Since it had previously been established that the substrate for dihydropteridine reductase is a quinonoid dihydropterin ( l o ) , the stimulation of tryptophan hydroxylation by the reductase provides strong support for the conclusion that, just as is the case with phenylalanine hydroxylase, the quinonoid dihydropterin is the product of tetrahydropterin oxidation during the enzymic hydroxylation of tryptophan. It is surprising that for several years after tryptophan hydroxylase had been described, the stoichiometry of the reaction catalyzed by the enzyme had not been determined. It was tacitly assumed that the stoichiometry was the same as that previously established for phenylalanine (180) and tyrosine (14) hydroxylases. Friedman et al. (15) showed that with BH, the assumption was correct. They observed that 1 mol of 5-hydroxytryptophan is formed for each mol of tetrahydrobiopterin oxidized. This result, taken together with their demonstration that the hydroxylation reaction is stimulated by dihydropteridine reductase, led to the formulation of the reaction as shown in Eq. 3, where BH, stands for tetrahydrobiopterin and q-BH, stands for quinonoid dihydrobiopterin. L-Tryptophan

+ BH4 + O2 -+ 5-hydroxy-~-tryptophan+ q-BH2 + H20

(3)

Subsequently, it was shown that the stoichiometry of the hydroxylation reaction in the presence of the two widely used pterin cofactor analogues, DMPH, and 6MPH,, is the same as that shown in Eq. (3) (i.e., 1 mol of 5-hydroxytryptophan formed for each mol of tetrahydropterin oxidized) (181). A catalytic property of the hydroxylase that is relevant to a discussion of its possible regulation by phosphorylation is the K, value for its substrate, Ltryptophan. For the soluble hydroxylase from both rat brain stem (176) and guinea pig brain stem (179), a value of 300 (in the presence of DMPH,) has been reported. This value of 300 pkf generated a considerable amount of speculative interest because it is far above the plasma or the brain levels of tryptophan, both of which are about 30 pibl for rats (182, 183). (The brain concentration was calculated from the reported value of 5 p g / g of tissue by assuming uniform distribution of

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SEYMOUR KAUFMAN

the amino acid in the tissue.) These considerations led to the conclusion that tryptophan hydroxylase activity in brain is severely limited by availability of its amino acid substrate [Fernstrom and Wurtman (182), and references therein]. What was not fully appreciated by these early speculations was that, just as it was first demonstrated for phenylalanine hydroxylase (7) with respect to the K , for phenylalanine, the K , of tryptophan hydroxylase for L-tryptophan varies markedly with the pterin cofactor used in the assay. With the partially purified rabbit brain enzyme, the following K,,, values for tryptophan (the pterin cofactor used in the assay is shown in parenthesis) were found to be 50 pM (BH,); 78 pM (6MPH,); and 290 pA4 (DMPH,) (15). There is little doubt that a K , value of 50 pM, the value obtained in the presence of BH,, is more consistent with in vivo observations than is the previously accepted value of 300 pM. It has been shown (182) that the concentration of serotonin in the brain increases when the concentration of tryptophan in the brain is increased from 5 pg/g (about 30 pM) to 15 pg/g (about 90 @ but I), further increases of tryptophan (up to 45 pg/g) do not lead to further increases in serotonin content. The hydroxylating system in brain, therefore, appears to be saturated with tryptophan at approximately 90 pM. These in vivo observations agree very well with the apparent K , value of 50 pM (15) but would be difficult to explain if one accepted the higher value of 300 pM. Thus, it does not appear that this hydroxylase operates in vivo under any unique disadvantage of substrate limitation, but rather it appears to function, as probably most other enzymes do (184), with tissue concentrations of its substrate in the region of the K , value of the substrate.

C. PHYSICAL PROPERTIES The partially purified enzyme from rabbit hindbrain was reported to have a molecular weight of 220,000 to 240,000 (181). On polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate, two major protein bands, M, = 57,500 and M, = 60,900, were detected (181). Based on these results, a tetrameric structure was proposed for the native enzyme. The hydroxylase from rat brain stem has been highly purified (185). During chromagraphy on DEAE-Sepharase CLdB, two forms of the enzyme were separated. On further purification, each of these forms was obtained in essentially pure form. On polyacrylamide gel electrophoresis in the presence of sodium dodecylsulfate, both forms gave a single sharp protein band with the same migration pattern. On electrophoresis in the absence of sodium dodecyl sulfate, however, both forms gave single, broad protein bands with slight differences in mobility. The specific activity of the more abundant species (form 11) was about 20%that of the less abundant form (form I). The molecular weight of form I was

275

8. AROMATIC AMINO ACID HYDROXYLASES

reported to be between 288,000 and 300,000, and appeared to be composed of four apparently identical subunits, M, = 59,000. Unlike rat liver phenylalanine hydroxylase (12) and bovine adrenal tyrosine hydroxylase (96), where there is direct evidence that these enzymes are iron proteins, with tryptophan hydroxylase direct evidence is lacking. There are, however, indications that this hydroxylase may also be an iron protein. It has been reported, for example, that both the brain and pineal enzymes are inhibited by iron chelators (174, 178, 179). Data have also been published showing that the hydroxylation of tryptophan can be stimulated by Fe2+. In this regard, Lovenberg et al. reported that the activity of the hydroxylase from pineal glands is stimulated twofold by the addition of Fez+ (173, 186), and concluded that this result constitutes a direct demonstration of the role of iron in tryptophan hydroxylase (176). By contrast, the activity of brain enzyme has been reported to be either not stimulated by Fe2 (173) (rabbit brain enzyme) or stimulated only to the extent of 50% (179) (guinea pig brain enzyme). Nakata and Fujisawa (185) have shown that the essentially homogeneous hydroxylase from rat brain stem can be stimulated about 3.5-fold by Fez+. At least part of the reported stimulation by added Fez has been traced to the ability of Fe2 to protect the hydroxylating system against the deleterious effects of H202, the latter compound being generated through the nonenzymic oxidation of tetrahydropterins (15). In the presence of an optimum amount of catalase and with the use of dihydropteridine reductase and NADPH to regenerate the tetrahydropterin, it was shown that the activity of partially purified tryptophan hydroxylase from rabbit hindbrain is not stimulated by the addition of 1 mM Fe2+ (15). Despite the fact that direct evidence on this point is not yet available, the inhibition by metal chelators, together with the many properties shared by the three pterin-dependent aromatic amino acid hydroxylases, lead to the strong expectation, as stated previously (12), that tryptophan hydroxylase will also prove to be an iron enzyme. +

+

+

D.

REGULATION BY PHOSPHORYLATION

As discussed in the previous section, the demonstration that neuronal tyrosine hydroxylase could be activated in response to nerve stimulation and that the activation was dependent upon the presence of Ca2+ in the medium, led to a series of studies that culminated in the demonstration that this acute activation of the hydroxylase probably involves its phosphorylation. Similarly, it had been shown that neuronal tryptophan hydroxylase could also be activated in response to electrical stimulation (187-189). Since the influx of Ca2+ into nerve terminals is known to be increased in response to stimulation of the nerve, the activation of the hydroxylase under these conditions prompted a search for a Ca2 -dependent activation of the enzyme. +

276

SEYMOUR KAUFMAN

This search appeared to be successful. Several groups reported that crude brain tryptophan hydroxylase could be activated by Ca2+, the activation, in general, being expressed as approximately a twofold decrease in K,,, for both the pterin cofactor and tryptophan, with little or no increase in V,,, (190-192). The very high concentrations of Ca2+ that were needed to elicit this activation, which were in the millimolar range and were therefore three to four orders of magnitude higher than normal cytoplasmic concentrations, indicated that this Ca2+ effect might not be of any physiological significance. Indeed, evidence was presented showing that activation of the hydroxylase by these high concentrations of Ca2 was, in all likelihood, due to the irreversible proteolytic activation of the enzyme (192). A MgATP-dependent activation of tryptophan hydroxylase in extracts of mouse brain, that was characterized by a decrease in K,,, for 6MPH4 with no change in K,,, for tryptophan and no change in Vmax, was reported in 1978 (193). The findings that hydroxylase activity was decreased by treatment with acid phosphatase and that the ATP could not be replaced by a nonutilizable analogue of ATP indicated that activation of the enzyme might involve its phosphorylation. Significantly, evidence was presented against the participation of CAMPdependent protein kinase in the activation. These results were extended when it was shown that the MgATP-dependent activation of the hydroxylase in rat brain stem extracts also showed a requirement for Ca2 , but, in contrast to the millimolar concentrations that had been implicated in the proteolytic activation of the enzyme, this MgATP-dependent process required only 5 to 10 pJ4 Ca2+ (194, 195), enhancing the likelihood that this activation did not involve the action of a Ca2 -dependent protease. Hamon et al. (194) found that the K , values for both 6MPH4 and tryptophan were decreased with V,,, remaining essentially unchanged, whereas Kuhn et al. (195) reported that activation was expressed exclusively as a decrease in the K , for 6MPH4 (from 0.21 mM to 0.09 d). The conclusion that the MgATP- and CaATP-dependent activation of the hydroxylase was due to its phosphorylation was supported by the finding that in rat brain stem extracts, the twofold activation that was apparent after incubation in the presence of ATP, Mg2+, and Ca2+ could be reversed by a second incubation in the presence of the Ca2+ chelating agent, EGTA. The deactivation was inhibited by the omission of Mg2+ and the addition of NaF, findings that indicated that the deactivation was due to the action of a phosphatase. This conclusion was also supported by the finding that the deactivated enzyme could be activated a second time by the addition of excess ATP and Ca2+ (196). The nature of the Ca2+ -dependent system responsible for the activation of tryptophan hydroxylase in rat brain stem was clarified (197) when it was shown that the protein kinase system is a complex one consisting of the protein kinase itself, which was purified from rat cerebral cortex and designated protein kinase I1 (198), calcium-dependent regulator protein (calmodulin) ( 1 9 9 , and an ac+

+

+

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tivator protein that was purified from rat brain to apparent homogeneity (148). As discussed in the previous section, this particular calmodulin-dependent protein kinase and the activator protein are also able to activate tyrosine hydroxylase, although with that enzyme it had been shown that the activator protein was not necessary for the phosphorylation of the hydroxylase but only for the expression of the enhanced activity of the phosphorylated enzyme. Although these studies by Fujisawa and his colleagues have identified the components of this kinase system that is capable of activating brain tryptophan hydroxylase, it must be emphasized that phosphorylation of the hydroxylase has not been demonstrated. In fact, the size of the gaps in our knowledge of the mechanism by which this hydroxylase is activated can be gauged by the report that pure preparations of rat brain tryptophan hydroxylase are only slightly activated by kinase I1 in the presence of activator protein (198). Unfortunately, the ability of kinase I1 to catalyze the phosphorylation of the pure hydroxylase was apparently not determined. It should also be noted that kinase I1 is not specific for tryptophan and tyrosine hydroxylases. Tubulin was identified as one of the endogenous substrates for this kinase in the soluble fraction from brain tissue (199). The K , for tryptophan hydroxylase, however, was found to be much lower (0.0003 p N ) than the K , for tubulin (1.7 pM) (147). Despite the many reports that exogenous cAMP does not activate brain tryptophan hydroxylase in crude, soluble fractions from brain (193-195), it has been found that dibutyryl cAMP administered intracerebroventricularly (200) or added to slices of rat brain stem (201) can activate the hydroxylase. The activation observed in the latter study was characterized by about a 30% decrease in the K,n for 6MPH, and for tryptophan, with a 23 to 29% increase in V,,,,, (201). Further studies, however, have provided little support for the idea that cAMP is involved in activating brain tryptophan hydroxylase. Thus, in contrast to the ability of dibutyryl cAMP to activate the enzyme, 8-bromo-CAMPwas found to be inactive in this system (202). Furthermore, exposure of the brain stem slices to several other treatments that would be expected to raise intraneuronal levels of CAMP, such as cholera toxin, adenosine, or 2-chloroadenosine, also failed to activate the hydroxylase (202). Other than the previously mentioned evidence that an enzyme is present in brain stem extracts that is capable of reversibly deactivating tryptophan hydroxylase that had previously been activated by an MgATP and CaATP (196), there have been no direct studies of a brain phosphatase that can act on the phosphorylated form of tryptophan hydroxylase.

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175. 176. 177. 178. 179. 180. 181. 182. 183. 184. 185. 186. 187. 188. 189. 190. 191. 192. 193. 194. 195. 196. 197. 198. 199. 200. 201. 202.

Section II

Control of Biological Processes

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Phosphorylation of Bra in Proteins S. IVAR WALAAS PAUL GREENGARD Laboratory of Molecular and Cellular Neuroscience The Rockefeller University New York, New York 10021-6399

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Types of Protein Phosphorylation Systems in Brain . . . . . . . . . . . . . . . . . B. Direct Evidence for Involvement of Protein Phosphorylation in Neuronal Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Protein Kinases in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Cyclic Nucleotide-Dependent Protein Kinases B. Calcium-Dependent Protein Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Calcium- and Cyclic Nucleotide-Independent Protein Kinases . . . . . . . . . 111. Phosphoproteins in the Brain A. Regional Distribution of Brain Phosphoproteins . . . . . . . . . . . . . . . . . . . . B. Classes of Neuronal Phosphoproteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Phosphorylation of Specific Brain Proteins . . . . . . . . . . . . . . . . . . . . . . . . IV. Protein Phosphatases in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

The nervous systems of multicellular organisms represent enormously complex tissues that are highly specialized for intercellular signal transmission and information processing (1-3). A major focus of neurobiological research for 285 THE ENZYMES, Val XVllI Copynght 0 1Y87 by Academic Preas. Inc All rights of repruducuon in any form reserved

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many years has been the elucidation of the mechanisms by which these tissues achieve intercellular communication, and in particular the molecular mechanisms by which extracellular signals such as neurotransmitters produce their biological responses in specific target cells. Work in numerous laboratories has indicated that protein phosphorylation is importantly involved in these signaling mechanisms. In this chapter, we review several aspects of the vital role of protein phosphorylation in nervous system function, and discuss some of the most thoroughly investigated protein phosphorylation systems found in mammalian brain. No complete coverage of the literature is intended, and the reader is referred to several recent reviews for more detailed information (4-7). Most intercellular communication in the brain takes place through first messengers (e.g., neurotransmitters, hormones, and neuromodulators), which are released from one cell in order to interact with receptors located on specific target cells. Two general types of responses are generated in nerve cells by such receptor activation. One consists of rapid, direct regulation of various ion channels. This in turn induces changes in membrane potential and action potential frequency, and represents the primary, mediatory response of nerve cells (2, 3 ) . A different type of response consists of more long-term regulation of intracellular biochemical systems in nerve cells. Research in the area suggests that protein phosphorylation is the most important of such intracellular biochemical mechanisms. Indeed, a great variety of physiological stimuli appear to produce diverse responses in nerve cells, including regulation of intermediary metabolism, modulation of neuronal excitability, neurotransmitter biosynthesis and release, and regulation of neuronal growth, differentiation, and morphology by regulating the state of phosphorylation of specific neuronal proteins (5-8). A. TYPESOF PROTEINPH~SPHORYLATION SYSTEMS IN BRAIN The protein phosphorylation systems found in mammalian tissues, including the brain, consist minimally of three components: protein kinases, which catalyze phosphorylation of proteins on specific amino acid residues; specific phosphoprotein substrates for these kinases, which change their biophysical and functional properties upon phosphorylation; and phosphoprotein phosphatases, which dephosphorylate the phosphoproteins and thereby return the particular protein phosphorylation system to its initial state. Activation of protein phosphorylation systems can be achieved either by activation of protein kinases and/or inhibition of protein phosphatases. Evidence suggests that most intercellular signaling in the brain achieves such activation by increasing the activity of distinct protcin kinases, although some brain protein phosphatases also appear to be under physiological control by extracellular messengers (see Section IV). Two general mechanisms for activating protein kinases have so far been defined in mammalian brain: activation through the generation of intracellular ‘‘second

9. PHOSPHORYLATION OF BRAIN PROTEINS

287

messengers,” or activation through direct binding of the first messenger to a receptor-protein kinase complex. Activation through second messenger generation appears to be the most prominent of these mechanisms. Nerve cells in the brain contain a variety of hormone and neurotransmitter receptors that can generate second messengers. The best-known second-messenger-generating systems include the well-known receptor-activated adenylate cyclases and guanylate cyclases, which synthesize the cyclic nucleotides, cyclic AMP, and cyclic GMP, respectively, and the receptor-linked or voltage-sensitive Ca2 channels, which upon receptor activation or depolarization, respectively, allow Ca2+ influx into the neurons (9-12). A receptor-linked phospholipase C activity has also been characterized (13, 14), which, upon receptor activation, generates inositol 1,4,5-trisphosphate and diacylglycerol from phosphatidylinositol 4,5-bisphosphate. The compounds generated by these systems (i.e., the cyclic nucleotides, Ca2 , diacylglycerol, and inositol trisphosphate) are all believed to be intracellular messengers. Furthermore, they all appear to achieve some or all of their effects through the regulation of specific protein kinases. Cyclic AMP-dependent, cyclic GMP-dependent , and the different classes of calcium-dependent protein kinases are therefore importantly involved in brain function. The second major mechanism through which brain protein phosphorylation systems are regulated is exemplified by the protein kinase activities that are associated with various growth factors and hormone receptors. In this case, binding of the extracellular messenger to its receptor appears to directly activate the protein kinase, which often is a part of the receptor itself, without any intervening second messenger (15, 16). The importance of these activation mechanisms for protein kinases in brain function is not well understood, however. +

+

B. DIRECTEVIDENCE FOR INVOLVEMENT OF PROTEIN PHOSPHORYLATION IN NEURONAL FUNCTION Direct evidence for the functional importance of protein phosphorylation in neuronal function has come from various electrophysiological studies (Table I). Early investigations predicted that cyclic nucleotides might be neurophysiologically active, since correlations between experimentally changed levels of cyclic AMP or cyclic GMP and specific electrophysiological neuronal properties were observed (9, 17). These predictions have been confirmed and extended to include the other second-messenger systems. Direct injections of various components of protein phosphorylation systems into identified invertebrate nerve cells (18-30) have shown, for ex. .nple, that the catalytic subunit of cyclic AMP-dependent protein kinase can regulate transmitter release from a sensory neuron in the mollusc Aplysia, apparently through phosphorylation and

TABLE I SYSTEMSI N WHICHDIRECTEVIDENCE HAS BEENOBTAINED FOR

Cell (genus)

Kinase injectedb

A

ROLEOF PROTEINPHOSPHORYLATION IN NEURONAL FUNCTION"

Inhibitor injectedC

Bag cell neurons (Aplysiu)

Cyclic AMP

PKI

Sensory neurons (Aplysiu)

Cyclic AMP

PKI

Neuron R15 (Aplvsiu)

PKI

N m 00

Conclusion of Studies

Reference

Kinase mediates effect of synaptic activation and of exogenous cyclic AMP in producing the afterdischarge; this action appears to be achieved through decreases in the conductance of calcium-dependent potassium channels and of early (IA) voltagedependent potassium channels. Kinase mediates effect of synaptic activation and of exogenous serotonin and cyclic AMP in facilitating neurotransmitter release in response to nerve impulses; this action appears to be achieved through decreases in the conductance of novel serotoninregulated potassium channels. Kinase mediates effect of exogenous serotonin and cyclic AMP in inhibiting bursting activity and in enhancing interburst hyperpolarization; this action appears to be achieved through increases in the conductance of novel serotonin-regulated. anomalously rectifying potassium channels.

(21, 22)

(18-20)

(23-25)

Unidentified neurons (Helix)

Cyclic AMP

Unidentified neurons (Helix) Photoreceptor cells (Herrnissenda)

Cyclic AMP

Hippocampal (CAI) pyramidal neurons (Cuvia) Bag cell neurons (Apiysiu)

Cyclic AMP

Terminal digits of giant synapse (Loligo) Cortical pyramidal neuron (Felix)

Calcium-calmodulin I1

~~

Tolbutamided

Cyclic AMP

Calcium-phospholipid

Cyclic GMP

~~

~

PKI

Kinase increases the conductance of calcium-dependent rectifying potassium channels. Kinase increases the conductance of voltage-dependent calcium channels. Kinase decreases the conductance of early (IA) and late (I,) voltage-dependent calcium channels. Kinase mediates effect of dopamine and cyclic AMP in producing a longlasting increase in input resistance. Kinase increases the height of action potentials; this action appears to be achieved through increases in the conductance of calcium channels. Kinase facilitates neurotransmitter release. Kinase mediates effect of acetylcholine and cyclic GMP in producing increase in input resistance. ~~

("Modified from Nestler and Greengard (7)and Hemmings et a/. (34). bProtein kinase abbreviations: cyclic AMP, catalytic subunit of cyclic AMP-dependent protein kinase; cyclic GMP, cyclic GMP-dependent protein kinase holoenzyme; calcium-phospholipid, calcium-phospholipid-dependent protein kinase holoenzyme; calcium-calmodulin 11, calcium-calmodulin-dependent protein kinase I1 holoenzyme. CPKI, specific protein inhibitor of cyclic AMP-dependent protein kinase. dThe authors claim that tolbutamide is a specific inhibitor of cyclic AMP-dependent protein kinase in v i m (26).

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regulation of K+channels (18-20). Similarly, the same protein kinase can regulate action potential firing frequency in the bag cell neurons of Aplysia, an effect which appears to be mediated through regulation of other K channels (21, 22), while the diacylglycerol-activated, Ca2 -phospholipid-dependent protein kinase (protein kinase C) appears to regulate Ca2+ channels in the bag cell neurons of Aplysia (30).Other studies have shown effects of activators of protein kinase C on ion channels in hippocampal pyramidal cells (31). Evidence also suggests that intracellular injection of cyclic GMP-dependent protein kinase can regulate input resistance, in this case in certain mammalian cortical pyramidal cells, thereby mimicking the effect of acetylcholine on these cells (33).Studies with Ca2 -calmodulin-dependent protein kinases have shown that another important nerve cell function, that is, neurotransmitter release (in this case from the squid giant axon), can be directly modulated by intracellular injection of components of protein phosphorylation systems, in this case without any apparent effect on ion channel properties (32).These and other studies have provided direct evidence that protein phosphorylation is importantly involved in signal transmission and information processing in invertebrate nerve cells. However, due to the small amount of tissue available in invertebrate preparations, only limited biochemical information is available about the molecular properties of the specific protein kinases, protein substrates, and protein phosphatases that are involved in these effects. In contrast, mammalian brain has turned out to be an unusually rich source of protein kinases, phosphoprotein substrates, and phosphoprotein phosphatases (8, 35, 36). +

+

+

II. Protein Kinases in the Brain The properties of protein kinases in the mammalian brain are generally similar to the properties of those present in nonnervous tissues (37, 38). Certain differences exist for some of the enzymes, however, especially in their relative concentrations and their cellular and subcellular distributions. In this section, we briefly describe some properties of the well-studied brain protein kinases. A, CYCLICNUCLEOTIDE-DEPENDENT PROTEINKINASES Mammalian brain contains the two distinct cyclic nucleotide-dependent protein kinases, namely cyclic AMP-dependent and cyclic GMP-dependent protein kinases (38), which have been found to be the principal intracellular binding proteins for these second messengers. Some structural homologies between these enzymes exist, but the differences (e.g., in substrate specificities, activation mechanisms, immunological cross reactivities, and tissue distributions) suggest that they have distinct physiological roles (37) (See also Volume XVII, Chapter 3).

9. PHOSPHORYLATION OF BRAIN PROTEINS

29 1

1. Cyclic AMP-Dependent Protein Kinase Early work established that cyclic AMP-dependent protein kinase, which appears to be the main or only mediator of the effects of cyclic AMP in vertebrate tissues, was highly enriched in brain (39). The main properties of the brain enzyme are identical to those of the enzyme prepared from peripheral tissues. Thus, it exists as a tetramer composed of two regulatory (R) (M,= 49,00055,000) and two catalytic (C) subunits (M,= 40,000, as determined by SDS-gel electrophoresis). In the absence of cyclic AMP, the two R-subunits, joined by a disulfide bond, are bound to the two inactive C-subunits. Each R-subunit has two binding sites for cyclic AMP, and exhibits cooperative binding of cyclic AMP. Upon binding, the C-subunits dissociate from the holoenzyme, and are then catalytically active (37, 38, 40). The isozymes of cyclic AMP-dependent protein kinase exhibit differences in their R-subunits. Two main isozymic forms are known, type I having R,-subunits (M,= 49,000) and type I1 having R,,-subunits (M,= 52,000-55,000). Differences between R, and R,, within each tissue have been extensively investigated (37, 38, 40). There is, in addition, evidence for differences between R,, from different tissues. For example, antibodies against brain R,, do not completely cross-react with cardiac R,, (41, 4 2 ) , comparative tryptic peptide mapping indicates differences between brain, skeletal muscle, and cardiac muscle R,, (43, 4 4 , and brain and muscle R,, appear to interact differently with the C-subunit (45).Finally, isoelectric variants of both R, and R,, have been found in brain (42, 46). Such differences presumably reflect adaptations of the isozymes for specific neuronal functions. Both the R, and R,, forms of cyclic AMP-dependent protein kinase have been found to be widely distributed throughout the brain (35, 36,47).Type I1 appears to be present predominantly in neuronal cells (48, 49), while type I has been found, for example, in myelin (50), and may be enriched in glial cells (48). Mammalian brain has, in contrast to other tissues, a high activity of cyclic AMPdependent protein kinase in both particulate and soluble subcellular fractions, with highest specific activities in the synaptic membrane and cytosol fractions (47). Thus, in the brain cyclic AMP may mediate effects on membranes, cytosolic structures, nuclei, and other intracellular compartments (51). Recent findings have shown that type I1 cyclic AMP-dependent protein kinase can be bound, apparently through the R,,-subunits, to specific cytoplasmic proteins in the brain, including the cytoskeletal protein MAP-2 (microtubule-associated protein 2) (52, 53); the Ca2 -calmodulin-dependent protein phosphatase, calcineurin ( 5 4 , and several other unknown proteins (55, 56). This type of interaction is presumably important in localizing and concentrating the enzyme close to its physiological substrates. For example, about 30% of cytosolic cyclic AMP-dependent protein kinase appears to be bound to MAP-2 (52, 53, 56), a high-molecular-weight microtubule-associated protein (M,= 280,000). MAP-2 +

292

S . IVAR WALAAS AND PAUL GREENGARD TABLE I1 ACTIVATION OF SECOND MESSENGER-REGULATED PROTEIN KINASESI N BRAIN

Type of first messengeP Neurotransmitters Hormones Neurotransmitters Depolarizing stimuli Drugs Nerve impulse Depolarizing stimuli Neurotransmitters Hormones Neurotransmitters Hormones Depolarizing stimuli

Second messenger

Cofactor

Cyclic AMP, Type I Cyclic AMP, Type 11 Cyclic GMP

Cyclic AMP Cyclic GMP

Calcium

Protein kinaseh

Calmodulin

Ca2+-calmodulin I. Ca2 -calmodulin 11 Myosin light chain Phosphorylase CaZ -phospholipid +

Diacylglycerol (Inositol triphosphate) Calcium

Phospholipid

+

“For recent reviews on signal molecules that increase the levels of second messengers in brain, see for example, Drummond (83),Downes (I.?), and Nishizuka (84). hProtein kinase abbreviations: Ca2 +-calmoddin I, calcium-calmodulin-dependent protein kinase I; myosin light chain, myosin light chain kinase; phosphorylase, phosphorylase kinase; others as in Table 1.

is an important substrate for cyclic AMP-dependent protein phosphorylation in brain (see Section III,C,2). and this phosphorylation appears to regulate both its microtubule assembly-promoting activity (57, 58) and its interactions with both actin (59) and neurofilaments (60). (MAP-2 is also phosphorylated by Ca2 dependent protein kinases, and is discussed further in Section III,C,2). The general mechanisms involved in the regulation (Table 11) of cyclic AMPdependent protein kinase in brain appear to be similar to those described in peripheral tissues (37, 38, 40). Adenylate cyclase, the membrane-bound enzyme that catalyzes the formation of cyclic AMP, is controlled by receptor-mediated stimulation and inhibition [see, e.g., Ref. (61)],and, in the brain, also by CaZ+ and calmodulin (62). As discussed above, increasing amounts of cyclic AMP generated by this enzyme dissociate the inactive holoenzyme and release the active C-subunits, while decreasing amounts of cyclic AMP (following hydrolysis by cyclic nucleotide phosphodiesterases) lead to reassociation and regeneration of the inactive holoenzyme. Autophosphorylation of R,,, which is stimulated by cyclic AMP, retards reassociation of the type I1 isozyme, and thereby enhances its response to cyclic AMP (63). In addition to the R-subunits, which can be regarded as inhibitory C-subunitbinding proteins, many tissues including brain contain a heat-stable protein in+

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293

hibitor of cyclic AMP-dependent protein kinase, which can bind to and inactivate the free C-subunits (64).Changes in the concentration of this inhibitor have been reported in brain following drug treatment (65, 66), and may play a role in chronic regulation of cyclic AMP-dependent protein kinase activity. 2 . Cyclic GMP-Dependent Protein Kinase Cyclic GMP-dependent protein kinase, which was initially observed in invertebrate tissues (67), is today known to be present in several mammalian tissues, including brain (68-70). The brain enzyme, partially purified from cerebellum (37, 38, 71), has been found to have properties similar to the lung or heart enzyme (38, 69, 71). The enzyme consists of a dimer, each subunit (M, = 74,000) of which has two binding sites for cyclic GMP (72, 73) and a catalytic domain. Upon binding of cyclic GMP, a conformational change occurs, and the catalytic domain is exposed to the substrate (74).The enzyme undergoes cyclic GMP-stimulated autophosphorylation, which appears to increase the V,,, of the phosphotransferase reaction (71a-73). Cyclic GMP-dependent protein kinase has an uneven tissue distribution, with only a few peripheral tissues showing significant activity (70).In the brain, the cerebellum contains the highest activity (75, 76), and studies have shown that this is due to a high enrichment of the enzyme in the Purkinje cells (76, 77). Indeed, an immunohistochemical analysis of cyclic GMP-dependent protein kinase immunoreactivity in the cerebellum of the rat allowed a complete anatomical mapping and analysis of the architecture, structure, and projections of all Purkinje cells (77). Immunoreactivity was found throughout the cytosol, in dendrites, axons, and perikaryon, but not in the nucleus, which is in agreement with the cytosolic distribution of this enzyme in most tissues (78). Cyclic GMPdependent protein kinase has not been conclusively demonstrated in any other nerve cell type in the mammalian brain (37). However, cyclic GMP appears to function as second messenger in many neurons in addition to Purkinje cells (79, 80), and studies have shown effects of cyclic GMP-dependent protein kinase injected into pyramidal cells in the cortex that mimic the effects of acetylcholine or of intracellular injection of cyclic GMP (33). A low level of cyclic GMPdependent protein kinase immunoreactivity has also been reported to be present in the medium-sized spiny neurons of the neostriatum (81). Therefore, the enzyme may also be present in low amounts in other neurons. The activity of cyclic GMP-dependent protein kinase is regulated (Table 11) by the level of cyclic GMP in the cells. A variety of neurotransmitters, drugs, hormones, and changes in cellular activity, have been found to increase the formation of cyclic GMP by guanylate cyclase (12, 82, 83).This effect is usually Ca2 -dependent, and often appears to be interrelated with phosphatidylinositol turnover and Ca2+ mobilization (13, 14, 84-86), possibly through fatty acid +

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derivatives (87). Inactivation of cyclic GMP-dependent protein kinase appears to be caused by cyclic nucleotide phosphodiesterase-catalyzed hydrolysis of the cyclic nucleotide (88) in brain as in other tissues.

B . CALCIUM-DEPENDENT PROTEINKINASES In nervous tissue, a role for Ca2 -dependent protein phosphorylation was first suggested by studies using purified nerve terminals from cerebral cortex (89, 89a). These so-called synaptosomes were first preincubated with inorganic 32P to label the intracellular ATP pools. Subsequent increase of intraterminal Ca2 concentration, by depolarization with veratridine or high K concentrations, led to an increase in the 32P content of several proteins. These effects of veratridine and K+ were totally dependent on the presence of Ca2 in the external medium and were rapid, reversible, and concomitant with Ca2+ influx into the nerve terminals. In addition, the effects were independent of cyclic nucleotides (89, 89a). Further studies on broken-cell preparations confirmed and extended these findings. The addition of Ca2 to synaptosomal lysates stimulated the phosphorylation of several proteins (90), and subsequent analysis showed that this Ca2 -regulated protein phosphorylation was mediated by the ubiquitous Ca2 binding protein calmodulin (91-94). Ca2 and calmodulin-dependent protein phosphorylation appears to be particularly important in brain. The levels of Ca2 -calmodulin-dependent protein kinase activity are very high in this tissue, and a large number of brain-specific endogenous substrates have been discovered (35, 36, 95). A second type of Ca2 +-dependent protein phosphorylation in intact brain tissue, catalyzed by the Ca2 -phospholipid-dependent protein kinase, has been described (96). This protein phosphorylation system may mediate certain effects of those Ca2 -mobilizing hormones and neurotransmitters that increase phosphatidylinositol turnover (84). The Ca2 -phospholipid-dependent protein kinase is present in brain in very high concentration (36, 9 3 , where it catalyzes the phosphorylation of a distinct set of proteins that have a wide distribution throughout the central nervous system (35, 36, 95). +

+

+

+

+

+

+ -

+

+

+

+

+

1. Ca2 -Calmodulin-Dependent Protein Kinases +

Following the observation that synaptosomal membranes contain a Ca2 calmodulin-dependent protein kinase (91, 92), Sieghart and co-workers (98) showed that two of the major substrates for Ca2+-calmodulin-dependent protein phosphorylation in synaptosomes were identical to the proteins synapsin Ia and Ib. Synapsin Ia and Ib (collectively known as synapsin I, see Section III,C,l) were previously known to be two of the major substrates for cyclic AMPdependent protein kinase in the nervous system, and had been purified and +

295

9. PHOSPHORYLATION OF BRAIN PROTEINS

extensively characterized (99).Further studies showed that synapsin I contained one site (Site I) that was phosphorylated by cyclic AMP-dependent protein kinase and by a Ca2 -calmodulin-dependent protein kinase, and two additional sites (collectively termed Site 11) that were phosphorylated by a Ca2 -calmodulin-dependent protein kinase (100, 101). Further studies have established that two distinct Ca2 -calmodulin-dependent protein kinases are responsible for this site specificity (101-105). Ca2+-calmodulin-dependent protein kinase I (Ca2 calmodulin kinase I) specifically phosphorylates Site I of synapsin I, and Ca2+calmodulin-dependent protein kinase I1 (Ca2 -calmodulin kinase 11) specifically phosphorylates Site 11. These two enzymes probably represent two of the major Ca2 -calmodulin-dependent protein kinases in brain. Ca2+-calmodulin kinase I has been purified (103, 106) to apparent homogeneity from bovine brain using synapsin I as substrate. Ca2+-calmodulin kinase I had a K,,,for synapsin I (site I) of approximately 2-3 pM, which is similar to that of cyclic AMP-dependent protein kinase, and a K , of approximately 50 phf for ATP. In addition to synapsin I, Ca2 -calmodulin kinase I phosphorylated Protein I11 (see Section III,C,l) and smooth-muscle myosin light chain. In contrast, skeletal-muscle myosin light chain, glycogen synthase, histone, tubulin, and MAP-2 were not phosphorylated to any significant extent by the enzyme. The molecular weight of the native enzyme was estimated to be 49,000 from gel filtration studies. The purified enzyme preparation contained three peptides (M,= 42,000, 39,000, and 37,000 upon SDS-PAGE), of which all bound calmodulin and copurified with the kinase activity through a number of chromatographic procedures. In addition, they were all phosphorylated with low stoichiometry in a Ca2 -calmodulin-dependent manner. These and other results suggest that enzyme activity is associated with these three polypeptides (37). Ca2 -calmodulin kinase I, although found in highest concentration in the brain, appears to have a widespread tissue distribution ( 3 3 , and has been found in the cytosol from all rat tissues examined, including (in decreasing order of activity) forebrain, pancreas, spleen, lung, adrenal gland, heart, skeletal muscle, liver, and kidney. Ca2+-calmodulin kinase I also appears to be widely distributed in brain, where it is probably present in all parts of the neuron (A. c . Nairn and P. Greengard, unpublished results). Since synapsin I is present only in the nerve terminal, the widespread distribution of Ca2 -calmodulin kinase I suggests the existence of additional physiological substrates for this enzyme in both neuronal and nonneuronal mammalian tissues. The mechanism of activation of Ca2 -calmodulin kinase I in brain is believed to be similar to those of most Ca2+-calmodulin-dependent enzymes, (i.e., calmodulin will, in the presence of micromolar concentrations of Ca2+, undergo a conformational change and expose hydrophobic domains, which will bind to a calmodulin-binding domain on the enzyme) (107,108). Various studies have shown that Ca2+-calmodulin kinase I is stimulated in intact nerve cell prepara+

+

+

+

+

+

+

+

+

+

+

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S. IVAR WALAAS AND PAUL GREENGARD

tions upon depolarization-induced Ca2+ influx (e.g., into nerve terminals) (loo),suggesting that this kinase may be involved in regulation of nerve cell function (37). Ca2 -calmodulin kinase I1 has been purified from rat forebrain and characterized, using synapsin I as a substrate (102, 109, 110). Several other Ca2+calmodulin-dependent protein kinases have also been prepared from rat forebrain which have physicochemical properties very similar to those of Ca2 -calmodulin kinase I1 (111-115). Despite several differences in substrate specificity among these different enzyme preparations it is likely that the different enzyme preparations used in these studies all represent Ca2+-calmodulin kinase I1 (116), and they will therefore be discussed together. The purified enzyme exhibited a broad substrate specificity (109, 110) with synapsin I (site 11) being the best substrate tested. In addition, MAP-2, glycogen synthase, smooth-muscle myosin light chain, tau proteins, tyrosine hydroxylase, tryptophan hydroxylase, myelin basic protein, ribosomal protein S 6, and the Ca2 -calmodulin-sensitive cyclic nucleotide phosphodiesterase are relatively good substrates, while tubulin has been found to be a relatively poor substrate for most of the enzyme preparations (37, 116). The purified Ca2 -calmodulin kinase I1 had a molecular weight of 600,000650,000 as measured by gel filtration, and contained a major 50,000-dalton polypeptide and less prominent polypeptides of 60,000 and 58,000 daltons. The 60,000- and 50,000-dalton polypeptides, although immunologically related (110, 117), were shown to be distinct polypeptides, while the 58,000-dalton polypeptide may have been generated from the 60,000-dalton polypeptide by proteolysis (110). Several results suggest that enzyme activity is associated with all three subunits. Each protein was autophosphorylated in a Ca2 -calmodulindependent manner, each was shown to bind calmodulin using a 1251-calmodulinoverlay technique, and each bound ATP (37). Evidence suggests that Ca2 -calmodulin kinase I1 from the forebrain may be an isozyme of a multifunctional Ca2 -calmodulin-dependent protein kinase. For example, Ca2 -calmodulin kinase I1 purified from rat cerebellum, although having properties almost identical to the enzyme prepared from forebrain, contained predominantly 60,000- and 58,000-dalton polypeptides (110). Furthermore, a number of Ca2 -calmodulin-dependent protein kinases that have properties similar to those of Ca2 -calmodulin kinase I1 have been prepared from tissues other than brain, including electric organ of Torpedo californica (118), turkey erythrocytes (119), bovine heart (120), liver (121, 122), and skeletal muscle (123). Glycogen synthase kinase from skeletal muscle, for example, which has a substrate specificity identical to that of Ca2+ -calmodulin kinase I1 (124), has a native molecular weight of 800,000 and contains autophosphorylated subunits of 59,000, 58,000, and 54,000 daltons (123); and monoclonal +

+

+

+

+

+

+

+

+

+

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9. PHOSPHORYLATION OF BRAIN PROTEINS

antibodies prepared against Ca2 -calmodulin kinase I1 have been shown to crossreact with each subunit of glycogen synthase kinase (124). The similar physiocochemical and immunological properties and broad substrate specificities of these various Ca2 -calmodulin-dependent protein kinases strongly suggest that they are isozymes of a multifunctional Ca2 -calmodulin-dependent protein kinase. Ca2 -calmodulin kinase I1 is present in very high concentration in the brain, comprising perhaps as much as 0.4% of total brain protein (109, 110). The enzyme is found throughout the cell, being particularly enriched in dendrites (125) and postsynaptic densities (117, 126, 127). In postsynaptic densities (PSD) the major PSD protein (128) has been shown to be identical to the 50,000-dalton subunit of Ca2+-calmodulin kinase I1 (17, 126. 127). In a regional survey of Ca2 -calmodulin kinase I1 activity in rat brain, the enzyme was found to have a widespread though variable distribution (35, 36). High activity was found in cortical regions, particularly in the hippocampus and in most subcortical forebrain regions, whereas relatively low activity was found in the cerebellum, brain stem, and spinal cord. It is an interesting possibility that different isozymes of Ca2+-balmodulin kinase I1 may be responsible for the activity found in particular brain regions (116). The widespread distribution of Ca2+-calmodulin kinase I1 in brain and its broad specificity suggests that the enzyme mediates or modulates a variety of Ca2 -regulated mechanisms in the nervous system. Myosin light chain kinase and phosphorylase kinase, two Ca2 -calmodulindependent protein kinases which have been purified from several nonneuronal tissues and characterized extensively, have been identified in brain. These two enzymes have narrow substrate specificities and are presumably not involved in a wide variety of nerve cell functions. They are therefore discussed only briefly. Myosin light chain kinase has been purified from forebrain (54, 129) and found to have properties similar to those of the smooth-muscle enzyme. The purified kinase has a molecular weight of 130,000 and it specifically phosphorylates smooth-muscle myosin light chain (54). Although myosin has been isolated from brain, its precise function in this tissue is unknown (130). However, phosphorylation of smooth-muscle or nonmuscle myosin light chain is believed to be a prerequisite for interaction between myosin and actin (131), and myosin light chain kinase probably plays a role in brain analogous to its role in smooth-muscle and nonmuscle cells. Phosphorylase kinase has been identified in, but not purified from, brain (132, 133). The brain enzyme, which appeared to be similar to phosphorylase kinase from skeletal muscle, was activated both by Ca2+ and by phosphorylation by cyclic AMP-dependent protein kinase (133). The brain kinase, however, is only partly cross-reactive with antibody prepared against the enzyme from skeletal muscle, suggesting that it may be a distinct phosphorylase kinase isozyme (133). +

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+

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S. IVAR WALAAS AND PAUL GREENGARD

It is likely that Ca2+ activation of phosphorylase kinase regulates glycogen breakdown in neurons, a process which has been shown to be enhanced by electrical stimulation (134). 2. Ca2 -Phospholipid-Dependent Protein Kinase +

The Ca2 -phospholipid-dependent protein kinase (also known as Protein Kinase C) was first purified from cerebellum as a cyclic nucleotide-independent protein kinase which could be activated by a Ca2 -dependent protease also found in brain (135, 136). The enzyme was later found to be activated by the addition of membrane phospholipid (of which phosphatidylserine and phosphatidylinositol were the best tested), Ca2+, and low concentrations of diacylglycerol, which (under optimal conditions) was shown to decrease the apparent K, value for Ca2+ from approximately 7 X 1OP5Mto 5 X 10-6M (137) (see also Chapter 5 in Volume XVII). The Ca2 -phospholipid-dependent protein kinase has been purified from brain (138, 139) and other tissues (140, 141), and the properties of the enzyme from different sources appear to be similar. The enzyme from brain is a monomer with M, = 82,000 estimated from SDS-PAGE and M, = 87,000 estimated from gel filtration (138). The enzyme has a broad substrate specificity that is distinctly different from those of both cyclic nucleotide-dependent and Ca2 -calmodulindependent protein kinases. Myelin basic protein (138, 142), MAP-2 (36), an 87,000-dalton protein (36, 96),the B-50 protein (143), and several other unidentified proteins (35, 36) have been identified as possible physiological substrates for the enzyme in brain. In addition, histone H1 and many other nonneuronal proteins have been shown to be good substrates for the enzyme (138, 140). Ca2 -phospholipid-dependent protein kinase has a broad species, tissue, and cellular distribution (97, 144). The enzyme is highly concentrated and widely distributed in brain, with the highest activity found in cortical regions and in the cerebellum, and the lowest activity found in the brain stem and spinal cord (36). The subcellular distribution of the kinase in brain is of interest. In studies of the distribution of kinase activity in brain (35, 36) the enzyme appeared to be predominantly soluble. However, other workers have suggested that a significant amount of the enzyme in brain is particulate (138, 145), and that the particulate kinase differs from the soluble enzyme in that it will not phosphorylate exogenously added substrates without the addition of a detergent, for example, Triton X-100. The nature of the association of the particulate kinase with the membrane is not known. Ca2 -phospholipid-dependent protein kinase is activated by K - or veratridine-induced Ca2 influx in intact synaptosomes (96).Similarly, addition to synaptosomes of tumor-promoting phorbol esters, which substitute for diacylglycerol (146), or activation of receptors that induce phosphatidylinositol turnover (13, 14), will activate the enzyme (147). Physiological activity in intact +

+

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+

+

+

+

+

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9. PHOSPHORYLATION OF BRAIN PROTEINS

central nervous system preparations therefore appears to activate Ca2 -phospholipid-dependent protein kinase. The brain contains high levels of the enzyme as well as a large number of specific substrate proteins. It is therefore likely that Ca2 -phospholipid-dependent protein phosphorylation is involved in many functions in the brain. For example, recent studies have shown that tumorpromoting phorbol esters promote the release of neurotransmitters from rat brain synaptosomes (148). Similarly, application of phorbol esters, or intracellular injection of the Ca2 -phospholipid-dependent protein kinase, enhances voltagesensitive Ca2+ currents in bag cell neurons of Aplysiu (30), and phorbol esters regulate ionic currents in hippocampal pyramidal cells (31). This provides evidence for the involvement of this protein phosphorylation system in the regulation of neuronal function. +

+

+

C. CALCIUM-AND CYCLICNUCLEOTIDE-INDEPENDENT PROTEINKINASES Mammalian brain contains various other protein kinases that can phosphorylate endogenous brain proteins (37).Most of these enzymes have not been shown to be importantly involved in specific brain functions, and they are therefore mentioned only briefly. Casein kinase I and casein kinase I1 are protein kinases that preferentially phosphorylate acidic proteins like casein (149). Brain-specific endogenous substrates for these kinases have not been reported, and their functions in brain are not known (37). Myelin protein kinase has been reported as a protein kinase activity independent of the known second messengers, highly enriched in myelin fractions, and very active towards myelin basic protein (150, 151). However, the relationship of this enzyme to proteolytic fragments of Ca2 -phospholipid-dependent protein kinase (see Section 111,C,3) is not known, and the functional effects of myelin basic protein phosphorylation are unclear. Coated vesicle kinase has been reported in brain coated vesicle preparations (152). This protein kinase may be related to a Ca2 -calmodulin-dependent coated vesicle kinase (153). Neurofilament protein kinase, an enzyme that can phosphorylate the three neurofilament proteins, is also second messenger-independent (154, 155). Work by Sternberger et ul. (156) has suggested that abnormal neurofilament phosphorylation may take place in the neurofibrillary tangles present in neurons in Alzheimer’s disease. The physiological importance of neurofilament phosphorylation is not well understood, however. Another class of protein kinases present in brain and probably of great functional importance is represented by the tyrosine-specific protein kinases (157). These enzymes phosphorylate their protein substrates on tyrosine residues instead of on the more commonly phosphorylated serine or threonine residues. Three types of tyrosine-specific kinases have been reported in neuronal preparations. One is probably related to pp60c-src, a normal cellular protein which is +

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homologous to a gene product of the Rous sarcoma virus (158). Another is found in preparations of insulin receptors prepared from brain (159) and is comparable to the growth factor receptor-associated tyrosine kinases (15). Third, the nicotinic acetylcholine receptor from Torpedo electric organ is phosphorylated by a tyrosine-specific protein kinase (160). However, it is not clear whether this particular protein kinase is present in mammalian brain. The functions of the tyrosine-specific protein kinases in mammalian brain are also not well understood. Finally, two specific protein kinases found in neuronal cells appear to be regulated by physiological activity. Rhodopsin kinase, a protein kinase present in photoreceptors and specific for rhodopsin, phosphorylates rhodopsin upon exposure to light and may be involved in visual signal transduction (161, 162). Pyruvate dehydrogenase kinase, a mitochondria1 enzyme phosphorylating specifically the a-subunit of pymvate dehydrogenase (see Chapter 3 in this volume.) appears to be activated by repeated electrical stimulation in, for example, the isolated hippocampal slice (163), presumably caused by metabolic changes induced by the electrical stimulation (164). The increased state of phosphorylation of pyruvate dehydrogenase has been suggested to be related to nerve terminal calcium metabolism (165).

111.

Phosphoproteins in the Brain

A complete molecular understanding of physiological effects in brain that are mediated or modulated by protein phosphorylation obviously requires the identification and characterization of the brain proteins that are phosphorylated by the different brain protein kinases. There are two distinct approaches to such studies. One is to characterize the possible phosphorylation of known proteins that have either established functions (such as enzymes, receptors, and ion channels) or are well characterized but whose exact function is unclear. This approach has been used successfully in the case of major proteins such as identified enzymes (e.g., tyrosine hydroxylase) (166), microtubule-associated proteins (e.g., MAP-2 and tau factor) (167-169), and myelin basic protein (170), and of certain quantitatively minor proteins, such as neurotransmitter receptors or ion channels (171). This approach is limited, however, since most of the proteins involved in specific nerve cell brain functions have not been identified. The second approach is to search for previously unknown phosphoproteins by virtue of their ability to be phosphorylated upon in vitro incubation with [y3*P]ATP. This approach is potentially open-ended in the sense that, depending on the type of preparation used, it should ultimately be possible to detect most or all of the substrates for the different protein kinases. These protein substrates may then be characterized with respect to their biochemical, physio-

9. PHOSPHORYLATION OF BRAIN PROTEINS

30 1

logical, and anatomical properties, and hypotheses about their functional importance formulated and experimentally tested (6, 7). Examples of investigations of protein phosphorylation in brain where this open-ended approach has been taken include studies with intact synaptosomes, synaptosomal lysates, synaptic membranes, total cytosol, synaptic vesicles, and synaptic junctions and postsynaptic densities (89-91, 96, 100, 1.53, 172-174). One such study, which analyzed the patterns of protein phosphorylation in particulate and soluble fractions from different brain regions by one-dimensional SDS-PAGE revealed over 70 proteins that were substrates for cyclic AMP-dependent, cyclic GMP-dependent, and/or calcium-dependent protein kinases and that appeared to be enriched in or specific to nervous tissue (35, 36). Analysis of brain protein phosphorylation by twodimensional electrophoresis has revealed the existence of an even larger number of phosphoproteins in the nervous system (8).The large number and diversity of neuronal proteins that undergo phosphorylation supports the view that protein phosphorylation plays numerous and varied physiological roles in the nervous system. OF BRAIN PHOSPHOPROTEINS DISTRIBUTION A. REGIONAL

Distinct patterns of phosphoproteins are observed when discrete regions of the brain are analyzed (35, 36). Such studies on the regional distribution of neuronal phosphoproteins have indicated that they can conveniently be divided into three categories, according to their patterns of regional distribution (35, 36). Category A phosphoproteins are widely and fairly evenly distributed throughout the nervous system, and presumably are involved in functions common to all nerve cells. Category B phosphoproteins are relatively widely, but unevenly, distributed throughout the nervous system; these phosphoproteins may be enriched in certain classes of nerve cells, and present in low levels or absent from other nerve cells. They may be importantly involved in functions restricted to classes of neurons. Category C phosphoproteins are restricted to one brain region only, where these proteins are presumably involved in functions specific to a single cell type. The distribution patterns for certain neuron-specific phosphoproteins have been studied in considerable detail. Category A phosphoproteins include the synaptic vesicle-associated proteins synapsin la, synapsin Ib, Protein IIla, and Protein IIIb. They appear to be present in virtually all axon terminals throughout the nervous system, and their development coincides with synaptogenesis. These synaptic vesicle-associated phosphoproteins have proven useful as general markers for the study of axon terminals in both developing and adult brain (8, 175). Furthermore, their presence in virtually all nerve terminals, and the regulation of their phosphorylation by neurotransmitters that act through second messengers, has made it possible to use these phosphoproteins to characterize presynaptic

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neurotransmitter receptors in various brain regions (176). Synapsin I and Protein I11 are discussed further in Section III,C, 1. Category B phosphoproteins include DARPP-32, a dopamine- and cyclic AMP-regulated phosphoprotein of M, 32,000 (see Section III,C,4), which is localized to dopaminoceptive neurons that possess the D, -receptor (dopamine receptor linked to adenylate cyclase) (177, 178). Anatomical studies of DARPP-32 have already revealed an intricate and specific distribution pattern for such dopaminoceptive neurons throughout the brain (178). Another Category B phosphoprotein is tyrosine hydroxylase (166), the rate-limiting enzyme in the biosynthesis of catecholamines. This phosphoprotein is restricted to nerve cells that utilize catecholamines as neurotransmitters, and has been used to map the localization of catecholaminergic neurons in the nervous system (179). Other examples of Category B phosphoproteins are the M, 50,000 and 60,000 autophosphorylated subunits of calcium and calmodulin-dependent protein kinase I1 described in Section II,A,l. The distribution of this protein kinase in brain is very similar to that of sodium-independentglutamate binding sites, which probably represent glutamate receptors (180). Since activation of glutamate receptors leads to calcium flux into neurons (184, this protein kinase may be involved in mediating certain of the transsynaptic effects of glutamate in the brain. Two substrate proteins specific to the cerebellar Purkinje cell are examples of Category C phosphoproteins. These are a cytosolic substrate for cyclic GMPdependent protein kinase (G-substrate), and a particulate substrate for cyclic AMP-dependent protein kinase (182, 183). In addition, as discussed in Section II,A,2, cyclic GMP-dependent protein kinase is also highly enriched in Purkinje cells. These three proteins therefore represent unique tools with which to analyze Purkinje cell anatomy and development. Furthermore, identification of the functions of these Category C phosphoproteins should reveal functional characteristics specific to this type of neuron.

B. CLASSESOF NEURONAL PHOSPHOPROTEINS A tentative classification of those neuronal phosphoproteins that have been most intensively studied and whose functions are at least partly known is presented in Table I11 (184-196). They include enzymes involved in neurotransmitter biosynthesis, enzymes involved in cyclic nucleotide metabolism, autophosphorylated protein kinases, protein phosphatase inhibitors, cytoskeletal proteins, synaptic vesicle-associated proteins, neurotransmitter receptors, and ion channels. Many of these proteins have unknown distribution. However, the functional roles of some of these phosphorylation reactions have already been established (7). Some general points are summarized here. Autophosphorylated protein kinases are among the most prominent substrates

TABLE I11 NEURONAL PROTEINS REGULATED BY PHOSPHORYLATION~,~ Proteinsa Enzymes involved in neurotransmitter biosynthesis Tyrosine hydroxylase Tryptophan hydroxylase Enzymes involved in cyclic nucleotide metabolism Adenylate cyclase Guanylate cyclase Phosphodiesterase Autophosphorylated protein kinases Cyclic AMP-dependent protein kinase Cyclic GMP-dependent protein kinase Calcium-calmodulin-dependent protein kinases Calcium-phospholipid-dependent protein kinase (protein kinase C) Casein kinases Tyrosine-specific protein kinases Rhodopsin kinase Protein phosphatase inhibitors Phosphatase inhibitor- 1 DARPP-32 G-substrate Cytoskeletal proteins Tubulin Tau factor Neurofilaments Fodrin Myosin light chain Synaptic vesicle-associated proteins Synapsin I Protein 111 Neurotransmitter receptors Nicotinic acetylcholine receptor

Muscarinic acetylcholine receptor P-adrenergic receptor Ion channelsC Sodium channel Potassium channels Calcium channels

References

(231, 232, 234) (227, 228) (190, 191, 233) (214, 215) (169) (154, 155) (112) (129, 131) (99, 100) (208, 211)

(I60, 193, 194

I

197, 198) (192) (199, 200) (195, 196) (18-25) (26-28, 30)

“Modified from Browning et al. (5) and Nestler et al. (8). bSome of the proteins included are specific to neurons. The others are present in many cell types in addition to being present in neurons and are included because the regulation of neuron-specific phenomena is among their multiple functions in the nervous system. Not included are many phosphoproteins present in diverse tissues (including brain) that play roles in generalized cellular processes, such as intermediary metabolism, and that do not appear to play roles in neuron-specific phenomena. =Several types of ion channels have been shown to be physiologically regulated by protein phosphorylation reactions, although it is not yet known whether such regulation is achieved directly through the phosphorylation of the ion channel, or indirectly through the phosphorylation of a modulatory protein that is not a constituent of the ion channel molecule.

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observed for kinases in endogenous phosphorylation assays in nervous and nonnervous tissue. Thus, prominent substrates for protein kinases activated by cyclic AMP, cyclic GMP, calcium, epidermal growth factor, and insulin have been shown to be autophosphorylated subunits of the respective protein kinases in certain target cells (7). It appears that autophosphorylation of protein kinases is the rule rather than the exception, although the physiological role of autophosphorylation is not clear. It seems likely, however, that the phosphorylated moieties of the protein kinases are involved in the regulation of kinase activity. Indeed, those phosphorylated moieties may be compared to inhibitors of protein phosphatases, which also are among the most prominent phosphoproteins that have been found in brain, in that both groups of phosphoproteins represent regulatory components of the protein kinases and protein phosphatases (8). Several neurotransmitter receptors have been reported to be associated with protein phosphorylation systems (Table III), although the exact nature of the association may vary. For example, the nicotinic acetylcholine receptor from Torpedo electroplax is rapidly and specifically phosphorylated on the y and 6 subunits by cyclic AMP-dependent protein kinase, on the 6 subunit by Ca2 phospholipid-dependent protein kinase, and on the p, y, and 6 subunits by a tyrosine-specific protein kinase endogenous to Torpedo electroplax (160, 197, 198). As another example, the P-adrenergic receptor is phosphorylated in intact turkey erythrocytes, where it appears to be a substrate for both cyclic AMPdependent and Ca2 -phospholipid-dependent .protein kinases. These phosphorylation reactions may mediate desensitization of the receptor (199, 200). Receptors for several types of steroid hormones, including estradiol, progesterone, and glucocorticoids, are also phosphorylated by various protein kinases in vitro or in vivo; and membrane receptors for immunoglobulin E, plasma glycoproteins, and transferrin are also phosphoproteins (7). The large number and diversity of receptors that undergo phosphorylation suggests that regulation of receptor function by phosphorylation is a common and physiologically important property of a wide variety of receptors. Many types of ion channels are also regulated by phosphorylation (Table 111). Some neurotransmitter-dependent ion channels, like the nicotinic acetylcholine receptor, are “receptor-ion channels” in that the neurotransmitter receptor and the ion channel involved are tightly coupled and reside in the same physical complex. With other neurotransmitter-dependent ion channels, the neurotransmitter receptor and the ion channel are distinct entities; the neurotransmitter regulates ion channel function indirectly through some second messenger. The diversity of ion channels altered by phosphorylation suggests that regulation of ion channel function by phosphorylation is a common and physiologically important property of a wide variety of ion channels [see Ref. (8)and Chapter 1 1, this volume]. +

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C. PHOSPHORYLATION OF SPECIFIC BRAINPROTEINS In this section, we describe the phosphorylation of some selected examples of major brain proteins in more detail. This is done in order to demonstrate the differences between various brain phosphoproteins, and the diversity of presumed functions they are involved in. Some of the proteins we discuss are related either to synaptic vesicles, to microtubules, or to myelin; others are cytosolic, cell-specific inhibitors of protein phosphatases; and still others are nerve terminal substrates specific for Ca2 -phospholipid-dependent protein kinase. +

1 . Synaptic Vesicle-Associated Proteins Synapsin I (previously called Protein I), which consists of two polypeptides of

M,86,000 and 80,000designated synapsin la and b, is a major brain substrate for cyclic AMP-dependent and Ca2 +-dependent protein kinases. It is found only in nervous tissue, where it is enriched in virtually all nerve terminals and appears to be associated with the external surface of synaptic vesicles (201, 202). It represents a major neuron-specific protein, making up about 0.4% of total brain protein, 1% of neuronal protein, and 6% of synaptic vesicle protein (175, 203). It appears to consist of a proline-rich “tail-region’’ and a globular “head-region,’’ both of which are phosphorylated. Cyclic AMP-dependent protein kinase and Ca2+-calmodulin kinase I phosphorylate Site I in the head region and Ca2+calmodulin kinase I1 phosphorylates Site I1 in the tail region (see Section II,B,I). Several studies have demonstrated that both Sites I and I1 in synapsin I can be phosphorylated in various intact tissue preparations. Examples include intact synaptosomes and brain slices (100, 204, 205), the superior cervical ganglion (206, 207), and the posterior pituitary (208). The protein was phosphorylated by application of neurotransmitter, by chemical depolarization and by electrically induced impulse conduction. In these studies the incorporation of 32P into both Sites I and I1 was found to be rapid, and reversible within seconds following termination of the stimulus. Synapsin I is bound through its tail region (209a) (which contains Site 11) to a specific, high affinity, saturable site on synaptic vesicles (203, 209). Phosphorylation of Site I1 has recently been found to reduce this binding (203, 209). Thus, the interaction between synapsin I and synaptic vesicles appears to be regulated by Ca2 -calmodulin kinase 11. Moreover, this interaction appears to be involved in the regulation of neurotransmitter release, since microinjection of synapsin I or Ca2 -calmodulin kinase I1 into presynaptic nerve terminals of the squid giant synapse potently modified the stimulated release of transmitter at this synapse (32). Protein I11 consists of Protein IIIa and Protein IIIb (M,= 74,000 and 55,000, respectively), which also are major substrates for cyclic AMP-dependent and Ca2+-dependent protein kinases in brain (210-212). Protein 111 appears to be a +

+

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synaptic vesicle-associated protein that shares some properties with synapsin I (212). It can be phosphorylated in intact cells by several neurotransmitters and stimuli which can increase cyclic AMP or Ca2+ levels (212). This phosphorylation appears to take place on one amino acid residue, and is catalyzed by cyclic AMP-dependent protein kinase or Ca2+-calmodulin kinase I (37). The function of Protein 111 in synaptic transmission is unknown. 2. Microtubule Proteins Microtubules are present throughout the nerve cell, both in association with postsynaptic membranes and in dendrites, somata, axons, and presynaptic terminals (128, 213). Microtubules consist of tubulin and microtubule-associated proteins (MAPS), which are phosphorylated by Ca2 -dependent and cyclic AMP-dependent mechanisms. Tubulin has been reported to be phosphorylated by both cyclic AMP-dependent and Ca2+-dependent protein kinases (169). For example, activation of endogenous Ca2 -calmodulin-dependent protein kinase activity in cytosol fractions, synaptosomes, and synaptic vesicles induced phosphorylation of two polypeptides which had apparent molecular weight similar to those of a-tubulin and P-tubulin (173, 214, 215). A brain Ca2 -calmodulin-dependent protein kinase that phosphorylates tubulin has also been purified and characterized (113). This protein kinase appears, however, to be similar to Ca2+-calmodulin kinase 11, an enzyme for which tubulin has been found to be a poor substrate (116). Studies have indicated that phosphorylation of tubulin by Ca2 -calmodulin kinase I1 but not by cyclic AMP-dependent kinase may decrease microtubule assembly (169). The significance of tubulin as a brain phosphoprotein is, however, not well understood (7). Microtubule-Associated Protein 2 (MAP-2), a high-molecular-weight protein (M, = 280,000) is enriched in neurons, where it is present mostly in dendrites (216, 217). MAP-2 was originally found to be a substrate for cyclic AMPdependent protein kinase (218) and subsequent studies have shown that the protein is also phosphorylated by both Ca2 -calmodulin kinase I1 and Ca2 phospholipid-dependent protein kinase in brain cytosol(36, 109, 110, 113). The protein is phosphorylated on multiple sites, the number of which has been reported to vary between 1-2 per molecule and up to 20-22 sites per molecule (218, 219). Phosphorylation of purified MAP-2 with Ca2+-calmodulin kinase I1 led to 32P-incorporation into at least 5-15 sites, which appeared to be distinct from those phosphorylated by cyclic AMP-dependent protein kinase (169, 220). However, methods capable of studying changes in the state of phosphorylation of the protein in intact tissue have not been reported, and the extracellular signals that might regulate MAP-2 phosphorylation have not been identified. Phosphorylation of MAP-2 by Ca2 -calmodulin kinase I1 or cyclic AMP-dependent protein kinase has been reported to induce disassembly of microtubules (169, +

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220). Phosphorylation of MAP-2 may therefore represent an important mechanism for the control of the assembly-disassembly cycle of microtubules. Tau factor is a group of microtubule-associated proteins (M,= 60,00055,000) which have been reported to be phosphorylated on multiple sites by Ca2 -calmodulin kinase I1 and cyclic AMP-dependent protein kinase (169). This phosphorylation appears to inhibit microtubule assembly, and tau factor may therefore play a role in microtubule function similar to that of MAP-2. +

3 . Myelin Basic Protein Myelin basic protein is a major component of myelin sheets, which in the brain are generated by oligodendrocytes and are wrapped around axons to insulate them and increase conduction efficiency. Several studies have shown that myelin basic protein is phosphorylated on multiple sites in vivo (221, 222). The protein kinase(s) responsible for this phosphorylation has not been conclusively identified. Myelin contains small amounts of cyclic AMP-dependent protein kinase (142), but myelin basic protein is not a good substrate for this enzyme in vitro (223). In contrast, myelin basic protein may be a substrate for Ca2+calrnodulin kinase I1 in vitro (113). More recently, however, brain myelin was reported to contain a highly active Ca2 -phospholipid-dependent protein kinase, which phosphorylated myelin basic protein in intact membranes, detergent extracts, and a partially purified preparation. These studies revealed no evidence for a Ca2 -calmodulin-dependent phosphorylation of myelin basic protein (142). Furthermore, a specific site phosphorylated in myelin basic protein in vivo (222) appears to correspond to a site phosphorylated by Ca2+phospholipid-dependent protein kinase in vitro (224). Further studies have shown that the state of phosphorylation of the protein is increased when myelinated axons in the optic nerve are depolarized with high K + or electrical stimulation (225, 226). However, the functional significance of myelin basic protein phosphorylation is unknown. +

+

4. Cell-Specific Protein Phosphatase Inhibitors DARPP-32 is an acid-soluble and heat-stable cytosolic phosphoprotein which is phosphorylated in intact nerve cells in response to dopamine or 8-bromo-cyclic AMP, and in broken cell preparations by endogenous cyclic AMP-dependent protein kinase (227). As described in Section III,A, the protein appears to be enriched in dopamine-innervatedneurons, particularly in the basal ganglia (173, and specifically in those dopaminoceptive cells which have D, -dopamine receptors (dopamine receptors stimulating adenylate cyclase) ( 1 78).It is phosphorylated on a single threonine residue (228, 229) which is surrounded by an amino acid sequence (230) very similar to those found in phosphatase inhibitor-1 (231, 232) and in G-substrate (233) (see following paragraph). Phosphorylated, but not dephosphorylated, DARPP-32 functions as a potent inhibitor of protein phos-

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phatase-1 (234).Therefore, DARPP-32 may be involved in mediating or modulating some transsynaptic effects of dopamine acting on D, -receptors by regulating the activity of protein phosphatase-I. In particular, DARPP-32 may be involved in the interaction between dopamine and other first messengers (e.g., glutamate) (8),since DARPP-32 would be able to modulate the state of phosphorylation of phosphoprotein substrates that are regulated by protein kinases activated by those first messengers. G-substrate is an acid-soluble and heat-stable protein (M,= 23,000) highly enriched in the cytosol of cerebellar Purkinje cells (182), which is phosphorylated by cyclic GMP-dependent protein kinase on two threonine residues. These residues have surrounding amino acid sequences very similar to those of DARPP-32 and phosphatase inhibitor- 1 (233).Furthermore, the phosphorylated form of G-substrate has been found to inhibit both protein phosphatase-1 and protein phosphatase-2A from skeletal muscle (37). Thus, G-substrate, like DARPP-32, may function as a neuronal cell-specific protein phosphatase inhibitor (8). 5.

Ca2 -Phospholipid-Regulated Nerve Terminal Proteins +

Studies of intact synaptosome preparations prelabelled with 32P have shown that depolarization-induced calcium influx or addition of the tumor-promoting phorbol esters results in the phosphorylation of an 87,000-dalton protein (96, 147). Further studies using crude cytosol or membrane preparations showed that this 87,000-dalton protein could be phosphorylated by the addition of Ca2+ plus phosphatidylserine, but not Ca2 -calmodulin or cyclic nucleotides (36, 96). These results suggest that the 87,000-dalton protein is a physiological and specific substrate for Ca2 -phospholipid-dependent protein kinase in brain. The 87,000-dalton phosphoprotein, which has been purified to homogeneity (139), appears to be widely distributed throughout the brain (35, 36). Recent studies have shown that it also may be present in low quantities outside the nervous system (K. A. Albert, S. I. Walaas, J . K. T. Wang, and P. Greengard, unpublished observations). Subcellular fractionation of rat cerebral cortex and cerebellum has shown that it is enriched in the crude synaptosomal fraction, and is present in the cytosol and membrane fractions (35, 36, 96). It is phosphorylated by endogenous and exogenous purified Ca2 -phospholipid-dependent, but not by Ca2 -calmodulin-dependent or cyclic nucleotide-dependent, protein kinase (36, 96, 139). The physiological function of the 87,000-dalton protein is not known. Extensive studies have indicated that activation of the Ca2 -phospholipid-dependent protein kinase in blood platelets, chromaffin cells, or pancreatic islets may be an important molecular mechanism for the regulation of hormone release (84, 235, 236). By analogy, Ca2 -phospholipid-dependent phosphorylation of the +

+

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87,000-dalton protein in nerve terminals may be involved in Ca2 -dependent modulation of the release of CNS neurotransmitters. B-50 is a brain-membrane protein of 48,000 daltons that is phosphorylated by an endogenous membrane protein kinase in a reaction which originally was found to be inhibited by high concentrations of ACTH (237). The B-50 protein appears to be a distinct, specific phosphoprotein substrate for Ca2 -phospholipid-dependent protein kinase, and is not phosphorylated by either cyclic AMP-dependent or Ca2 -calmodulin-dependent protein kinase (36, 143, 237). Incubation of intact synaptosomes with depolarizing concentrations of K or with tumor-promoting phorbol esters have been found to increase the state of phosphorylation of B-50 (147). The B-50 protein has been localized specifically to nervous tissue by both biochemical and immunochemical methods, and appears to be associated with presynaptic plasma membranes, possibly as an integral membrane protein (238-240). Partially purified B-50-protein kinase complex was reported to have phosphatidylinositol 4-phosphate kinase activity, and this activity appeared to be regulated by the state of phosphorylation of B-50 (241). Therefore, the Ca2+dependent phosphorylation of B-50 may be involved in the regulation of the phosphatidylinositol-linked type of signal transmission (13) in the brain. +

+

+

+

IV. Protein Phosphatases in the Brain Although protein phosphorylation systems in the brain in most cases appear to be rapidly regulated by activation of protein kinases, evidence suggests that protein phosphatases may also constitute targets for regulatory agents (242245). The protein phosphatases involved in cellular regulation have been divided into four enzymic activities (244, 245), which can be classified into two types. Type 1 phosphatases (protein phosphatase- 1) selectively dephosphorylate the psubunit of phosphorylase kinase and are inhibited by phosphatase inhibitor- 1 or phosphatase inhibitor-2, while type 2 protein phosphatases (protein phosphatase-2A, -2B, and -2C), selectively dephosphorylate the &-subunit of phosphorylase kinase, and are insensitive to these inhibitors (232, 233). Recently, phosphotyrosyl-protein phosphatases have also been found in various tissues; these appear to be distinct from the main phosphoseryl and phosphothreonyl protein phosphatases (246, 247). (For further discussion about the general properties of protein phosphatases, see Chapter 8 in Vol. XVII). Protein phosphatase-1, -2A, -2B, -2C and the phosphotyrosyl phosphatase activities have all been found in brain (248, 249). Furthermore, a mitochondria1 phosphatase dephosphorylating pyruvate dehydrogenase has been reported (250), and phosphatase activities are also present in myelin (251, 252) and in partially purified

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preparations of rhodopsin (253), nicotinic acetylcholine receptor (194), and MAP-2 (254). The exact identities of these phosphatase activities are not known, however. The presence in brain of protein phosphatase-1, -2A, -2B, and -2C indicates that these enzymes probably dephosphorylate distinct brain phosphoproteins. Protein phosphatase-1, -2A, and -2C have relatively broad substrate specificities, while the substrate specificity of protein phosphatase-2B (also known as calcineurin) is more restricted (255, 256). Interestingly, phosphatase-2B (calcineurin) which is a Ca2+ -calmodulin-dependent enzyme, has been found to dephosphorylate several neuronal phosphoproteins, including DARPP-32 and Gsubstrate, with high efficiency (256). Thus, the state of phosphorylation of these two brain phosphoproteins may be regulated both by cyclic nucleotides (activating protein kinases) and by Ca2+ and calmodulin (activating a protein phosphatase) (8). It is particularly interesting that the phosphorylated forms of DARPP-32 and G-substrate, in turn, are potent inhibitors of protein phosphatase-1 (see Section III,C,4). This suggests that protein phosphatase-2B in the brain may be involved in modulating the activity of other protein phosphatases, thereby extending its influence to phosphoproteins which are not direct substrates for the enzyme itself. These results indicate that the regulation of protein phosphatase activity may be particularly important in the brain. Such regulation appears to be anatomically differentiated, since phosphatase inhibitor-1 is present throughout the brain, while DARPP-32 and G-substrate are region- and cell-specific (see Section III,C,4). These findings indicate that certain neurotransmitters may produce some of their physiological effects in brain by regulating protein phosphatases in specific cells, in some cases by molecular mechanisms that may be specific to nervous tissue.

V. Conclusion The overriding importance of reversible protein phosphorylation in cellular regulation is generally acknowledged. The data reviewed in this chapter demonstrate the existence of a rich variety of protein kinases, protein phosphatases, and phosphoprotein substrates in the mammalian brain, and summarize evidence for regulation of these systems by various regulatory agents such as neurotransmitters, hormones, and the nerve impulse itself. The data show that brain protein phosphorylation is involved in a variety of nerve cell functions, including specific informational processes related both to rapid signal transmission and, possibly, to long-term processes that may be associated with memory and learning (257, 258). The many individual molecular pathways involving protein phosphorylation

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that have been demonstrated in nervous tissue, and the numerous interactions between these pathways, attest to the extraordinary complexity of physiological processes at the molecular level in the nervous system. Identification of the specific phosphoprotein(s) involved in a particular biological response is crucial to the elucidation of the molecular pathway through which that response is achieved. A large number of phosphoproteins have been found in nervous tissue, and in several instances their functional role is known. These results suggest that further studies of neuronal phosphoproteins, including the characterization of their biochemical, anatomical, and physiological properties will greatly improve our understanding about the molecular basis of nervous system function.

REFERENCES I . Shepherd, G. M. (1979). “The Synaptic Organization of the Brain.” Oxford Univ. Press, London and New York. 2. Kuffler, S. W., Nicholls, J. G., and Martin, A. R. (1984). “From Neuron to Brain.” Sinauer, Sunderland, Massachusetts. 3. Shepherd, G. M. (1983). “Neurobiology.” Oxford Univ. Press, London and New York. 4. Greengard, P. (1978). Science 199, 146. 5. Browning, M. D., Huganir, R., and Greengard, P. (1985). J . Neurochem. 45, 1 1 . 6. Nestler, E. J . , and Greengard, P. (1983). Nature (London) 305, 583. 7. Nestler, E. J . , and Greengard. P. (1984). “Protein Phosphorylation in the Nervous System.” Wiley, New York. 8. Nestler, E. J., Walaas, S. I., and Greengard, P. (1984). Science 225, 1357. 9. Greengard, P. (1976). Nature (London) 260, 101. 10. Greengard, P. (1978). “Cyclic Nucleotides, Phosphorylated Proteins, and Neuronal Function.” Raven Press, New York. 11. Greengard, P. (1981). Harvey Lect. 75, 277. 12. Goldberg, N. D., and Haddox, M. K. (1977). Annu. Rev. Biochem. 46, 823. 13. Downes, C. P. (1982). Cell Calcium 3, 413. 14. Berridge, M. 3. (1984). BJ220, 345. 15. Cobb, M. H., and Rosen, 0. M. (1984). BBA 738, 1. 16. Cohen, S . , Carpenter, G., and King, L. (1980). JBC 255, 4834. 17. Bloom, F. E. (1975). Rev. Physiol. Biochem. Pharmacol. 74, 1. 18. Castellucci, V., Kandel, E. R., Schwartz, J. H., Wilson, F. D., Nairn, A. C., and Greengard, P. (1980). PNAS 77, 7492. 19. Castellucci, V . , Nairn, A. C., Greengard, P., Schwartz, J. H., and Kandel, E. R. (1982). J . Neurosci. 2, 1673. 20. Siegelbaum, S. A., Camardo, J. S., and Kandel, E. R. (1982). Nature (London) 299, 413. 21. Kaczmarek, L. K., Jennings, K. R., Strumwasser, F., Nairn, A. C., Walter, U., Wilson, F. D., and Greengard, P. (1980). PNAS 77, 7487. 22. Kaczmarek, L. K . , Nairn, A. C., and Greengard, P. (1984). SOC. Neurosci. Absrr. 10, 895. 23. Adams, W. B., and Levitan, I. B. (1982). PNAS 79, 3877. 24. Benson, J. A , , and Levitan, I. B. (1983). PNAS 80, 3522. 25. de Peyer, J. E., Cachelin, A. B., Levitan, I. B., and Reuter, H. (1982). PNAS 79, 4207. 26. Doroshenko, P. A., Kostyuk, P. G., Martynyuk, A. E., Kursky, M. D., and Vorobetz, Z. D. (1984). Neuroscience 11, 263.

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232. Nimmo, G . A., and Cohen, P. (1978). EJE 87, 341. 233. Aitken, A,, Bilham, D., Cohen, P . , Aswad, D. W., andGreengard, P. (1981). JEC256,3501. 234. Hemmings, H. C., Jr., Greengard, P., Tung, H. Y. L., and Cohen, P. (1984). Nature (London) 310, 503. 235. Baker, P. F., and Knight, D. E. (1984). Trends Neurosci. 7, 120. 236. Tanigawa, K., Kuzuya, H., Irnura, H., Taniguchi, H., Baba, S., Takai, Y., and Nishizuka, Y. (1982). FEES Lett. 138, 183. 237. Zwiers, H., Veldhuis, H. D., Schotman, P., and Gispen, W. H. (1976). Neurochem. Res. 1, 699. 238. Sorenson, R. G., Kleine, L. P., and Mahler, H. R. (1981). Erain Res. Bull. 7, 57. 239. Kristjansson, G . I . , Zwiers, H., Oestreicher, A. B., and Cispen, W , H. (1982). J . Neurochem. 39, 37 I . 240. Oestreicher, A. B., Zwiers, H . , Schotrnan, P., and Gispen, W. H. (1981). Bruin Res. Bull. 6, 145. 241. JollBs, J., van Dongen, C. J., Schotman, P., Wirtz, K. W. A,, and Gispen, W. H. (1980). Nature (London) 286, 623. 242. Li, H.-C. (1982). Curr. Top. Cell. Regul. 21, 129. 243. Cohen, P. (1982). Nature (London) 296, 613. 244. Ingebritsen, T. S., and Cohen, P. (1983). Science 221, 331. 245. Ingebritsen, T. S., and Cohen, P. (1983). EJE 732, 255. 246. Horlein, D., Gallis, B., Brautigan, D. L., and Bomstein, P. (1982). Biochemistry 21, 5577. 247. Foulkes, J. G., Erikson, E., and Erikson, R. L. (1983). JEC 258, 431. 248. Ingebritsen, T. S., Stewart, A. A., and Cohen, P. (1983). EJE 132, 297. 249. Wallace, R. W., Tallant, E. A , , and Cheung, W. Y. (1980). Biochemistry 19, 1831. 250. Sheu, K.-F. R . , Lai, J. C. K., and Blass, J. P. (1983). J. Neurochem. 40, 1366. 251. Miyamoto, E . , and Kakiuchi, S . (1975). EEA 384, 458. 252. McNamara, J. O., and Appel, S. H. (1977). J. Neurochem. 29, 27. 253. Goridis, C., and Weller, M. (1976). Adv. Eiochem. Psychopharmucol. 15, 391. 254. Coughlin, B. A,, White, H. D., and Purich, D. L. (1980). EERC 92, 89. 255. Stewart, A. A., Ingebritsen, T. S . , Manalan, A., Klee, C. B., and Cohen, P. (1982). FEES Lett. 137, 80. 256. King, M. M., Huang, C. Y., Chock, P. B., Naim, A. C., Hemmings, H. C., Jr., Chan, K.-F. J . , and Greengard, P. (1984). JEC 259, 8080. 257. Greengard, P., and Kuo, J. F. (1970). Adv. Eiochem. Psychopharmacol. 3, 287. 258. Kandel, E. R., and Schwartz, J. H. (1982). Science 218, 433.

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10

Regulation of Receptor Function JEFFREY L. BENOVIC ROBERT J. LEFKOWITZ Howard Hughes Medical insiiiute Deparimenis of Medicine (Cardiology) and Biochemistry Duke University Medical Center Durham, North Carolina 27710

I. Introduction and Perspectives ................................. 11. The P-Adrenergic Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Heterologous Desensitization B. Homologous Desensitization .................................... 111. Rhodopsin .................................... IV. The Nicotinic Acetylcholine Receptor ............................... ................. V. The Receptors for EGF and Insulin . . . . . VI. Other Membrane Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . .................

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Introduction and Perspectives

Receptors for hormones, drugs, and other biologically active molecules are crucially positioned to serve as important points of cellular metabolic regulation. They represent the critical locus of interaction of ligands with cells. Thus receptors control the responsiveness of cells to hormonal and pharmacological agonists or the entrance of key substances into cells via receptor-mediated endocytosis pathways. Mechanisms for regulation of receptor function have obvious potential for controlling these important cellular metabolic events. 319 THE ENZYMES, Vol. XVllI Copyright 0 1987 by Academic Press, Inc. All rights of reproduction in any form reserved

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The diverse and widespread role of phosphorylation in regulating the function of many enzymes is amply demonstrated by previous chapters in this volume. Recognition of enzyme regulation preceded appreciation of the role of protein phosphorylation in the regulation of receptor function. A key to progress in this area has been the ability to study the receptors for a wide variety of biologically active substances by techniques such as ligand binding, affinity chromatography, and photoaffinity labeling. This has permitted the study of the modification of receptor structure and function by phosphorylation. In this chapter we focus on the phosphorylation of several important plasma membrane receptors: the P-adrenergic receptor coupled to adenylate cyclase; rhodopsin, the archetypal “light” receptor of the rod outer segment; the nicotinic cholinergic receptor; the IgE receptor; and the transferrin receptor, which is involved in the internalization of iron-transferrin complexes. In addition, several cell surface receptors that possess tyrosine kinase activity (e.g., insulin and EGF receptors) are briefly discussed since they are dealt with in detail elsewhere in this volume. These specific cases provide examples of several mechanisms by which phosphorylation may regulate receptor function.

II. The f3-Adrenergic Receptor The P-adrenergic receptor is an ubiquitous plasma membrane glycoprotein that mediates catecholamine stimulation of the enzyme adenylate cyclase [reviewed in Refs. (1, 2 ) ] . This stimulation occurs via an agonist-induced, GTPdependent, interaction of receptor and N,, the stimulatory guanine nucleotide regulatory protein; N, can then directly activate adenylate cyclase leading to increased intracellular cAMP levels and subsequent activation of CAMP-dependent protein kinase. The P-adrenergic receptor is the only one of the adenylate cyclase-coupled receptors that has been purified and about which structural information is currently available. When isolated from mammalian sources it is visualized on SDSpolyacrylamide gel electrophoresis (SDS-PAGE) as a single polypeptide of M, 64,000. The receptor isolated from avian erythrocytes consists of two peptides, M, 40,000 and 50,000, present in variable proportions with the smaller peptide likely derived from the larger by proteolysis. Nonetheless both peptides appear to contain the intact P-adrenergic-ligand binding site. The amphibian erythrocyte P-adrenergic receptor has an apparent M, of 58.000. One of the striking features of the P-adrenergic receptor-adenylate cyclase system is that prolonged incubation of catecholamines with a cell leads to a diminution or blunting of the response to further challenge by the agonist. This process, termed “desensitization,” leads to reduced cAMP levels in the cell and consequently to a reduced cellular response to the hormone.

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Several different patterns of desensitization or refractoriness may be observed. In one type, generally termed “homologous desensitization,” exposure to a pagonist leads to diminished responsiveness only to subsequent stimulation by pagonists. The ability of other agonists to stimulate the adenylate cyclase through distinct receptors is preserved. In “heterologous desensitization” a more general blunting of responsiveness to other hormonal activators is also observed. In some situations even the actions of agents that bypass receptors in their stimulatory actions on the enzyme are reduced (e.g., NaF, forskolin). Heterologous desensitization appears to be modulated, at least in part, by CAMP, whereas homologous desensitization is not CAMPmediated. Considerable additional evidence, which has been recently reviewed elsewhere (3, 4), strongly suggests that different mechanisms are involved in homologous and heterologous desensitization. Homologous desensitization clearly seems to involve alterations in receptor activity whereas heterologous desensitization may be mediated by changes in the receptors or more distal components of the system. Homologous desensitization invariably involves sequestration or internalization of the receptors whereas heterologous desensitization involves receptor uncoupling from adenylate cyclase stimulation without receptor sequestration.

A. HETEROLOGOUS DESENSITIZATION Evidence has been presented that strongly implicates P-adrenergic receptor phosphorylation in the mechanism of heterologous desensitization. Stadel er al. (5) were the first to directly examine this question using turkey erythrocytes, a convenient model for the study of a heterologous form of desensitization. Using photoaffinity labeling techniques, it was found that the electrophoretic mobility on SDS-PAGE of (3-adrenergic receptors derived from desensitized turkey erythrocytes was different from that of receptors derived from control cells. The desensitized receptors appeared to migrate with decreased mobility. This alteration correlated with desensitization and could be mimicked by incubating the cells with cyclic nucleotide analogs. These findings were the first to suggest that phosphorylation of the p-adrenergic receptor might occur during desensitization since comparable alterations in electrophoretic mobility of other proteins after phosphorylation have been described (6, 7). In order to test this hypothesis, intact turkey erythrocytes were incubated with ’*Pi to label the intracellular ATP pool (8).Cells were then incubated with either buffer alone, agonist to promote desensitization or agonist plus antagonist to block the desensitization. The P-adrenergic receptors from the three groups were then partially purified by solubilization and affinity chromatography before characterization by SDS-PAGE. Figure 1 is an autoradiogram depicting such an experiment. Lanes 1 and 2 show [Iz5I]pABC (a p-adrenergic specific pho-

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2

3

4

5

FIG. 1. Autoradiogram of a SDS-PAGE of 32P-labeled P-adrenergic receptors partially purified from control and isoproterenol-desensitized turkey erythrocytes. Turkey erythrocytes were preincubated with [32P]orthophosphate for 20 h; 32P-labeled cells were then incubated for 4 h in buffer alone (lane 3), with 1 pi4 isoproterenol (lane 4),or with 1 pi4 isoproterenol plus 10 pi4 propranolol (lane 5 ) . P-Receptors were partially purified from membranes prepared from these cells by solubilization and affinity chromatography. Included in this panel for comparison are P-receptors labeled with [1251]pABCin membranes prepared from control (lane I ) and isoproterenol-desensitized (lane 2) erythrocytes from a separate experiment run on the same gel.

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toaffinity probe) labeled receptor from the membranes of control and desensitized cells. Lanes 3, 4 and 5 show 32P-labeledreceptor from control, desensitized, and agonist plus antagonist preincubated cells. This experiment demonstrates that the turkey erythrocyte P-adrenergic receptor is a phosphoprotein and that following desensitization the phosphate content of the receptor is increased two to three times compared to control. The ability of antagonist to block both the altered receptor mobility and increased 32P incorporation correlates well with the ability of the antagonist to prevent agonist-induced desensitization. This study demonstrates that the P-adrenergic receptor is specifically phosphorylated in turkey erythrocytes in response to prolonged challenge by agonist. Further evidence that desensitization of turkey erythrocyte adenylate cyclase is correlated with phosphorylation of the P-adrenergic receptor was provided by the work of Sibley et ul. (9). They found that under basal conditions the receptor contains 0.75 mol phosphate/mol while under maximally desensitized conditions the ratio increases to 2.34 mol/mol. They also demonstrated that the agonist dose-response curve as well as the time courses for desensitization and resensitization were identical for adenylate cyclase desensitization and receptor phosphorylation. The finding that incubation of turkey erythrocytes with cAMP and cAMP analogs only partially mimics isoproterenol in promoting adenylate cyclase desensitization and receptor phosphorylation suggests that only part of this process is mediated by the CAMP-dependent protein kinase. To directly assess the functionality of the P-adrenergic receptor from desensitized turkey erythrocytes, studies using receptor purified from both control and desensitized cells were performed (10). The purified receptors were implanted into phospholipid vesicles which were subsequently fused with Xenopus luevis erythrocytes. These cells contain N, and adenylate cyclase but little or no padrenergic receptor. As shown in Fig. 2, the fusion of reconstituted P-adrenergic receptor prepared from control cells establishes a sevenfold isoproterenol-induced stimulation of adenvlate cyclase activity in the previously unresponsive X. luevis erythrocytes. By contrast, receptor prepared from desensitized turkey erythrocytes establishes only a fourfold stimulation of adenylate cyclase. This represents a 45% decrease in the response and correlates very well with the extent of isoproterenol-induceddesensitization of the adenylate cyclase observed in the original crude membranes. These results thus demonstrate a direct relation between a stable modification of the P-adrenergic receptor due to hormoneinduced desensitization and a functional impairment of the receptor in a reconstituted system. Several studies have demonstrated that incubation of phorbol esters with duck and turkey erythrocytes leads to both desensitization of adenylate cyclase and phosphorylation of the P-adrenergic receptor (11, 12). That phorbol esters lead to desensitization of adenylate cyclase also had previously been shown in C6

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X&X

X&C

X&D

FIG.2. Fusion of X . luevis erythrocytes with affinity chromatography-purified and reconstituted P-adrenergic receptors from control and desensitized turkey erythrocytes. Receptor preparations were reconstituted into phospholipid vesicles and fused with X . laevis erythrocytes as described in (10). The same number of control and desensitized receptors were used in the reconstituion protocol and the same efficiency of insertion (20 to 25%) was observed in both cases. Abbreviations: B, basal: Iso, 50 )LM (-) isoproterenol; Is0 + Pro, 50 )LM (-)isoproterenol plus 50 pA4 (+)propranolol; PGE], 3 )LM prostaglandin El; and NaF, 10 mM NaF.

glioma (13) and mouse skin and epidermis (14). Since phorbol esters appear to act by stimulating the enzyme protein kinase C [reviewed in Ref. (IS)],these studies suggest that protein kinase C may be able to directly modulate cAMP levels via phosphorylation and desensitization of the adenylate cyclase-coupled 6-adrenergic receptor. In an attempt to more directly determine what protein kinases are involved in receptor phosphorylation, a cell-free system for desensitization of adenylate cyclase was developed (16, 17). Desensitization of adenylate cyclase in isolated turkey erythrocyte membranes was shown to require the presence of ATP, Mg2+, and factor(s) present in the soluble fraction of the cell. In the cell-free system, isoproterenol promoted a 40-60% desensitization while cAMP led to a 20-30% decrease in adenylate cyclase activity. When the soluble fraction of the cell was not included, no desensitization of adenylate cyclase was observed. However, when isolated turkey erythrocyte membranes were incubated with purified CAMP-dependent protein kinase an 20-30% desensitization was observed. This effect is completely abolished by the specific inhibitor of the kinase. In contrast, only about one-half of the 40-60% desensitization induced in a turkey erythrocyte lysate by isoproterenol is blocked by the inhibitor of the CAMP-dependent protein kinase. Similarly only about one-half of the phosphorylation of the P-adrenergic receptor induced in the lysate system by isoproterenol is blocked by the specific CAMP-dependent protein kinase inhibitor. In this cell-free lysate system phorbol esters also promoted P-adrenergic receptor desensitization and phosphorylation ( I 7). This desensitization could be

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mimicked by incubating isolated turkey erythrocyte membranes with partially purified preparations of protein kinase C plus phorbol esters. Calmodulin also promoted receptor phosphorylation and desensitization but to a much lesser extent than either isoproterenol or phorbol esters. The effect of calmodulin was entirely blocked by EGTA. These findings with the cell-free system suggest that (a) multiple protein kinase systems are capable of regulating P-adrenergic receptor function via phosphorylation reactions, and (b) CAMP may not be the sole mediator of isoproterenol-induced heterologous desensitization in these cells. We have directly studied the ability of several different protein kinases to phosphorylate the purified P-adrenergic receptor (18). In these studies, purified hamster lung P-adrenergic receptor was incubated with [ Y ~ ~ P I A TMg2+, P, and several different protein kinases. It was found that cGMP protein kinase. myosin light chain kinase, casein kinases I and 11, and rhodopsin kinase were unable to phosphorylate the receptor. Conversely, CAMP-dependent protein kinase was

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I INCUBATION TIME (HR)

2

FIG.3. Effect of isoproterenol on the time course of CAMP-dependent protein kinase-catalyzed padrenergic receptor phosphorylation. Purified hamster lung p-adrenergic receptor (0.14 @) was incubated with the catalytic subunit of CAMP-dependent protein kinase (0.34 @) in the presence or absence of 20 @ (-)isoproterenol at 25°C for the indicated period of time. Reactions were stopped by the addition of SDS sample buffer followed by electrophoresis on a 10%SDS-polyacrylamidegel. After drying and autoradiography, the gel was cut and counted to determine the stoichiometry.

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able to phosphorylate the receptor in a stoichiometric fashion. The time course of receptor phosphorylation by the CAMP-dependent protein kinase is shown in Fig. 3. It is seen that inclusion of the P-agonist, isoproterenol, promotes a 2- to 3-fold increase in the rate of receptor phosphorylation. This effect can be blocked by Pantagonists and suggests that the agonist induces a conformational change in the receptor which exposes the phosphorylation site(s). The stoichiometry of 0.5 mol 32P/mol receptor seen in Fig. 3 was found to be very dependent on the incubation conditions. When the receptor used was initially reconstituted in phospholipid vesicles a stoichiometry of 2 mol/mol could be attained. Highperformance liquid chromatography (HPLC) tryptic mapping of 32P-labeled receptor revealed two major phosphorylation sites both at serine residues. The phosphorylated receptor can be completely dephosphorylated by a high-molecular-weight phosphoprotein phosphatase. The rate of receptor dephosphorylation is also specifically enhanced 2- to 3-fold by isoproterenol again suggesting a conformational change in the receptor that exposes the phosphorylation sites. The functional significance of receptor phosphorylation was studied using ligand binding and reconstitution techniques. The P-adrenergic receptor, in addition to binding specific ligands, is able to interact with N, in a manner that promotes N, activation of adenylate cyclase. The interaction of purified receptor and N, in phospholipid vesicles is promoted by agonists and may be monitored by measuring the GTPase activity of N,. Figure 4 demonstrates that phosphorylated P-adrenergic receptor has a diminished ability to interact with N, with no apparent change in ligand binding. When the receptor is phosphorylated an 25% decrease in isoproterenol-promoted GTPase activity is seen relative to control receptor. The concentrations of isoproterenol which promoted 50% of the maximum GTPase activities were 136 and 125 nM for control and phosphorylated receptor, respectively. These studies provide a direct demonstration of regulation of the function of the isolated P-adrenergic receptor by CAMP-dependent protein kinase. Moreover, they suggest one possible general mechanism for heterologous regulation of adenylate cyclase-coupled receptors. Cyclic AMP generated in response to agonist stimulation of the enzyme activates the CAMP-dependent kinase. This kinase presumably phosphorylates multiple receptors, thus uncoupling them from productive interaction with N,. Although the structures of the various adenylate cyclase-coupled receptors are not known, it is reasonable to speculate that there are regions of strong homology since they all couple to the same N, molecules. Thus, all these receptors might serve as substrates for the CAMPdependent kinase. Moreover, the ability of an agonist to change the conformation of the receptor in such a way as to make it a better substrate for the kinase provides an additional mechanism for fine-tuning the regulation. Thus, in addition to phosphorylation and desensitization of a variety of receptors there might be even more profound alterations of the particular type of receptor that is

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0 CONTROL B A R

0 PHOSPHORYLATED B A R

n 5l-

/.I

n ”

9

8

7

-Log,,

s sop rote re no^]

6

5

4

M

FIG.4. Isoproterenol-promoted GTPase activity in phospholipid vesicles containing N, and phosphorylated or control receptor. Phospholipid vesicles containing P-adrenergic receptor, N,, and the catalytic subunit of CAMP-dependent protein kinase were incubated with 30 mM sodium phosphate, 10 mM tris-HC1, pH 7.2, 100 mM NaCI, 5 mM MgC12, 5 mM p-nitrophenyl phosphate, 50 pM AppNHp, and 250 k g of soybean phosphatidylcholine. Phosphorylation incubations additionally contained 50 pM ATP whereas control samples contained no ATP. After incubating for 2 hr at 25°C the samples were assayed for GTPase activity as a function of the isoproterenol concentration.

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occupied by its specific agonist. Finally, it is worth stressing, as previously noted, that kinases other than the CAMP-dependent kinase may well participate in these phosphorylation events.

B . HOMOLOGOUSDESENSITIZATION The possible involvement of P-adrenergic receptor phosphorylation in homologous desensitization has come under investigation. As previously noted, this form of desensitization is more specific for one class of agonist, is not CAMPmediated, and involves receptor sequestration within a poorly defined internalized compartment of membrane. Some evidence suggests that once “sequestered,’’ the receptors may be functionally normal [Ref. (19), as assessed by reconstitution approaches]. However, other evidence suggests that, especially at very early time points in the desensitization process, a functional alteration in the receptors may occur (20). Sibley et al. (21) have documented that homologous desensitization of the padrenergic receptor in the frog erythrocyte is associated with stoichiometric

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phosphorylation of the receptor. Other agents that raise cAMP levels in the cells without desensitizing the P-adrenergic receptor, such as PGE, , do not lead to padrenergic receptor phosphorylation under these conditions. This is consistent with the apparent lack of dependence of homologous desensitization on cAMP generation. Whether the receptor phosphorylation occurring during homologous desensitization is involved in triggering receptor sequestration, in directly uncoupling the receptor, or both, remains to be determined. There is also no evidence bearing on the nature of the kinases involved, other than the presumption that it is not the CAMP-dependent protein kinase.

111. Rhodopsin The rod outer segment of the retina converts a light stimulus into a change in the intracellular concentration of cGMP [reviewed in Ref. (22)l.This process is mediated by the GTP-dependent interaction of rhodopsin, a receptor for light, and transducin, a guanine nucleotide regulatory protein which is structurally similar to N, and Ni (23). Activated transducin is able to directly stimulate the enzyme cGMP phosphodiesterase leading to a decrease in intracellular cGMP concentrations. A possible role of phosphorylation in the regulation of this system was noted by three independent groups in th.: early 1970s when they discovered a light-dependent phosphorylation of rhodopsin (24-26). The kinase involved in the phosphorylation appears to be !iighly specific for freshly bleached rhodopsin as a substrate (27-29). Under optimal conditions as many as 9 or 10 phosphate groups are incorporated into serine and threonine residues in the carboxyl-terminal region of rhodopsin (30, 31). Several groups have suggested that phosphorylation of rhodopsin is a mechanism to turn off phosphodiesterase activity (32-36). This was based on their findings that ATP suppresses light-stimulated activation of phosphodiesterase and GTP binding to rod membranes. The inability of phosphorylated rhodopsin to activate phosphodiesterase (via a reduced interaction with transducin) was directly demonstrated by Shichi er al. (37). In these studies, rhodopsin species containing either 0, 1, or 2 mol phosphate/mol rhodopsin were isolated by electrofocusing. The ability of these three rhodopsin species to activate transducin in phospholipid vesicles was then studied. Rhodopsin with 1 mol phosphate/mol was only 4% as active as control (no phosphate) whereas rhodopsin with 2 mol/mol was completely inactive. While the phosphorylation of rhodopsin inhibits its interaction with transducin (37), it also appears to enhance its interaction with a 48,000-dalton protein (38).This major soluble protein in rod cells appears to compete with transducin for binding to phosphorylated rhodopsin and thus may also be involved in regulating phosphodiesterase activity (38). These studies support the notion that light-activated phosphorylation of rhodop-

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sin directly leads to decreased activation of cGMP phosphodiesterase. This may represent a mechanism for light adaptation analogous to the desensitization of adenylate cyclase-coupled receptors discussed in Section 11.

IV. The Nicotinic Acetylcholine Receptor The nicotinic acetylcholine receptor (also see Chapter 11) is an integral membrane protein that functions as a ligand-gated ion channel at the vertebrate neuromuscular junction (39). The acetylcholine receptor has been most extensively studied in the Torpedo electric organ and exists as an oligomer of four polypeptide chains with masses of 40 (a),50 (p), 60 (y), and 65 ( 6 ) kilodaltons with a stoichiometry of a2 py6 (40).Phosphorylation of the acetylcholine receptor has been shown to occur both in vivo and in vitro with as rnany as 9 phosphoserines being identified, distributed 1 , 1, 2, and 5 among the a,p, y, and 6 subunits, respectively (41). Postsynaptic membranes rich in the acetylcholine receptor appear to contain both endogenous protein kinase (42-44) and protein phosphatase activities (45). The endogenous protein kinases include the CAMP-dependent protein kinase which phosphorylates the y and 6 subunits of the receptor on serine residues (46, 47). This phosphorylation can be completely blocked by the specific inhibitor of the CAMP-dependent protein kinase. Phosphorylation studies of purified acetylcholine receptor with the catalytic subunit of CAMPdependent protein kinase also demonstrate phosphorylation of the y and S subunits with stoichiometries of 1.O and 0.89 mol 32P/mol receptor, respectively (46). Endogenous protein kinase C can also phosphorylate the y and 6 subunits of the receptor on serine residues (48).In addition, an endogenous tyrosine protein kinase which phosphorylates the p, y, and S subunits of the receptor has been reported (49).This kinase phosphorylates a single tyrosine residue on each of the subunits with stoichiometries of 0.5 mol/mol attained in postsynaptic membranes. It has also been reported that purified pp60""' of Rous sarcoma virus and A431 cell membranes, which are rich in EGF receptor, both specifically phosphorylate purified acetylcholine receptor on the p, y, and 6 subunits (49).While the acetylcholine receptor can be extensively phosphorylated both in vitro and in vivo, the effects of phosphorylation on the function of the receptor are unknown.

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V. The Receptors for EGF and Insulin The EGF receptor mediates a wide array of biological effects including increased glycolytic activity, stimulation of amino acid and sugar transport, and increased rates of protein, RNA, and DNA synthesis (50) (also see Vol. XVII,

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Chap. 7). One of the earliest events after EGF binding is activation of a tyrosinespecific protein kinase which appears to be intrinsic to the EGF receptor (51-54) (see Vol. XVII, Chapter 6). Autophosphorylation of the EGF receptor occurs at one tyrosine residue in vivo (52, 55), whereas in vitro as many as four tyrosines are phosphorylated (55). In addition, extensive phosphorylation of the receptor at both serine and threonine residues occurs in vivo (52). Several lines of evidence suggest that protein kinase C is involved in the phosphorylation of the EGF receptor. It has been demonstrated that tetradecanoyl phorbol acetate (TPA) and other active tumor-promoting phorbol esters can modulate the binding of EGF to its receptor (56-65). Depending upon the cell type, phorbol esters appear to cause either a reduction in the number of EGF receptor sites (56-59) or a decrease in the affinity of the receptors for EGF (60-65). Phorbol ester treatment also stimulates phosphorylation of the EGF receptor at serine and threonine residues and appears to inhibit the tyrosine kinase activity of the receptor leading to reduced phosphotyrosine levels (64-67). Similar results are observed when diacylglycerol is used, again implicating protein kinase C (68). Protein kinase C can directly phosphorylate purified EGF receptor at three threonine residues and again results in decreased tyrosine kinase activity (66). Since EGF can enhance Ca2 influx and diacylglycerol formation (69), both endogenous activators of protein kinase C, this may represent a mechanism for feedback inhibition of the receptor tyrosine kinase. Phosphorylation of the EGF receptor by CAMP-dependent protein kinase has also been demonstrated in vitro (70, 71). However, a physiological role of EGF receptor phosphorylation by this kinase remains to be established. Insulin binding to its receptor also triggers a wide variety of biological effects (72). One of the earliest responses is phosphorylation of the P-subunit of the insulin receptor. Like the EGF receptor, the insulin receptor is phosphorylated on tyrosine by a receptor-associated tyrosine kinase (73-77) (see Vol. XVII, Chapter 7). The receptor is also extensively phosphorylated on serine and threonine residues. Several studies suggest that protein kinase C is involved in insulin receptor phosphorylation. Phorbol esters appear to decrease the affinity of insulin for its receptor in several cell lines (78, 79) although in some cells no effect on insulin binding was observed (57, 60. 80). Phorbol esters also promote phosphorylation of the insulin receptor (80, 81) and appear to inhibit insulin-induced receptor phosphorylation and insulin action (80). These studies suggest that phosphorylation of the insulin receptor regulates insulin action. +

VI. Other Membrane Receptors Mast cells and basophils have cell-surface receptors that bind IgE. The IgE receptor is composed of an a component of M, 45,000 and a P component of

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33,000. Phosphorylation studies with a rat basophilic leukemia cell line have demonstrated a non-IgE-mediated phosphorylation of the p component of the receptor (82). In contrast to these findings Hempstead et al. (83, 84) have shown phosphorylation of both the a and p components of the receptor in rat mast cells. Additionally, when these cells were stimulated with antigen a rapid increase (55% in 15 s) in the phosphorylation of the a component was observed with no apparent change in p component phosphorylation. These studies suggest that increased phosphorylation of the a component of the receptor may be part of the antigen-stimulated process leading to mediator secretion. The transferrin receptor is a 180-kDa phosphorylated glycoprotein consisting of two identical subunits. This receptor mediates endocytosis of transferrin resulting in cellular uptake of Fe2+. The signal for receptor-mediated endocytosis, other than ligand binding, is unknown. Two groups have shown, however, that phorbol esters induce a 10- to 20-fold increase in phosphorylation of the transferrin receptor as well as an 50% decrease in the number of cell-surface transferrin receptors (85, 86). While it is tempting to speculate that phosphorylation may be the signal for receptor-mediated endocytosis this does not appear to be the case in HL60 cells (85).In these cells, transferrin does not induce phosphorylation of the receptor nor inhibit phorbol ester-induced phosphorylation of the transfenin receptor. However, phorbol ester-induced phosphorylation of the transferrin receptor may well be acting as a signal for receptor internalization. M,

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REFERENCES 1. 2. 3. 4. 5.

6. 7. 8. 9. 10. 11. 12. 13.

14. 15.

Lefkowitz, R. J., Stadel, J. M., and Caron, M. G. (1983). Annu. Rev. Biochem. 52, 159. Stiles, G. L., Caron, M. G., and Lefkowitz, R. J . (1984). Physiol. Rev. 64, 661. Harden, T. K. (1983). Pharmacol. Rev. 35, 5. Sibley, D. R., and Lefkowitz, R. J . (1985). Nature (London) 317, 124. Stadel, J . M., Nambi, P., Lavin, T. N., Heald, S. L., Caron, M. G., and Lefkowitz, R. J. (1982). JBC 257, 9242. Zallor, M. J . , Kerlavage, A. R.,and Taylor, S. S. (1979). JBC 254, 2408. Shih, T. Y., Weeks, M. O., Young, H. A., and Scolnick, E. M. (1979). Virology 96, 64. Stadel, J . M., Nambi, P., Shorr, R. G. L., Sawyer, D. F., Caron, M. G., and Lefkowitz, R. J. (1983). PNAS 80, 3173. Sibley, D. R., Peters, J. R., Nambi, P., Caron, M. G., and Letkowitz, R. J. (1984). JBC 259, 9742. Strulovici, B., Cerione, R. A., Kilpatrick, B. F., Caron, M. G., and Lefkowitz, R. J. (1984). Science 225, 837. Sibley, D. R., Nambi, P., Peters, J . R., and Lefkowitz, R. J . (1984). BBRC 121, 973. Kelleher, D. I . , Pessin, J . E., Ruoho, A. E., and Johnson, G. L. (1984). PNAS 81, 4316. Mallorga, P., Tallman, J. F., Hennebury, R. C., Hirata, F., Strittmatter, W. T., and Axelrod, J. (1980). PNAS 77, 1341. Belman, S., and Carte, S. J. (1980). Nature (London) 284, 171. Nishizuka, Y. (1984). Narure (London) 308, 693.

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16. Nambi, P., Sibley, D. R., Stadel, J . M., Michel, T., Peters, J. R., and Lefkowitz, R. J. (1984). JBC 259, 4629. 17. Nambi, P., Peters, J. R., Sibley, D. R., and Lefkowitz, R. J. (1985). JBC 260, 2165. 18. Benovic, J. L., Pike, L. J., Cerione, R. A., Staniszewski, C., Yoshimasa, T., Codina, J., Bimbaumer, L., Caron, M. G., and Lefkowitz, R. J. (1985). JBC 260, 7094. 19. Strulovici, B., Stadel, J. M.,and Lefkowitz, R. J. (1983). JBC 258, 6410. 20. Waldo, G. L., Northup, J. K., Perkins, J . P., and Harden, T. K. (1983). JBC 258, 13900. 21. Sibley, D. R., Strasser, R. H., Caron, M. G., and Lefkowitz, R. J. (1985). JBC 260, 3883. 22. Knowles, A. (1984). Prog. Retinal Res. 3, 157. 23. Manning, D. R., and Gilman, A. G. (1983). JBC 258, 7059. 24. Kuhn, H., and Dreyer, W. J. (1972). FEES Lett. 20, 1. 25. Bownds, D., Dawes, J., Miller, J., and Stahlman, M. (1972). Nature (London), New B i d . 237, 125. 26. Frank, R. N., Cavanagh, H.D., and Renyon, K. R. (1973). JBC 248, 596. 27. Shichi, H., Somers, R. L., and O’Brien, P. J. (1974). BBRC 61, 217. 28. Kuhn, H., and McDowell, J. H. (1977). Biophys. Struct. Mech. 3, 199. 29. Shichi, H., and Somers, R. L. (1978). JBC 253, 7040. 30. Wilden, U., and Kuhn, H. (1982). Biochemistry 21, 3014. 31. Hermolin, J., Karell, M. A., Hamm, H. E., and Bownds, M. D. (1982). J. Gen. Physiof. 79, 633. 32. Kuhn, H., Cook, J. H., and Dreyer, W. J. (1973). Biochemistry 12, 2495. 33. Liebman, P. A . , and Pugh, E. N. (1980). Narure (London) 287, 734. 34. Liebman, P. A., and Pugh, E. N. (1982). Vision Res. 22, 1475. 35. Sitaramayya, A., and Liebman, P. A. (1983). JBC 258, 1205. 36. Sitaramayya, A., and Liebman, P. A. (1983). JBC 258, 12106. 37. Shichi, H., Yamamoto, K., and Somers, R. L. (1984). Vision Res. 24, 1523. 38. Kuhn, H., Hall, S. W., and Wilden, U. (1984). FEBSLett. 176, 473. 39. Changeux, J.-P. (1981). Harvey Lect. 75, 85. 40. Reynolds, J. A., and Karlin, A. (1978). Biochemistry 17, 2035. 41. Vandlen, R. L., Wu, W. C.-S., Eisenach, J . C., and Raftery, M. A. (1979). Biochemistry 18, 1845. 42. Gordon, A. S., Davis, C. G., Milfay, D., and Diamond, I. (1977). Nature (London) 267, 539. 43. Teichberg, V. I., Sobel, A,, and Changeux, J.-P. (1977). Nature (London) 267, 540. 44. Saitoh, T., and Changeux, J.-P. (1981). PNAS 78, 4430. 45. Gordon, A. S . , Milfay, D., Davis, C. G., and Diamond, I. (1979). BBRC 87, 876. 46. Huganir, R. L., and Greengard, P. (1983). PNAS 80, 1130. 47. Zavoico, G. B., Comerci, C., Subers, E., Egan, J. J., Huang, C.-K., Feinstein, M. B., and Smilowitz, H. (1984). BBA 770, 225. 48. Huganir, R. L., Albert, K. A., and Greengard, P. (1983). Soc. Neurosci. Abstr. 9, 578. 49. Huganir, R. L., Miles, K., and Greengard, P. (1984). PNAS 81, 6968. 50. Carpenter, G., and Cohen, S. (1979). Annu. Rev. Biochem. 48, 193. 51. Cohen, S., Carpenter, G., and King, L. (1980). JBC 255, 4834. 52. Hunter, T., and Cooper, J. A. (1981). Cell (Cambridge, Mass.) 24, 741. 53. Cohen, S . , Ushiro, H., Stoscheck, C., and Chinkers, M. (1982). JBC 257, 1523. 54. Kasuga, M., Zick, Y.,Blithe, D. L.,Krettaz, M., and Kahn, C. R. (1982). Nature (London) 298, 667. 5 5 . Weber, W., Bertics, P. J., and Gill, G. N. (1984). JBC 259, 14631. 56. Lee, L.-S., and Weinstein, I. B. (1978). Science 202, 313. 57. Lee, L.-S., and Weinstein, I. B. (1979). PNAS 76, 5168. 58. Murray, A. W., and Froscio, M. (1980). Carcinogenesis (London) 31, 681.

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59. Salomon, D. S. (1981). JBC 256, 7958. 60. Shoyab, M., De Larco, J. E., and Todaro, G.J. (1979). Nature (London) 279, 387. 61. Brown, K. D., Dicker, P., and Rozengurt, E. (1979). BBRC 86, 1037. 62. Magun, B. E., Matrisian. L. M., and Bowden, G.T. (1980). JBC 255, 6373. 63. King, A. C., and Cuatrecasas, P. (1982). JBC 257, 3053. 64. Friedman, B. A., Frackelton, A. R., Jr., Ross, A. H., Connors, J. M., Fujiki, H., Sugimura, T., and Rosner, M. R. (1984). PNAS 81, 3034. 65. Davis, R. J . , and Czech, M. P. (1984). JBC 259, 8545. 66. Cochet, C., Gill, G.N., Meisenhelder, J., Cooper, J. A., and Hunter, T. (1984). JBC 259, 2553. 67. Iwashita, S . , and Fox, C. F. (1984). JBC 259, 2559. 68. McCaffrey, P. G.,Friedman, B. A., and Rosner, M. R. (1984). JBC 259, 12502. 69. Sawyer, S . T., and Cohen, S. (1981). Biochemistry 20, 6280. 70. Rackoff, W. R., Rubin, R. A , , and Earp, H. S. (1984). Mol. Cell. Endocrinol. 34, 113. 71. Dastidar, P. G.,and Fox, C. F. (1984). JBC 259, 3864. 72. Czech, M. P. (1977). Annu. Rev. Biochem. 47, 359. 73. Kasuga, M., Karlsson, F. A., and Kahn. C. R. (1982). Science 215, 185. 74. Kasuga, M., Zick, Y., Blithe, D. R., Karlsson, F. A., Haring, H. U., and Kahn, C. R. (1982). JBC 257, 9891. 75. Zick, Y . , Kasuga, M., Kahn, C. R., and Roth, J. (1983). JBC 258, 75. 76. Zick, Y., Whittaker, J., and Roth, J. (1983). JBC 258, 3431. 77. Zick, Y., (hunberger, G.,Podskalny, J . M., Moncada, V., Taylor, S . I., Gorden, P., and Roth, J. (1983). BBRC 116, 1129. 78. Grunberger, G.,and Gordon, P. (1982). Am. J. Physiol. 243, E319. 79. Thomopoulos, P., Testa, U., Gourdin, M.-F., Hervy, C., Titeux, M., and Vainchenker, W. (1982). EJB 29, 389. 80. Takayama, S . , White, M. F., Lauris, V., and Kahn, C. R. (1984). PNAS 81, 7797. 81. Jacobs, S . , Sahyoun, N. E., Saltiel, A. R., and Cuatrecasas, P. (1983). PNAS 80, 6211. 82. Fewtrell, C., Goetze, A., and Metzger, H. (1982). Biochemistry 21, 2004. 83. Hempstead, B. L., Kulczycki, A,, Jr., and Parker, C. W. (1981). BBRC 98, 815. 84. Hempstead, B . L., Parker, C. W., and Kulczycki, A., Jr. (1983). PNAS 80, 3050. 85. May, W. S . , Jacobs, S . , and Cuatrecasas, P. (1984). PNAS 81, 2016. 86. Klausner, R. D., Harford, J., and van Renswoude, J. (1984). PNAS 81, 3005.

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Regulation of Ionic Channels SANDRA ROSSIE WILLIAM A. CATTERALL Department of Pharmacology University of Washington Seattle, Washington 981 95

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Calcium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Electrophysiological Evidence for Modulation of Calcium Channels by Phosphorylation . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . B. Biochemical Studies of Phosphorylation ........... 111. Potassium Channels .................... .................. A. Electrophysiological Evidence for Modulation of K Channels by Protein Phosphorylation . . . . . . . B. Functional Significa C. Biochemical Analysis of Potassium Channel Phosphorylation . . . . . . . . . IV . Acetylcholine Receptor A. Biochemical Studies of Phosphorylation .......................... B. Functional Sig ion . . . . . . . . . . . . ....... V. Sodium Channels ............... ...... A. Biochemical S B. Functional Significance of Phosphorylation ........................ VI . Conclusions . . . . . . . . . . References ..........................................

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Introduction

Most animal cells, including neurons, maintain large ionic gradients across their surface membranes such that the intracellular fluid contains a high con335 THE ENZYMES, Vol. XVlll Copyright 0 1987 by Academic Press, Inc. All rights of reproduction in any form reserved.

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centration of potassium ions and low concentrations of sodium ions and calcium ions relative to the extracellular fluid. These large ion gradients are maintained by the action of energy-dependent ion pumps specific for Na and K , or for Ca2+. In addition, nearly all cells maintain an internally negative membrane potential of the order of -60 millivolts because their surface membranes are specifically permeable to K , allowing K to leak out of cells faster than Na and Ca2+ can leak in. These ion gradients and the resting membrane potential provide the electrochemical driving force for ion movements mediated by ionic channels involved in electrical signaling. Ionic channels mediating electrical signaling are intrinsic membrane proteins that form ion-selective pores through which ions can move down their electrochemical gradients into or out of cells. The ionic channels can be divided into two classes, voltage-regulated and ligand-regulated. These primary regulatory or “gating” processes occur on the millisecond time scale and allow voltagesensitive ion channels to respond rapidly to changes in membrane voltage and ligand-gated ion channels to respond rapidly to synaptically released neurotransmitters. Voltage-sensitive ion channels are regulated by two experimentally separable processes: activation, which controls the voltage and time dependence of opening of the ion channel after depolarization; and inactivation, which controls the voltage- and time-dependence of channel closure during maintained depolarization. Ligand-gated ion channels are regulated by two formally analogous processes: activation, which leads to channel opening upon binding of a neurotransmitter or related ligand; and desensitization, which leads to channel closure during exposure to agonists. The responsiveness of ion channels to membrane potential and neurotransmitters is also regulated on the time scale of seconds to minutes. Protein phosphorylation is now implicated as an important component of this second-order regulation of ion channel function. As discussed elsewhere in this volume, many hormones and neurotransmitters act by altering intracellular levels of cyclic AMP, cyclic GMP and calcium. Cyclic AMP and cyclic GMP are thought to produce cellular responses mainly, if not exclusively, through activation of specific cyclic nucleotide-dependent protein kinases ( 1 , 2). Many of the effects of calcium are mediated by stimulation of calcium-dependent protein kinases ( 3 ) and calcium-dependent phosphoprotein phosphatases ( 4 ) . Several examples of ion channel modulation by phosphorylation events initiated by neurotransmitter action are now documented. In this chapter, electrophysiological and biochemical evidence for regulation of ion channels by phosphorylation are reviewed. Four major topics are considered: voltage-sensitive calcium channels, potassium channels, sodium channels, and the nicotinic acetylcholine receptor. Electrophysiological observations have documented modulation of both calcium and potassium channels by cyclic AMPdependent phosphorylation. Little is known about the biochemical identity of these channels, however, and it is not known whether regulation is achieved by +

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direct phosphorylation of the channel itself or of some associated regulatory protein. In contrast, the acetylcholine receptor and the voltage-sensitive sodium channel are biochemically identified channels known to be phosphoproteins in vivo. Both of these channels have been demonstrated to be substrates for several distinct kinases in vitro. The role of phosphorylation in modulating the acetylcholine receptor has recently been directly demonstrated.

II. Calcium Channels Voltage-sensitive calcium channels mediate the influx of extracellular calcium into excitable cells during membrane depolarization and provide the link between electrical signals and other physiological events ( 5 ) . Elevation of intracellular calcium initiates such specialized responses as secretion in neuronal and endocrine tissue and contraction of muscle (6). The free intracellular calcium concentration influences many cellular processes, and the level of cytoplasmic calcium is regulated in a complex manner. Not surprisingly, voltage-sensitive calcium channels, which help control concentration of this important intracellular messenger, are themselves subject to modulation. Three calcium channel subtypes have been identified in different tissues (129133). A channel responsible for long-lasting calcium currents l(L-type, 134) has been described in smooth, cardiac, and skeletal muscle and in many types of neurons. This channel is distinguished by its sensitivity to blockade by organic calcium channel antagonists such as the dihydropyridines, verapamil, and diltiazem. Modulation of L-type channels by a wide variety of hormones and neurotransmitters has been demonstrated in both cardiac muscle and neuronal tissue (8). The sensitivity of channels to modulation by different hormones and neurotransmitters varies with tissue source. The mechanism underlying modulation has not been elucidated in most cases that have been described. Only those cases in which regulation of calcium channel activity is clearly linked to protein phosphorylation are discussed in this section. A.

ELECTROPHYSIOLOGICAL EVIDENCE FOR MODULATION OF CALCIUM CHANNELS BY PHOSPHORYLATION

P-Adrenergic modulation of the slow inward calcium current in cardiac muscle is the first described and most thoroughly studied example of calcium current regulation. Voltage-clamp experiments performed on mammalian (9) and frog (10) cardiac tissue showed that P-adrenergic stimulation enhances calcium current. This effect contributes significantly to the increase in cardiac contractility, beat rate, and amplitude of the cardiac action potential caused by P-adrenergic

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agonists (9). The potentiation of the slow inward current appears to be mediated by cyclic AMP and consequent activation of cyclic AMP-dependent protein kinase. Incubation of tissue with cyclic AMP derivatives (IZ-I.?), or phosphodiesterase inhibitors to prevent cyclic nucleotide degradation (12, 13), mimic the action of 6-adrenergic agonists on cardiac tissue. Cardiac tissue can be dissociated into single, electrically competent myocytes which can be used for recording of whole-cell and single-channel currents, as well as for microinjection studies. Direct injection of cyclic AMP (14) or the purified catalytic subunit of cyclic AMP-dependent protein kinase (15) into isolated cardiac myocytes influence the action potential and the calcium current in a manner identical to that of P-adrenergic stimulation. The response of guinea

100ms

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+lOmV

-55mv r

B

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FIG. I . The response of isolated guinea pig cardiac myocytes to intracellular injection of purified catalytic subunit of cyclic AMP-dependent protein kinase. (A) Evoked action potentials 30, 60, and 120 s after injection of catalytic subunit. (B) The response of the slow inward current in a voltageclamped cell to injection of catalytic subunit. The top tracing shows the voltage step, the bottom tracing shows current responses to the voltage step before (upper trace) and after (lower trace) injection. Data are reproduced from Ref. (26).

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A

-80

200 ms

FIG. 2. Effects of protein kinase and adrenaline are not additive. (A) Action potentials recorded from an isolated guinea pig cardiac myocyte after multiple injections of catalytic subunit of cyclic AMP-dependent protein kinase. (B) After maximal response to catalytic subunit has been induced, adrenaline caused no further increase in the amplitude of the action potential recorded from this cell. Data are reproduced from Ref. (16).

pig cardiac myocytes to injection of the catalytic subunit of cyclic AMP-dependent protein kinase is depicted in Fig. 1 (16). Prolongation of the action potential (Fig. 1A) and the enhancement of the slow inward current (Fig. 1B) are illustrated. Figure 2 shows that maximal stimulation of a cell by injection of the catalytic subunit prevents additional response to application of adrenaline. This is consistent with the hypothesis that P-adrenergic agonists act through cyclic AMP-dependent phosphorylation in causing this response. Analysis of P-adrenergic enhancement of calcium current under voltage-clamp conditions showed that the increase was not due to a change in the sensitivity of channels to voltage, but rather to an increase in the maximal calcium conductance (17-19). If total current is viewed as a summation of current through individual channels, then an increase in total current could be caused by either an increase in unit channel conductance, in the number of functional channels, or in

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the probability of channel opening at any voltage. Records of single calcium channel activity have been used to further examine these possibilities (20-23). Calcium channels monitored by the patch-clamp method of Hamill ef al. (24) are seen to respond to depolarizing voltage steps with bursts of activity consisting of rapid opening and closing, with prolonged periods of residence in a closed state between bursts (20). Single channel recording from cultured rat cardiac myocytes (20-22) and from isolated frog ventricular myocytes (23) showed that the unit channel conductance did not change in the presence of P-adrenergic agonists. Small increases in the mean channel open time and in the frequency of channel opening were observed in rat myocytes (20-22). In most observations of channel activity in cultured rat myocytes, the number of functional channels seen within a recording patch did not increase on treatment with P-adrenergic agonists (22) or 8-bromo cyclic AMP (21, 22). Thus, in this preparation an increase in the probability of channel opening is the major underlying cause of the P-adrenergicinduced increase in calcium current. In contrast, an increase in the number of functional channels may contribute to, and perhaps dominate, the enhanced frequency of channel openings seen in other preparations (17, 18, 23). Isolated frog ventricular myocytes have a much larger response to P-adrenergic stimulation than rat myocytes. In these cells, an increase in the number of functional channels was clearly observed in recordings from cell-attached patches treated with isoproterenol (23). Analysis of fluctuation in voltage-clamped whole cell calcium currents also led to the conclusion that P-adrenergic stimulation caused an increase in the number of functional channels in frog ventricular myocytes (23). Thus, the increase in inward calcium current during the cardiac action potential induced by P-adrenergic agents probably results from a combination of two effects: an increase in the probability of activation of functional calcium channels that are in an “activable” pool, and an increase in the number of functional channels by transfer from an “inactivable” pool to the activable pool. The results raise the possibility that calcium channels may require cyclic AMPdependent phosphorylation to be activable. Other observations also indicate that cyclic AMP-dependent phosphorylation is important for the maintenance of active calcium channels. In cells whose cytoplasm was removed by internal perfusion (25-28) and in isolated membrane patches (27, 29), calcium current rapidly degenerated. These findings suggest that metabolic processes are required to sustain functional calcium channels. In voltage-clamp studies on snail neurons (26) and rat dorsal root ganglion cells (25), calcium conductance was stabilized by intracellular perfusion with cyclic AMP, ATP and Mg2+, or F-, implying that cyclic AMP-dependent phosphorylation is required to maintain calcium channel activity. In snail neurons, intracellular perfusion with purified catalytic subunit of cyclic AMP-dependent protein kinase in combination with ATP and Mg2+ stabilized calcium conductance (30). In cardiac myocytes, injection of purified regulatory subunit of cyclic

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34 I

AMP-dependent protein kinase reduced the plateau phase of depolarizationevoked action potentials (15). These results indicate that cyclic AMP-dependent phosphorylation is involved in the maintenance of functional calcium channels in the basal state. Several observations suggest that phosphorylation by protein kinase C also modulates calcium channels. In Aplysia bag cell neurons, application of the phorbol ester 12-0-tetradecanoyl-phorbol-13-acetate, or intracellular injection of protein kinase C increased inward calcium current during action potentials (31). In chick dorsal root ganglion cells, treatment with activators of protein kinase C reduced voltage-sensitive calcium current (135). This response is similar to that seen with norepinephrine and y aminobutyric acid (136, 137) in this preparation. After pretreatment with a protein kinase C activator, norepinephrine produced no further reduction in calcium current (135), suggesting that protein kinase C dependent phosphorylation events may mediate transmitter-induced reduction of calcium current in these cells.

B. BIOCHEMICAL STUDIES OF PHOSPHORYLATION The results discussed above indicate that cyclic AMP-dependent phosphorylation is involved in both the maintenance and recruitment of functional calcium channels. The calcium channel itself or an associated regulatory protein may be the substrate for phosphorylation. Several approaches have been taken to identify proteins whose phosphorylation may regulate the calcium channel. Rinaldi et al. (32, 33) identified a proteolipid of 23 kDa in cardiac sarcolemma, calciductin, that is phosphorylated in a cyclic AMP-dependent manner. Phosphorylation of calciductin was correlated with 45Ca2+ influx into vesicles (32, 3 3 , leading to the hypothesis that calciductin phosphorylation mediates regulation of calcium channels. However, the finding in this study of a large accumulated calcium concentration suggested that an active uptake mechanism for calcium, rather than passive diffusion down an electrochemical gradient, accounted for these results (34). In bovine heart sarcolemmal vesicles, Flockerzi et al. (35) also observed cyclic AMP-dependent phosphorylation of a protein of 23 kDa, but found no effect of phosphorylation on voltage-dependent calcium uptake. In addition, calciductin has not been unequivocally distinguished from phospholamban (36, 37). Thus, the relevance of calciductin phosphorylation to regulation of calcium channels is uncertain. Home et al. (38) proposed that polypeptides of 42 and 45 kDa are components of the cardiac calcium channel based on covalent labeling with a dihydropyridine isothiocyanate. Nitrendipine, a calcium channel antagonist, and isoproterenol increased phosphorylation of a protein of similar size in cardiac membranes. However, phosphorylation was not dependent on cyclic AMP and the effect of isoproterenol was poorly blocked by P-adrenergic antagonists, suggesting that

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this phosphoprotein does not mediate the effects of P-adrenergic agonists on calcium channels. Curtis and Catterall purified a protein from skeletal muscle transverse-tubule membranes that binds dihydropyridine calcium channel antagonists with high affinity and consists of three subunits: a (162 kDa), P (50 kDa) and y (33 kDa) (39). This purified protein complex mediates 45Ca2 influx when reconstituted into phospholipid vesicles (138) and exhibits electrophysiological properties similar to those of calcium channels when reconstituted into phospholipid bilayers and studied in single channel mode (139). Thus, this complex likely represents the skeletal muscle calcium channel. Recent results have shown that activation of skeletal muscle calcium channels is also enhanced by cyclic AMP (140, 141). Two observations suggest that the calcium channel itself is directly regulated by phosphorylation. Both the CY and P subunits of the purified skeletal muscle calcium channel were shown to be good substrates for cyclic AMP-dependent protein kinase (40). The P subunit could also be phosphorylated in intact membranes (40). In electrophysiological studies of purified, reconstituted skeletal muscle channel, Flockerzi et af. (139) found that incubation of reconstituted channel with cyclic AMP-dependent protein kinase and ATP-y-S resulted in an increased probability of channel opening. These results support the conclusion that phosphorylation of CY or P subunits of the calcium channel itself is the mechanism of channel regulation by cyclic AMP-dependent protein phosphorylation. +

111.

Potassium Channels

The resting membrane potential of most excitable cells is near the equilibrium potential for potassium. Potassium currents act to set and stabilize the membrane potential of excitable cells. Thus, they decrease cellular excitability, limit the degree and duration of depolarization, control repetitive bursts of action potentials, and return the cell to its resting membrane potential after excitation (41). There is a wide variety of potassium channels differing in unit channel conductance, gating kinetics, sensitivity to voltage, and activation by calcium [for review, see Ref. (42)].By regulating the activity of potassium channels, neurotransmitters and hormones may predispose the level of excitability and electrical behavior of neurons, endocrine, or muscle cells. Modulation of several distinct types of potassium channels has been described [see Ref. (43)],although the mechanism of regulation is poorly understood in most cases. In several instances, potassium channel regulation appears to be mediated by cyclic AMPdependent phosphorylation (44-50). Potassium channel regulation by calcium and calmodulin-dependent (51) and protein kinase C-dependent (142, 143) phosphorylation have also been described.

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FOR MODULATION OF K A. ELECTROPHYSIOLOCICAL EVIDENCE CHANNELS BY PROTEIN PHOSPHORYLATION

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Neurotransmitter-Regulated K in Sensory Neurons of Aplysia

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Three examples of potassium channel modulation by phosphorylation involve identified neurons in the marine mollusc Aplysia californica. One such case involves behavioral sensitization of the defensive gill withdrawal reflex in Aplysia (52). This behavioral adaptation is mediated by release of neurotransmitter onto the sensory nerve terminals that regulate the firing of the motor neurons initiating gill withdrawal. Initial studies suggested that facilitory transmission onto the sensory nerve terminals is mediated by serotonin. Application of serotonin prolongs the action potential (53) and increases excitatory transmitter output from these terminals (54). Serotonin treatment of the sensory neurons was found to increase intracellular levels of cyclic AMP (55). Application of dibutyryl cyclic AMP (54), or injection of cyclic AMP (54) or the purified catalytic subunit of cyclic AMP-dependent protein kinase (56) into the sensory neuron caused a response identical to that of serotonin. Injection of protein kinase inhibitor, which blocks cyclic AMPdependent protein kinase activity ( 5 3 , decreased or reversed the effect of serotonin on action potential duration and neurotransmitter output (58). Figure 3 depicts serotonin-induced prolongation of action potentials evoked in these sensory neurons (Fig. 3A), and inhibition of the effect of serotonin by injection of protein kinase inhibitor (Fig. 3B). Thus, continual activation of cyclic AMPdependent protein kinase is required for sensitization. Voltage-clamp studies identified a potassium channel as the target of modulation during sensitization. Both serotonin and cyclic AMP caused an increase in membrane resistance (53) and decreased outward potassium current (52, 59). Decreased potassium conductance delayed repolarization of the membrane po-

SEROTONIN 10-6Y

FIG.3. Effect of protein kinase inhibitor on modulation of action potential duration by serotonin. (A) The action potential of an Aplysia sensory neuron is prolonged by application of serotonin. (B) Prior injection of protein kinase inhibitor prevents the action of serotonin. Data are reproduced from Ref. (57).

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l

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FIG. 4. Single channel records from a cell-attached patch of sensory cell membrane show that serotonin induces a decrease in the number of active potassium channels. (A) Fmr channels are active before application of serotonin; after drug treatment, only one channel is active. (B) and (C). Expanded tracings from the indicated span of (A). Comparing (B) and (C), single channel current is unchanged by treatment with serotonin. Data are reproduced from Ref. (43).

tential to the resting level during the action potential, thus prolonging inward calcium current and causing enhanced release of neurotransmitter from the sensory neuron (52). The serotonin-sensitive potassium channel, called the S channel, was distinguished from previously identified potassium channels in being active at the resting membrane potential, insensitive to regulation by calcium, and only weakly voltage-dependent (60). Single channel recordings indicated that application of serotonin or intracellular injection of cyclic AMP caused a decrease in the number of active S channels, without effecting the mean channel current (59). The effect of serotonin on the activity of single S channels is shown in Fig. 4. Further observations have shown that S channels in excised patches of membrane can be closed by treatment with the catalytic subunit of cyclic AMPdependent protein kinase in the presence of ATP (61),implying that either the S channel itself or another membrane protein is subject to phosphorylation. 2. K + Channels Mediating the A Current Another example of cyclic AMP-induced depression of potassium conductance is found in the behavior of electrically coupled neurosecretory bag cells in the abdominal ganglion of Aplysia. Brief stimulation of these cells causes a prolonged discharge of repetitive action potentials lasting several minutes, accompanied by secretion of hormones that induce egg-laying behavior (62). Cyclic AMP-dependent phosphorylation is thought to promote the repetitive firing of these cells. Intracellular levels of cyclic AMP increased twofold during

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the prolonged period of discharge (63). Analogues of cyclic AMP could initiate repetitive discharge (64, 65), and the discharge could be prolonged by inhibiting phosphodiesterase activity (63). Using dispersed bag cells maintained in culture, it was shown that microinjection of the catalytic subunit of cyclic AMP-dependent protein kinase also induced a train of repetitive discharges (65). Voltageclamp studies of individual bag cells in culture implicated modification of potassium conductance. Treatment of cells with a cyclic AMP analogue (66) or a combination of forskolin and a phosphodiesterase inhibitor, Ro20- 1724 (67), caused a decrease in I,, a rapidly activated, transient K + current considered to be mediated by a distinct A current K + channel. The decrease in I, prevented rapid repolarization of the cell membrane following the action potential and thereby caused bursts of repetitive action potentials. Alkon and colleagues have described an example of classical conditioning in which I, may be modified by calcium and calmodulin-dependent phosphorylation. In the mollusc Hermissenda crassicornis, paired presentation of light with rotation results in long-term reduction of I, in type B photoreceptors (51). This decrease in I, is calcium-dependent (68) and can be mimicked by injection of phosphorylase kinase, a calcium and calmodulin-dependent enzyme (69). These results suggest that calcium and calmodulin-dependent phosphorylation may play a role in blocking I, after behavioral conditioning in this neuronal pathway. 3. Inward Rectifier K + Channels in Aplysia Neurons

In the identified neuron R15 of the abdominal ganglion of Aplysia, cyclic AMP-dependent phosphorylation induces an increase in potassium conductance. This neuron exhibits endogenous rhythmic cycles of repetitive action potentials with periods of hyperpolarization between bursts (70). Serotonin was shown to cause a decrease in membrane resistance, suppression of repetitive firing, and hyperpolarization of the cell (46). The potassium current modified by serotonin was identified as an inward rectifying current distinct from the rapid transient current I,, the delayed current I,, and the calcium-activated potassium current (71). These inward rectifying channels are activated by hyperpolarization. Therefore, they act to hold the membrane potential near the resting value and prevent bursts of action potentials. This response to serotonin is mediated by cyclic AMP. Analogs of cyclic AMP mimicked the action of serotonin on cellular activity (46). Injection of protein kinase inhibitor blocked the effect of serotonin on potassium conductance (47).

4. Calcium-Activated K

+

Channels

Calcium-activated K channels are regulated by both membrane potential and intracellular calcium concentration (42). The voltage dependence of channel activation is shifted to more negative membrane potentials as cytosolic calcium increases, resulting in synergistic activation by depolarization and calcium (41). +

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These channels play a critical role in determining the interval between bursts of action potentials by mediating prolonged, calcium-dependent hyperpolarization. De Peyer and colleagues (48) showed that injection of the purified catalytic subunit of cyclic AMP-dependent protein kinase increased the calcium-activated potassium current during evoked action potentials in neurons of the land snail Helix roseneri. Calcium-activated potassium channels may also be modified by cyclic AMPdependent protein kinase as part of the neuronal processes leading to simple learning. In the learning paradigm described earlier for Herrnissenda crassicornis, calcium-activated potassium channels may be influenced by cyclic AMPdependent phosphorylation. Injection of the catalytic subunit of cyclic AMPdependent protein kinase (49) induced a decrease in late potassium currents of type B photoreceptors which are considered to be mediated by calcium-activated potassium channels. These results point to convergent regulation of this ion channel by two different second messengers, calcium and cyclic AMP. Two examples of phosphorylation-dependent regulation of channel activity in vertebrate neurons have been reported. In hippocampal pyramidal cells from mammalian brain, noradrenaline was shown to inhibit calcium-activated potassium conductance and this effect was mimicked by intracellular perfusion of cyclic AMP and its stable derivatives (50). Treatment of pyramidal cells with phorbol esters, which activate protein kinase C, blocked the slow, calciumdependent potassium current responsible for the afterhyperpolarizing potential in these cells (142, 143). Thus, modulation of calcium-activated potassium channels as described in molluscs also occurs in vertebrates. Further investigation may show that phosphorylation-dependent regulation of other potassium channels is prominent in vertebrates as well.

B . FUNCTIONAL SIGNIFICANCE OF PHOSPHORYLATION It is clear from the previous discussion that a variety of potassium channels are modified by calcium- or cyclic AMP-dependent phosphorylation. The unique characteristics of each potassium current and its contribution to the overall electrical response of the cell defines what aspect of cellular function is modified. In this way, phosphorylation events can influence diverse aspects of cellular behavior. In the case of Aplysia neuron R15, changes in potassium conductance modify the form of ongoing electrical activity of the cell. The broader physiological consequences of this action are not known. For the neurosecretory bag cells, inhibition of potassium channels allows the initiation of repetitive electrical activity which triggers hormone release and initiation of egg-laying behavior in Aplysia (65). The inhibition of calcium-activated potassium channels in hippocampal pyramidal cells regulates the adaptation of these cells to high-frequency stimulation (50).

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The alteration of potassium channel function by cyclic AMP-dependent phosphorylation has perhaps its most far-reaching consequences in mediation of behavioral adaptation of the gill-withdrawal reflex. Evidently, this forms the basis for sensitization of this behavioral pathway which is a simple form of learning. The short-term “memory” of this learned behavior lasting up to an hour seems to reside in a prolonged elevation of cellular cyclic AMP rather than in stable phosphorylation events (58). This represents the first description of a biochemical pathway underlying learned behavior.

C. BIOCHEMICAL ANALYSIS OF POTASSIUM CHANNEL PHOSPHORYLATION Voltage-sensitive potassium channels have not been isolated and their protein components are not identified. Therefore, it is not known whether direct phosphorylation of protein components of potassium channels is a likely mechanism of regulation. However, the modulation K channel properties in excised membrane patches implies that a membrane protein is the site of phosphorylation (61). Cyclic AMP-dependent phosphorylation of membrane proteins that occurs concomitant with regulation of potassium channels has been described in several molluscan neuronal preparations (72-76). However, the low density of ionic channels in neuronal membranes makes it unlikely that the major phosphoproteins observed in these studies are components of potassium channels. Further work with purified preparations of potassium channels will be required to determine whether regulation involves direct phosphorylation of channel components. +

IV. Acetylcholine Receptor The nicotinic acetylcholine receptor is a neurotransmitter-activated ion channel that mediates excitation at the neuromuscular junction, postganglionic neurons of the autonomic nervous system, and electric organs of electrogenic fish. This receptor has been studied extensively, both biochemically and electrophysiologically. The nicotinic receptor purified from fish electroplax consists of four structurally homologous subunits, a (50 kDa), p (54 kDa), y (56 kDa), and S (58 kDa), associated as a pentamer, a2pyS (77). Upon reconstitution into artificial liposomes or planar lipid bilayers, the purified receptor exhibits full biological activity, indicating that this pentameric structure is sufficient to account for the behavior of the native channel (78-81). The acetylcholine receptor from electroplax can be phosphorylated by several different kinases (82-84), suggesting regulation by protein phosphorylation.

348

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SANDRA ROSSIE AND WILLIAM A. CATTERALL

BIOCHEMICALSTUDIESOF PHOSPHORYLATION

Preparations enriched in electroplax postsynaptic membranes contain both protein kinase (85-87) and protein phosphatase activity (87, 88). The acetylcholine receptor itself was found to be phosphorylated by endogenous kinase activity (86, 89). Initial studies indicated that both the y- (90) and $-subunits (89,90) of the receptor were phosphorylated, and indirect evidence suggested that the a- and P-subunits may also be substrates for phosphorylation (91-93). Moreover, chemical analysis of the purified subunits of the acetylcholine receptor showed that all subunits contain phosphoserine (94). Attempts to identify the kinase activity responsible for phosphorylating the receptor were confounded by two problems: the presence of phosphorylated contaminants in the membrane preparation which were within the same molecular weight range as the receptor subunits, and the presence of several kinases in the preparation active toward different subunits of the receptor. Thus, despite reports that cyclic nucleotides had no effect on receptor phosphorylation (85, 91, 92), it is now clear that endogenous cyclic AMP-dependent protein kinase phosphorylates both the y- and 6-subunits of the receptor in membrane preparations (82). Basal endogenous phosphorylation that occurred under the experimental conditions used in this study was inhibited by protein kinase inhibitor, and was thus attributed to cyclic AMP-dependent protein kinase. Using purified receptor, both y and 6 were phosphorylated by the purified catalytic subunit of cyclic AMP-dependent protein kinase to the extent of 1 mol 32P per mol receptor. The initial rates of phosphorylation of these subunits were comparable to those of other proteins effectively phosphorylated by this enzyme under the same experimental conditions. Since physiological substrates for this enzyme are generally phosphorylated more rapidly than other proteins, this result suggests that cyclic AMP-regulated phosphorylation of the receptor may occur in vivo. Although early studies reported conflicting results concerning phosphorylation of the acetylcholine receptor in Torpedo membranes by calcium and calmodulindependent protein kinase (92, 93), it appears now that phosphorylation of the receptor is not regulated by calcium and calmodulin. There are proteins in electroplax membrane preparations that are phosphorylated by endogenous calcium and calmodulin-dependent protein kinase and comigrate with subunits of the receptor during SDS gel electrophoresis. These phosphoproteins do not copurify with the acetylcholine receptor through affinity chromatography, however, and are therefore distinct from the receptor subunits (82). Preliminary studies (83)indicate the presence of endogenous protein kinase C in preparations of Torpedo cafifornica electroplax membranes. The 6-subunit of the acetylcholine receptor was phosphorylated in membranes by this endogenous activity. Protein kinase C purified from brain phosphorylated both the a-and 6subunits of purified receptor on serine residues (95).The activity of this enzyme

II.

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349

in membranes and in purified preparations was dependent on calcium and phosphatidylserine. Endogenous tyrosine kinase activity has also been measured in Torpedo californica postsynaptic membranes (84). When the activity of cyclic AMP-dependent protein kinase and protein kinase C in this preparation was inhibited, residual phosphorylation of the acetylcholine receptor was still detected. Phosphorylation under these conditions was stimulated by Mn2+, a characteristic of tyrosine kinase activity (96), and resulted in the incorporation of 32P into tyrosine residues of the p, y, and 6 subunits of the receptor. Thus, the Torpedo acetylcholine receptor is a phosphoprotein in vivo. The receptor may be phosphorylated by several distinct kinases present in postsynaptic membranes (Table I). Multiple sites exist on the receptor for each kinase. Huganir and colleagues have proposed that all of these phosphorylation sites reside near one another in a conserved region of each subunit which represent major intracellular loops in structural models of the receptor (84). Site-directed antibody studies support the proposed identification of cyclic AMP-dependent phosphorylation sites on the S and y subunits (144). B. FUNCTIONAL SIGNIFICANCE OF PHOSPHORYLATION The functional significance of phosphorylation of the acetylcholine receptor at several distinct sites by multiple protein kinases is not yet fully understood despite extensive investigation. Dynamic phosphorylation events are not responsible for channel gating or the development of desensitization, since the reconstituted purified receptor exhibits these properties in the absence of kinase or phosphatase activity (80, 81). However, it is possible that the state of receptor phosphorylation regulates transmitter binding, channel activation, ion conductance, or desensitization. Alternatively, phosphorylation events may regulate biochemical maturation, metabolic stability, or localization of the receptor. Hypotheses on the role of phosphorylation in the disposition of the acetylcholine receptor center on the biochemical and electrophysiological distinctions between mature receptors found at the vertebrate muscle motor endplate and extrajunctional receptors found in embryonic and denervated muscle. Acetylcholine receptors in embryonic and denervated muscle are dispersed over the entire cell surface (97), and are free to diffuse laterally within the membrane (98). In adult innervated muscle, receptors are clustered at the neuromuscular junction (97), and are immobilized (99).Junctional receptors are more metabolically stable than are extrajunctional receptors (100-102) and have a shorter mean channel open time (103, 104). Although both forms of receptor are the same apparent size, they can be distinguished immunologically (105, 106) and by isoelectric point (107). The molecular bases of these differences in receptor characteristics are not understood. It has been suggested that the state of receptor

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SANDRA ROSSIE AND WILLIAM A. CATTERALL TABLE I PHOSPHORYLATION OF SUBUNITS OF THE ACETYLCHOLINE RECEPTOR Subunit

Kinase activity

References

a

Protein kinase C Tyrosine kinase Cyclic AMP-dependent protein kinase Tyrosine kinase Cyclic AMP-dependent protein kinase Protein kinase C Tyrosine kinase

(83, 95)

P Y 6

(84)

(82) (84)

(82) (83) (84)

phosphorylation may account for or contribute to these properties. For example, the state of phosphorylation of the receptor may influence its metabolic stability or mobility by altering its ability to interact with the cytoskeletal matrix. Alternatively, the attachment of phosphate groups may modify channel gating properties. Saitoh and Changeux (90) showed that the adult and neonatal forms of the acetylcholine receptor from Torpedo murmorutu differed with respect to thermal stability and isoelectric point, the mature receptor being less sensitive to heat denaturation and more acidic than the neonatal form. Adult receptor appeared to be phosphorylated to a greater degree than was neonatal channel. Treatment of mature receptor with alkaline phosphatase rendered it more basic and more susceptible to heat denaturation. These results suggested that differences in phosphorylation underlay some of the distinguishing characteristics of adult and neonatal forms of the channel. A report by Rubin and colleagues (108) raises the possibility that tyrosine phosphorylation may prevent clustering of acetylcholine receptors. These investigators found that transformation of cultured chicken myotubes by Rous sarcoma virus TSNY68 prevented receptor clustering. This response was specific to infection with a transforming virus, suggesting that the src gene product, pp6OSrc.,a tyrosine kinase (109), mediates inhibition of clustering. In light of the discovery of tyrosine phosphorylation of receptor subunits (84), it will be interesting to consider the involvement of tyrosine phosphorylation of acetylcholine receptors in the clustering mechanism. Recent evidence indicates that cyclic AMP-dependent phosphorylation modifies the rate of desensitization of the acetylcholine receptor. In electrophysiological studies using intact muscle, forskolin, which stimulates cyclic AMP synthesis, increased the rate of receptor desensitization (145, 146). No effect on single channel conductance, mean open time, or the frequency of channel openings was observed (146). Huganir and colleagues (147)showed that an enhanced rate of receptor desensitization could result from phosphorylation of

I I.

REGULATION OF IONIC CHANNELS

35 I

the receptor itself. Measuring ion flux into vesicles containing reconstituted Torpedo receptor, they found that receptors desensitized more rapidly if they had been phosphorylated by cyclic AMP-dependent protein kinase prior to reconstitution. Treatment of cultured myotubes with activators of protein kinase C also enhanced desensitization of acetylcholine receptors (148). The region of the receptor subunits phosphorylated by these two kinases (84) may regulate desensitization in response to two distinct second messenger systems.

V. Sodium Channels The voltage-sensitive sodium channel mediates voltage-dependent sodium conductance that is responsible for the initial phase of the action potential in most excitable tissues. The sodium channel has been isolated and purified from several sources. Purified channels from eel electroplax (110-11.3), rat brain ( 1 1 4 , skeletal muscle (I 15, 116), and chicken heart (117)contain a similar high-molecularweight component (260 kDa). The sodium channel purified from rat brain consists of three glycoprotein subunits, designated a (260 kDa), p l (36 kDa), and p2 (33 kDa) (118, 119). A.

BIOCHEMICAL STUDIESOF SODIUM CHANNEL PHOSPHORYLATION

In contrast to the calcium and potassium channels, the initial evidence that sodium channels might be subjected to modulation by phosphorylation-dephosphorylation came from biochemical studies of the purified protein. Costa et al. (120) found that the a subunit of partially purified rat brain sodium channel could be specifically phosphorylated by the purified catalytic subunit of cyclic AMP-dependent protein kinase, to the extent of 3-4 mol 32P per mol functional channel. The rate of phosphorylation of the purified channel was comparable to that for an established substrate for this enzyme, suggesting that cyclic AMPregulated phosphorylation of the sodium channel may occur in vivo. Phosphorylation of the sodium channel by cyclic AMP-dependent protein kinase was also demonstrated in intact synaptosomes and lysed synaptosomal membranes (121). Incubation of lysed synaptosomal plasma membranes with added catalytic subunit of cyclic AMP-dependent protein kinase followed by isolation of the sodium channel subunits by immunoprecipitation and SDS-gel electrophoresis resulted in incorporation of 32P into a protein of 260 kDa (Figure 5A, lane 1). This was shown to be the a-subunit of the sodium channel by block of immunoprecipitation with highly purified sodium channel protein (Figure 5A, lane 2). If synaptosomes were first incubated with 8-Br-cyclic AMP to stimulate phosphorylation by endogenous protein kinase and ATP, subsequent phos-

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A

B

Fic. 5 . 8-Br-cyclic AMP stimulation of sodium channel phosphorylation in synaptosomes. (A) Lane I , SDS-PAGE of rat brain sodium channel after phosphorylation and immunoprecipitation, showing radioactively labeled a-subunit. Lane 2, incubation in the presence of excess unlabeled purified sodium channel prevents immunoprecipitation of labeled channel. Lane 3, sodium channel solubilized from synaptosomes, immunoprecipitated, then phosphorylated by exogenous purified catalytic subunit of cyclic AMP-dependent protein kinase. Lane 4,after stimulation of intact synaptosomes with 8-Br-cyclic AMP, sodium channel cannot be further phosphorylated by exogenous catalytic subunit, indicating that channels had been fully phosphorylated in siru during stimulation. (B) The time course of synaptosome stimulation with 8-Br-cyclic AMP. Data are reproduced from Ref. (121).

phorylation of the sodium channel by treatment with excess catalytic subunit and [y-32P]ATPafter synaptosomal lysis was completely blocked (Figure 5A, lanes 3 and 4). This result indicates that the sites on the a-subunit of the sodium channel that are phosphorylated in vitro are also phosphorylated in situ when cyclic AMP levels change. Stimulation of phosphorylation by 8-Br-cyclic AMP was complete within 15 s (Figure 5B) and was dose-dependent. Chromatographic analysis of tryptic digest fragments showed that four major sites of phosphorylation occurred on the cytoplasmic face of the sodium channel during endogenous phosphorylation. All four sites were protected against rephosphorylation in channels isolated from stimulated synaptosomes. Purified rat brain sodium channel and channel present in synaptosomal membranes can also be phosphorylated by purified protein kinase C (122). Phosphorylation of the a-subunit of purified channel occurred to the extent of 3-4 mol phosphate per mol channel. The initial rate of phosphorylation of purified sodium channel by this enzyme was comparable to that of histone H1, suggesting that the channel may be a good substrate for protein kinase C. Addition of diacylglycerol and phosphatidylserine did not enhance the activity of protein kinase C toward the purified sodium channel or channel in lysed synaptosomal membranes. The requirements of the enzyme were presumably satisfied by the

1I .

REGULATION OF IONIC CHANNELS

353

presence of detergent and lipid in the purified preparation and lipid present in membranes. Phosphorylation of the sodium channel in lysed synaptic membranes was calcium-dependent. Analysis of radioactively labeled phosphopeptides after tryptic digest indicated that two of the fragments detected after phosphorylation by protein kinase C comigrated with phosphopeptide fragments generated from cyclic AMP-dependent phosphorylation, whereas other fragments did not. It has not been determined whether the phosphorylated sites on these two proteolytic fragments are distinct or shared by cyclic AMP-dependent protein kinase and protein kinase C.

B . FUNCTIONAL SIGNIFICANCE OF PHOSPHORYLATION The relevance of phosphorylation events to sodium channel function is not known. As for the acetylcholine receptor, dynamic phosphorylation events would not be expected to mediate channel opening and closing. Phosphorylation may, however, modulate some electrically sensitive aspect of the channel, such as gating properties or conductance in the open state. Windisch and Tritthart found that norepinephrine and phosphodiesterase inhibitors increased inactivation of the cardiac action potential during prolonged depolarizations (123). This technique provides an indirect measure of sodium channel inactivation, but is not as rigorous a measure of this process as voltage-clamp procedures. Ribeiro and Sebastiao (124) found that cyclic AMP derivatives enhanced the block of the action potentials in sciatic nerve by tetrodotoxin, a specific sodium channel blocker (125).Costa and Catterall(121) showed that veratridine-stimulated 22Na influx in synaptosomes was significantly inhibited by treatment with 8-Br-cyclic AMP. This may be the result of a reduction in either the total number of open sodium channels or the unit conductance of open channels, or may reflect a change in the efficacy of neurotoxin activation of channels. Considered together, these results raise the possibility that cyclic AMP-dependent phosphorylation reduces sodium channel activation, but further work using voltage-clamp methods will be required to confirm this possibility. Phosphorylation events may modulate other aspects of the sodium channel, such as biosynthesis and processing, metabolic stability, or mobility and localization of the channel within the cytoplasmic membrane. Stuhmer and Almers (126) have shown that the sodium channel in frog skeletal muscle is immobilized within the sarcolemma, as was found in the case of junctional acetylcholine receptors (99). It is possible that phosphorylation is involved in association of the channel with the cytoskeletal matrix. Earlier, evidence for differential phosphorylation of neonatal and adult acetylcholine receptors was discussed. Immature forms of the sodium channel have also been described in fetal mammalian muscle (127), and in fetal rat brain (128). These channel subtypes may differ in phosphorylation state. Complete examination of possible physiological roles of

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phosphorylation of the sodium channel will require consideration of these modes of action in addition to effects on channel function on the millisecond time scale.

VI. Conclusions Protein phosphorylation is the most widely distributed process for regulation of protein function in biology. Ionic channels are a relatively recent addition to the long list of molecules regulated directly or indirectly by phosphorylation reactions. Physiological studies have implicated this pathway in the action of neurotransmitters in regulating the heartbeat and in a wide range of functions in the nervous system, including the acquisition and retention of simple learned behaviors. At present, there is no example where the molecular mechanism of this regulation is fully understood. In the case of calcium and potassium channels, the physiological consequences of protein phosphorylation are widely documented but sites of critical phosphorylation events are not fully characterized. In the case of acetylcholine receptors and sodium channels, regulated phosphorylation reactions have been described in vivo and in vitro but their physiological consequences are only partially defined. Future work in this area will focus on filling in the gaps in these regulatory pathways and on identifying new regulatory pathways through which protein phosphorylation may regulate ion channel function.

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56. Castellucci, V. F., Kandel, E. R., Schwartz, J. H., Wilson, F. D., Nairn, A. C., and Greengard, P. (1980). PNAS 77, 7492. 57. Krebs, E. G., and Beavo, J. A. (1979). Annu. Rev. Biochem. 48, 929. 58. Castellucci, V. F., Nairn, A., Greengard, P., Schwartz, J. H., and Kandel, E. R. (1982). J. Neurosci. 2, 1673. 59. Seigelbaum, S. A., Camardo, J. S., and Kandel, E. R. (1982). Nature (London) 299, 413. 60. Klein, M., Camardo, J., and Kandel, E. R. (1982). PNAS 79, 5713. 61. Shuster, M. J., Camardo, J. S., Siegelbaum, S. A,, and Kandel, E. R. (1985). Nalure (London) 313, 392. 62. Stuart, D. K., Chiu, A. Y., and Strumwasser, F. (1980). J. Neurophysiol. 43, 488. 63. Kaczmarek, L. K., Jennings, K. R., and Strumwasser, F. (1978). PNAS 75, 5200. 64. Kaczrnarek, L. K., and Strurnwasser, F. (1981). J. Neurosci. 1, 626. 65. Strumwasser, F., Kaczmarek, L. K . , and Jennings, K. R. (1982). FP 41, 2933. 66. Kaczmarek, L. K., and Strurnwasser, F. (1984). 1.Neurophysiol. 52, 340. 67. Strong, J. A. (1984). J. Neurosci. 4, 2772. 68. Alkon, D. L., Shoukimas, J . J., and Heldrnan, E. (1982). Eiophys. J. 40, 245. 69. Acosta-Urguidi, J., Alkon, D. L., and Neary, J. T. (1984). Science 224, 1254. 70. Levitan, I. B., and Adams, W. B. (1981). Adv. Cyclic Nucleotide Res. 14, 647. 71. Benson, J. A., and Levitan, I. B. (1983). PNAS 80, 3522. 72. Levitan, I. B., and Barondes, S. (1974). PNAS 71, 1145. 73. Paris, C. G., Castellucci, V. F., Kandel, E., and Schwartz, J. (1981). Cold Spring Harbor Conf. Cell Proliferation 8, 1361. 74. Jennings, K. R., Kaczmarek, L. K., Hewick, R. M., Dreyer, W. J., and Strumwasser, F. (1982). J . Neurosri. 2, 158. 75. Lemos, J. R., Novak-Hofer, I., and Levitan, I. B. (1982). Nature (London) 298, 64. 76. Lemos, J. R., Novak-Hofer, I., and Levitan, I. B. (1984). PNAS 81, 3233. 77. Changeux, J.-P., Devillers-Thiery, A., and Chemouilli, P. (1984). Science 225, 1335. 78. Wu, W. C.-S., and Raftery, M. A. (1979). EERC 89, 26. 79. Changeux, J.-P., Heidrnann, T., Popot, J.-L., and Sobel, A. (1979). FEBS Lett. 105, 181, 80. Huganir, R. L., Schell, M. A., and Racker, E. (1979). FEBS Lerr. 108, 155. 81. Nelson, N., Anholt, R., Lindstrom, J . , and Montal, M. (1980). PNAS 77, 3057. 82. Huganir, R. L., and Greengard, P. (1983). PNAS 80, 1130. 83. Huganir, R. L., Albert, K. A., and Greengard, P. (1983). Soc. Neurosci. Abstr. 9, 578. 84. Huganir, R. L., Miles, K., and Greengard, P. (1984). PNAS 81, 6968. 85. Gordon, A. S . , Davis, C. G., and Diamond, I. (1977). PNAS 74, 263. 86. Teichberg, V. I . , Sobel, A., and Changeux, J.-P. (1977). Narure (London) 267, 540. 87. Teichberg, V. I., and Changeux, J.-P. (1977). FEES Lett. 74, 71. 88. Gordon, A. S., Milfay, D., Davis, C. G., and Diamond, I. (1979). BERC 87, 876. 89. Gordon, A. S . , Davis, C. G., Milfay, D., and Diamond, I. ( I 977). Nature (London) 267,539. 90. Saitoh, T., and Changeux, J.-P. (1981). PNAS 78, 4430. 91. Saitoh, T.,and Changeux, J.-P. (1980). EJE 105, 5 1 . 92. Smilowitz, H., Hadjian, R. A., Dwyer, J., and Feinstein, M. B. (1981). PNAS 78, 4708. 93. Davis, C. G., Gordon, A. S., and Diamond, I. (1982). PNAS 79, 3666. 94. Vandlen, R. L., Wu, W. C.-S., Eisenach, J. C., and Raftery, M. A. (1979). Biochemistry 18, 1845. 95. Huganir, R. L., unpublished observations. 96. Swarup, G., Dasgupta, J. D., and Garbers, D. L. (1984). Adv. Enzyme Regul. 22, 267. 97. Fambrough, D. (1979). Physiol. Rev. 59, 165. 98. Axelrod, D., Ravdin, P., Koppel, D. E., Schlessinger, J., Webb, W. W., Elson, E. L., and Podleski, T. R. (1976). PNAS 73, 4594.

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99. Tank, D. W., Wu, E. S., and Webb, W. W. (1981). Biophys. J. 33, 74A. 100. Merlie, J.-P., Changeux, J.-P., and Gros, F. (1976). Nature (London) 264, 74. 101. Frank, E., Gautvik, K., and Sornmerschild, H. (1975). Acfa Physiol. Scand. 95, 66. 102. Linden, D. C., and Fambrough, D. (1979). Neuroscience 4, 527. 103. Michler, A , , and Sakmann, B. (1980). Dev. Biol. 80, 1. 104. Fischbach, G. D., and Schuetze, S. M. (1980). J . Physiol. (London) 303, 125. 105. Almon, R. R., and Appel, S. H. (1975). BBA 393, 66. 106. Weinberg, C. G., and Hall, Z. (1979). PNAS 76, 504. 107. Brockes, J. P., and Hall, Z. W. (1975). Biochemistry 14, 2100. 108. Anthony, D. T., Schuetze, S. M., and Rubin, L. L. (1984). PNAS 81, 2265. 109. Hunter, T., and Sefton, B. M. (1980). PNAS 77, 131 1. 110. Agnew, W. S., Moore, A. C., Levinson, S. R., and Raftery, M. A. (1980). BBRC 92, 860. 1 11. Nakayama, H., Withy, R. M., and Raftery, M. A. (1982). PNAS 79, 7575. 112. Miller, J. A,, Agnew, W. S., and Levinson, S. R. (1983). Biochemistry 22, 462. 113. Norman, R. I., Schmid, A,, Lombet, A,, Barhanin, J., and Lazdunski, M. (1983). PNAS 80, 4164. 114. Hartshorne, R. P., and Catterall, W. A. (1981). PNAS 78, 4620. 115. Barchi, R. L. (1983). J . Neurochem. 40, 1377. 116. Casadei, J. M., Gordon, R. D., Lampson, L. A , , Schotland, D. L., and Barchi, R. L. (1984). PNAS 81, 6227. 117. Lombet, A,, and Lazdunski, M. (1984). EJB 141, 651. 1 18. Hartshorne, R. P., Messner, D. J., Coppersmith, J. C.. and Catterall, W. A. (1982). JBC 257, 13888. 1 19. Hartshorne, R. P., and Catterall, W. A. (1984). JBC 259, 1667. 120. Costa, M. R. C., Casnellie, J. E., and Catterall, W. A. (1982). JBC 257, 7918. 121. Costa, M. R. C., and Catterall, W. A. (1984). JBC 259, 8210. 122. Costa, M. R. C., and Catterall, W. A. (1984). Cell. Mol. Neurobiol. 4, 291. 123. Windisch, H . , and Tritthart, H. A. (1982). J . Mol. Cell. Cardiol. 14, 431. 124. Ribeiro, J. A , , and Sebastiao, A. M. (1984). Br. J . Pharmarol. 81, 277. 125. Narahashi, T. (1974). Physiol. Rev. 54, 813. 126. Stuhmer, W., and Almers, W. (1982). PNAS 79, 946. 127. Kidokoro, Y., Heinemann, S., Schubert, D., Brandt, B. L., and Klier, F. G. (1975). CSHSQB 40, 373. 128. Schmidt, J., Rossie, S., and Catterall, W. A. (1985). PNAS 82, 4847. 129. Carbone, E., and Lux, H. D. (1984). Nature (London) 310, 501. 130. Nilius, B., Hess, P., Lansman, J . B., and Tsien, R . W. (1985). Nature (London) 316, 443. 13I . Bean, B. P. ( 1985). J. Gen. Physiol. 86, I . 132. Armstrong, C. M., and Matteson, D. R. (1985). Science 277, 65. 133. Cognard, C., Lazdunski, M., and Romey, G. (1986). PNAS 83, 517. 134. Nowycky, M. C., Fox, A,, and Tsien, R. W. (1985). Nature (London) 316, 440. 135. Rane, S. G., and Dunlap, K. (1986). PNAS 83, 184. 136. Dunlap, K., and Fischbach, G. (1978). Nature (London) 276, 837. 137. Dunlap, K., and Fischbach, G. (1981). J. Physiol. (London) 317, 519. 138. Curtis, B. M., and Catterall, W. A. (1986). i9iochemistr.y 25, 3077. 139. Flockerzi, V . , Oeken, H.-J., Hofmann, F., Pelzer, D., Cavalie, A,, and Trautwein, W. (1986). Nature (London) 323, 66. 140. Schmid, A,, Renaud, J.-F., Lazdunski, M. (1985). J. B i d . Chem. 260, 13041. 141. Arreola, J . , Calvo, J., Garcia, M. C., and Sanchez, J . A. (1986). Biophys. J. 49, 197a. 142. Baraban, J. M., Snyder, S. H . , and Alger, B. E. (1985). PNAS 82, 2538. 143. Malenka, R. C., Madison, D. V., Andrade, R., and Nicoll, R. A. (1986). J. Neurosci. 6,475.

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144. Souroujon, M., Neumann, D., Pizzighella, S . , Fridkin, M., and Fuchs, S. (1986). EMBOJ. 5, 543. 145. Middleton, P., Jaramillo, F., and Schuetze, S. M. (1986). PNAS 83, 4967. 146. Albuquerque, E. X., Deschpande, S. S . , Aracava, Y . , Alkondon, M., and Daly, J . W. (1986). FEBS Lett. 199, 113. 147. Huganir, R. L., Delcour, A. H., Greengard, P., and Hess, G. P. (1986). Nature (London)321, 114. 148. Eusebi, F . , Molinaro, M., and Zani, B . M. (1985). J . Cell. Biology 100, 1339.

12

Regulation of Protein Synthesis IRVING M. LONDON* DANIEL H. LEVIN? ROBERT L. MATTS* N. SHAUN B. THOMAS* RAYMOND PETRYSHYN" JANE-JANE CHEN*

* Harvard-M.I.T. Division of Health Sciences and Technology fDepartment of Biology Massachusetrs Institute of Technology and Harvard Universiiy Cambridge. Massachusetts 02 139

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Initiation of Protein Synthesis in Eukaryotic Cells ..................... 111. Role of eIF-2 in Eukaryotic Protein Chain Initiation and the Effect of eIF-2a Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. eIF-2 Recycling .................................. B. Effect of eIF-2a n on the Interaction of RF and eIF-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Does the Formation of an RF.eIF-2(aP) Complex Take Place in ....... Situ and Account for the Inhibition of Protein Synthesis? D. Site of Action of RF in the Recycling of eIF-2 .................... IV. Heme-Regulated eIF-2a Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. dsRNA-Dependent eIF-2a Kinase VI. Biological Significance of HRI and VII. Guanine Nucleotide-Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

360 360 362 362 364 366 368 369

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359 THE ENZYMES, Vol. XVlll Copyright 8 1987 by Academic Press, Inc. All rights of reproduction in any form reserved.

360

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introduction

The findings that heme controls the synthesis of protein in reticulocytes, and that in heme-deficiency diminished initiation with disaggregation of polyribosomes occurs ( I , 2) were made in the mid- 1960s. The mechanisms involved in this regulation are gradually being elucidated. The principal mechanism is the inhibition of initiation of protein synthesis as the result of phosphorylation of the a-subunit of the eukaryotic initiation factor 2, eIF-2a. In addition to heme-deficiency, oxidized glutathione (GSSG) and low levels of double-stranded RNA inhibit initiation by promoting phosphorylation of eIF-2a. This chapter deals with the regulation of initiation by the phosphorylation of eIF-2a; the effects of this phosphorylation on the exchange of GTP for GDP which is essential for the recycling of eIF-2; the activation, activity, and characteristics of the eIF-2a kinases; and the distribution and function of these kinases in nonerythroid cells. To begin, however, it may be useful to present very briefly the accepted view of the initiation of protein synthesis in eukaryotic cells [for reviews, see Refs. (3-5)l.

Ii. Initiation of Protein Synthesis in Eukaryotic Cells The translation of mRNA in eukaryotic cells occurs in the cytoplasm. In the first step of initiation, free 80 S ribosomes are in equilibrium with their 40 S and 60 S subunits (Fig. I). In the presence of eIF-3,40 S subunits bind the eIF-3 and eIF-4C to form a 43 S ribosomal complex; the binding of eIF-3 and eIF-4C to the 40 S subunit inhibits the joining of the 60 S subunit. In the next step, eIF-2 binds GTP and the initiator tRNA, Met-tRNA,, in a ternary complex. The binding by eIF-2 is specific for both guanine nucleotides and for Met-tRNA,. The ternary complex now binds to the 43 S ribosomal complex to form the 43 S preinitiation complex. It is to the 43 S preinitiation complex that mRNA is bound. This binding occurs in an ATP-dependent reaction in which eIF-4A, eIF-4B, and eIF-4F form a complex with mRNA. The product of the binding of mRNA to the 43 S preinitiation complex is the 48 S preinitiation complex. In this complex the cap structure is bound close to the ribosome and the AUG initiator codon is downstream from the cap structure. The anticodon of the Met-tRNA, probably participates in the scanning and recognition of the AUG codon. The joining of the 48 S preinitiation complex and the 60 S subunit is catalyzed by eIF-5 which has a ribosome-dependent GTPase activity. The joining reaction is accompanied by the release of the initiation factors eIF-3, eIF-4C, and eIF-2;

36 1

12. REGULATION OF PROTEIN SYNTHESIS

m @\@* @ @u

0

+

GTP t Met- tRNAf

43s RIBOSOMAL COMPLEX

4

C" 2 .GTP.Mel-tRNA,

TERNARY COMPLEX -1RNAf

@@@GTP.Met

u

43s PRElNlTlATlON COMPLEX

mRNA @@@.GTP.

Met-1RNAf

485 PRElNlTlATlON COMPLEX

'LL

@.GDP BINARY + P,,

Q Met-1RNAf

@, @, @

COMPLEX

80s INITIATION COMPLEX

FIG.1. Summary of eukaryotic initiation. Numbers in circles refer to eukatyotic initiation factors.

elF-2 is released as a binary complex, eIF-2.GDP. The product of the joining reaction is the 80 S initiation complex. Formation of the active 80 S initiation complex is the final step in initiation. The Met-tRNA, is positioned in the P (peptidyl) site on the ribosome for the start of polypeptide elongation. The sequence of steps in the process of initiation affords several opportunities for the exercise of regulatory mechanisms. These include the recycling of eIF-2 after its release as the eIF-2.GDP complex in the joining reaction; the formation of the ternary complex; and the relative affinities of mRNAs for eIF-2 and for eIF-4A, -4B, and -4F in determining the relative rates of translation of the mRNAs (6). Our discussion is concerned with the regulatory effects of phosphorylation of eIF-2a on the recycling of eIF-2.

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Role of elF-2 in Eukaryotic Protein Chain Initiation and the Effect of elF-2a Phosphorylation

The initiation of protein synthesis in reticulocytes and their lysates is inhibited by heme-deficiency, double-stranded RNA, or oxidized glutathione (7-1 1). In lysates, the inhibitions are characterized by a brief period of control linear synthesis, followed by an abrupt decline in this rate and by disaggregation of polyribosomes. These inhibitions of protein synthesis are associated with a decrease in the formation of the eIF-2*Met-tRNAf*GTPternary complex and the 40 S.eIF-2*Met-tRNAc-GTP 43 S initiation complex (12-23). The fundamental mechanism responsible for the inhibition is the activation of CAMP-independent protein kinases that specifically phosphorylate the 38-kDa a-subunit of eIF-2 (eIF-2a) (12,24-29). Dephosphorylation of eIF-2a accompanies the recovery of protein synthesis upon addition of hemin to inhibited heme-deficient lysates (30). Early studies suggested that the principal effect of the inhibitor, activated during heme-deficiency (HRI) in the reticulocyte lysate, was the impairment of the recycling of eIF-2 (31). Subsequently, partially purified factors, which stimulated ternary complex formation by interacting with eIF-2, were isolated in several laboratories from reticulocytes (18-21). In vitro studies at that time prompted the suggestion that the interaction of eIF-2 with these stimulatory factors was inhibited by phosphorylation of eIF-2a. However, protein synthesis in reticulocyte lysates was found to be shut off when only 20-40% of the eIF-2a present was phosphorylated (32, 33). The suggestion of inhibited interaction of phosphorylated eIF-2 with the stimulatory factors failed to explain why the 60% of more of the eIF-2 which was not phosphorylated could not function in the cycle of initiation. Clearly, another mechanism must be involved and some other factor must become rate-limiting as a result of phosphorylation of eIF-2a. A.

EIF-2 RECYCLING

The recycling of eIF-2 requires GTP-GOP exchange, which is catalyzed by the reversing factor (RF). A binary complex between eIF-2 and GDP is formed at the final step of initiation as the GTP present in the ternary complex is hydrolyzed upon the joining of the 48 S initiation complex and the 60 S ribosomal subunit (34-37). eIF-2.GDP is a highly stable complex, such that the GDP bound to eIF-2 dissociates at an unusually slow rate (t,,, > 30 min at 30°C) at physiological Mg2+ concentrations (38-42). In addition, the binding affinity of eIF-2 for GDP is approximately 100-fold greater than that for GTP (Kd GDP-3.1 X lop8 M versus Kd GTP-2.5X l o p 6 M ) (43, 44). This difference in affinity and the slow dissociation rate of the eIF-2.GDP complex under physiological conditions account for the very potent ability of GDP to inhibit ternary complex formation

12. REGULATION OF PROTEIN SYNTHESIS

363

(38-45).Therefore, the (eIF-2.GDP) complex formed at the final step of initiation in vitro would be essentially inactive in subsequent rounds of the initiation cycle in the absence of any mechanism to effect GTP-GDP exchange. How does phosphorylation of eIF-2a relate to this GTP-GDP exchange? Phosphorylation of eIF-2a has been found to have no effect on the binding affinity of eIF-2 for GDP (44).GDP also spontaneously dissociates from eIF-2 in the absence of Mg2 , a property of eIF-2 which is also not affected by the phosphorylation of eIF-2a (39, 40). These findings led to the suggestion that the mechanism by which eIF-2a phosphorylation brings about the inhibition of protein synthesis is not by a direct effect on the guanine-nucleotide-binding properties of eIF-2, but rather on the functional availability of RF. The eukaryotic reversing factor is a multipolypeptide factor that consists of 5 asymmetric subunits with approximate molecular weights of 82, 65, 5 5 , 40, and 32 kDa (40, 41, 44, 46, 47). RF was originally purified from rabbit reticulocyte lysates by assaying its ability to stimulate protein synthesis in heme-deficient lysates. Purified from both postribosomal supernatants and ribosomal salt washes, RF has been isolated in either a free form or complexed in a 1 :1 stoichiometry with eIF-2 (40, 41, 44, 46-49). RF is apparently similar, if not identical, to the partially purified factors discussed previously which stimulated ternary complex formation by eIF-2 (18-21). Addition of RF to heme-deficient reticulocyte lysates has been found to stimulate protein synthesis catalytically, with 20-40 pmol of globin synthesized in 30 min for each pmol of RF added (40, 44). Addition of eIF-2 to heme-deficient lysates has been found to result in, at best, a stoichiometric synthesis of globin chains (44). The RF functions in the initiation cycle to catalyze the recycling of eIF-2 by markedly stimulating the rate of dissociation of GDP from the eIF-2.GDP comW to eIF-2 lowers the binding affinity of eIF-2 plex (39-42, 50). The binding of F for GDP (Kd GDP=1.8X M ) and increases the affinity of eIF-2 for GTP (Kd GTP=l.7X M ) (44, 51). Since GTP is present physiologically at concentrations 10- to 20-fold greater than GDP (52, 53), the net effect is that upon the binding of RF to eIF-2.GDP, GTP is exchanged for GDP and, in the presence of Met-tRNA,, ternary complex formation is promoted (38-41, 50). The catalytic action of RF on the recycling of eIF-2 may be summarized as shown in reactions (1)-(4). +

+ (eIF-2GDP) e (RF.eIF-2) + GDP (RF.eIF-2) + GTP e RF + (eIF-2GTP) (eIF-2.Met-tRNArGTP) (eIF-2GTP) + Met-tRNA, (eIF-2.Met-tRNArGTP) + (40 S.eIF-3.eIF-4C) -+ 43 S initiation complex RF

(1)

(2) (3) (4)

Reactions (1)-(3) are apparently reversible. The reaction sequence is suggested by the following observations:

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LONDON, LEVIN, MATTS, THOMAS, PETRYSHYN, AND CHEN

1. The dissociation of the RF-eIF-2 complex is stimulated by GDP or GTP

(39, 40, 54). 2. RF, at high concentrations, has been reported to stimulate ternary complex dissociation in the presence of GDP (55, 56). 3. In the presence of 40 S ribosomal subunits the net reaction is RF-catalyzed formation of the 43 S initiation complex (55-57); no RF has been detected associated with the 43 S complex (44, 57-59).

However, the specific sequence of reactions involved in RF-catalyzed recycling of eIF-2 has not yet been agreed upon and alternative schemes have been suggested (57, 60, 61).

B. EFFECTOF EIF-2a PHOSPHORYLATION O N THE INTERACTION OF RF AND EIF-2 The observation that protein synthesis is inhibited by more than 95% when only 20-40% of the eIF-2a present in heme-deficient or dsRNA-treated lysates is phosphorylated led to the suggestion that the function of some factor, other than eIF-2, has become limiting (33). In v i m the phosphorylation of the asubunit of eIF-2 in the eIF-2GDP complex was found to inhibit the ability of RF to dissociate GDP catalytically from the eIF-2GDP complex (38-41). RF was found to bind tightly to eIF-2(aP)GDP complexes (40, 54) to form a complex from which GDP is readily exchangeable (58). However, the RF.eIF-2(aP) complex formed is essentially nondissociable with the result that the RF present in the complex is not functional (40, 54). This was demonstrated in in v i m assay systems in which the RF present in the RF.eIF-2(aP) complex was found to be unavailable to catalyze the dissociation of GDP from unphosphorylated eIF2GDP complex. As noted previously, the binding of RF to unphosphorylated eIF-2 is a reversible reaction, such that the dissociation of the RF.eIF-2 complex is stimulated in the presence of GTP or GDP. However, the addition of high levels of GTP or GDP does not stimulate the dissociation of the RFVeIF2(aP) complex, an indication that the binding of RF to phosphorylated eIF-2 is not a readily reversible reaction (62). High levels of GTP or GDP also do not stimulate the dissociation of GDP from eIF-2 by a mass action effect in in v i m systems (39, 60, 62). The observations on the interactions of RF and eIF-2 in in v i m systems led to the following hypothesis of the mechanism by which phosphorylation of eIF-2(a) results in the inhibition of protein synthesis in the reticulocyte lysate (40, 51, 54, 63): 1. The binary complex (eIF-2GDP) is formed at the final step of initiation, the joining of the 48 S complex with the 60 S ribosomal subunit.

2. The critical role of RF in the initiation cycle is to catalyze the dissociation

365

12. REGULATION OF PROTEIN SYNTHESIS

of eIF-2GDP to permit the formation of the ternary complex (eIF-2.MettRNA,GTP). 3. The binary complex, eIF-2GDP, is the primary site of action of eIF-2a kinases. 4. RF interacts with phosphorylated binary complex to form a RF.eIF-2(aP) complex that effectively sequesters RF so that it can not function in the recycling of the remaining unphosphorylated eIF-2GDP. Therefore, since RF is present in lysates at a much lower concentration than eIF-2, phosphorylation of only 20-40% of the eIF-2a present is sufficient to bind all the available RF in a nonfunctional complex with phosphorylated eIF-2. The hypothesis further predicts that restoration of protein synthesis in inhibited lysates requires the presence of a phosphatase which can dephosphorylate the RF.eIF-2(aP) complex to release functional RF. These reactions are shown schematically in Fig. 2 and may be summarized as shown in reactions (5)-(8). 48 S

+ 60 S

eIF-2GDP

-+

+ eIF-2GDP R F d F - 2 + GDP

80 S

+ RF S

(5)

(6)

+ ATP el~-2(a)kinaS)eIF-2(aP).GDP+ ADP RF.eIF-2(aP) + GDP eIF-2(aP)GDP + RF

eIF-2GDP

-+

(7)

(8)

High nonphysiological concentrations of GTP reverse or prevent inhibition of protein synthesis in heme-deficient lysates (64, 65). However, the observations [48S COMPLEX]

+ 60s

t-

[8OS COMPLEX]

+ el?,)

H R I or dsI

+ ATr

1 eIF-2(aP). GDP

FIG. 2. Effect of phosphorylation of eIF-2a on availability of RF for recycling of eIF-2.

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LONDON, LEVIN, MATTS, THOMAS, PETRYSHYN, AND CHEN

summarized above on the interaction of RF, eIF-2, and guanine nucleotides in vitro indicate that GTP does not act to reverse the inhibition of protein synthesis in heme-deficient lysates by a mass action displacement of GDP from the eIF-2.GDP complex (61, 62) or of RF from the RF*eIF-2(aP)(62) complex as has been suggested (44, 60). Data from several laboratories indicate that high levels of GTP act in heme-deficient lysates to inhibit the activation and/or the activity of HRI (62, 64-67), thus allowing the dephosphorylationof eIF-2(aP) in the RF-eIF-2(aP) complex, the release of functional RF, and the restoration of protein synthesis (62).

c.

DOESTHE FORMATION OF AN R F ' E I F - ~ ( ~COMPLEX P) TAKE PLACEI N SITU AND ACCOUNT FOR THE INHIBITION OF PROTEIN SYNTHESIS?

Verification of the hypothesis, derived from in vitro observations, for the mechanism by which eIF-2a phosphorylation leads to the inhibition of protein synthesis requires that the following conditions be met: 1. Changes in RF activity should correlate with alterations in the rate of protein synthesis and eIF-2a kinase activity. 2. An RF-eIF-2(aP) complex should be demonstrated, and only phosphorylated eIF-2 should be bound to RF when protein synthesis is fully inhibited. 3. Restoration of protein synthesis in inhibited lysates should be accompanied by the dephophorylation of eIF-2(aP) in the RF.eIF-2(aP) complex. 1.

Correlation of RF Activity, etF-2a Phosphorylation, and Protein Synthesis in the Reticulocyte Lysate The dissociation of GDP from eIF-2 is specifically catalyzed by RF, such that RF activity in whole or fractionated lysates can be measured by their capacity to stimulate the dissociation of exogenously added eIF-2.[3H]GDP (68). The relationship between protein synthesis and eIF-2-[3H]GDPdissociation activity (RF activity) can therefore be examined to determine whether there is a correlation between these two parameters as inhibition and restoration of protein synthesis are taking place. In reticulocyte lysates, when protein synthesis is inhibited by over 90% by heme-deficiency, dsRNA (20 ng/ml) or oxidized glutathione (500 pW), the rate of eIF-2.[3H]GDP dissociation was found to be reduced to 5% of that found in hemin-supplemented control lysates (68).The resumption of protein synthesis in heme-deficient lysates, which occurs upon the addition of hemin or high levels of GTP after shutoff, was found to correlate with the restoration of RF activity in these lysates (68). The loss of RF activity in the reticulocyte lysate, therefore, correlates with the known activation of eIF-2a kinase and the inhibition of protein synthesis. The finding that RF activity is lost in inhibited lysates, when

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only 20-40% of the eIF-2a is known to be phosphorylated, demonstrates that RF becomes a limiting component in the initiation cycle when protein synthesis is inhibited due to the activation of eIF-2a kinases. The observation that RF activity is lost and that eIF-2a phosphorylation reaches a plateau at approximately the same time at which protein synthesis shuts off provides further evidence of a relationship between RF activity and the level of eIF-2a phosphorylation in the reticulocyte lysate. 2. Localization of RF Activity and Phosphoiylated elF-2a in the Reticulocyte Lysate

When heme-deficient reticulocyte lysates are separated by centrifugation into ribosomal and nonribosomal (S 100) fractions, 90-95% of the eIF-2(aP) is associated with the ribosomal fraction, and the remaining 5-10% with the S 100 (58). A similar distribution of eIF-2(aP) has been reported in Ehrlich ascites cells (69). In hemin-supplemented lysates fractionated on sucrose density gradients, RF activity, localized by assaying for eIF-2.[3H]GDP dissociation, is present predominantly as a nonribosomal 15 S complex (58). In heme-deficient lysates or in lysates inhibited by dsRNA, the RF activity in nonribosomal fractions is markedly reduced (58). Alkaline phosphatase treatment of sucrose density gradient fractions derived from inhibited lysates results in the dephosphorylation of 8090% of the proteins present, including eIF-2(aP). Significant RF activity is released from nonribosomal fractions only (58).These findings indicate that RF exists functionally in the reticulocyte lysate primarily as a nonribosomal complex, and that active RF can be released upon dephosphorylation of the eIF-2(aP) to which the RF is bound. To establish the existence of the RF.eIF-2(aP) complex, heme-deficient reticulocyte lysates were fractionated on glycerol gradients to separate free eIF-2 (6 S) from eIF-2 associated with RF (15 S). All nonribosomal phosphorylated eIF-2(aP) was found to be associated in a 15 S complex with RF. In fully inhibited lysates all the eIF-2 associated with RF was found to be phosphorylated, an indication that when protein synthesis is shut off all available RF is sequestered in a non-functional 15 S complex with phosphorylated eIF-2 (58). 3. Restoration of Protein Synthesis and Dephosphorylation of e l F - Z ( d ) in Reticulocyte Lysates

In earlier studies, a decrease in the level of eIF-2a phosphorylation was observed when protein synthesis was restored in inhibited lysates upon addition of hemin or high levels of GTP (30, 66). Our working hypothesis requires that the eIF-2(aP) present in the RF-eIF-2(aP) complex be dephosphorylated for protein synthesis to resume; dephosphorylation of the bulk of the eIF-2(aP) present in the lysate would result simply in the generation of eIF-2.GDP com-

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plexes which are incapable of recycling in the absence of functional RF. Nonribosomal eIF-2(aP) has been reported to be dephosphorylated much more rapidly than the eIF-2(aP) present in the total lysate (70). The observation that all nonribosomal eIF-2(aP) is found associated in a 15 S complex with RF (58) suggests that the eIF-2(aP) bound in the RF-eIF-2(olP) is the primary site of eIF-2(aP) dephosphorylation in the reticulocyte lysate. Addition of hemin or high levels of GTP to heme-deficient lysates, after the shut-off of protein synthesis, results in a time-dependent release of RF activity and restoration of protein synthesis (62, 68). Ten minutes after the addition of 2 mM GTP, 70% of the RF activity has been found to be restored in these lysates, with a concomitant 65-75% decrease in the amount of the eIF-2(aP) associated with the 15 S RFaeIF-2 complex (62);RF activity correlates directly with the dephosphorylation of eIF-2(aP) in the RFaeIF-2 complex. The dephosphorylation of the RF.eIF-2(aP) complex and the release of functional RF are responsible for the restoration of protein synthesis by GTP. D.

SITE OF

ACTIONOF RF IN

THE

RECYCLINGOF EIF-2

The observation that the only nonribosomal eIF-2(aP) present in the lysate is associated with RF led to the proposal that ribosome-associated eIF-2(aP)GDP complexes may be released from the ribosome on interaction with RF (58).This hypothesis is supported by recent work on the localization of eIF-2a phosphorylation in inhibited lysates (59). During the initial stage of incubation of heme-deficient or dsRNA-treated reticulocyte lysates, when protein synthesis is occurring at control linear rates and polyribosomes are still intact, eIF-a(aP) is found associated with several ribosomal sites. eIF-a(aP) is observed in 40 S complexes, but surprisingly, it is also observed bound to free 60 S ribosomal subunits, and to 60 S subunits of 80 S ribosomes and polyribosomes (59, 71). After disaggregation of polyribosomes and shutoff of protein synthesis, eIF2(aP) accumulates primarily on free 60 S ribosomal subunits and on the 60 S subunits of 80 S ribosomal couples (59, 71). While this observation suggests a function for polyribosome-bound eIF-2(aP) in the mechanism of inhibition of protein synthesis, it is not clear if the 60 S-bound polyribosomal eIF-2(aP) derives from the incorporation into polyribosomes of 40 S initiation complexes containing eIF-2(aP), or is due to the direct phosphorylation of polyribosomal eIF-2 during elongation or termination. The eIF-2(aP) associated with the 60 S subunit appears to be present as the binary complex [eIF-2(aP).GDP] (59, 71). Reversal of the inhibition of protein synthesis in heme-deficient lysates by the addition of RF results in a rapid binding of RF to 60 S subunits, a concomitant dissociation of 60 S-bound eIF-2(aP)GDP, and a decrease in the amount of free 60 S ribosomal subunits present in the lysate as polyribosomes re-form (59).Unphosphorylated eIF-2 can

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bind to 60 S ribosomal subunits and recycle through polyribosomes in normal protein synthesizing lysates (59, 71). Therefore, the binding of eIF-2 to 60 S ribosomal subunits is likely to reflect a normal physiological function, and is not simply a characteristic of inhibition or a unique property of eIF-2(aP). The phosphorylation of eIF-2a and the sequestration of RF result not only in the inhibition of recycling of eIF-2 because of diminished GTP-GDP exchange but also in inhibition of release of eIF-2.GDP from 60 S ribosomal subunits. The relationship of the binding of eIF-2 by 60 S subunits to the sequence of events in protein chain initiation remains to be elucidated. One hypothesis assumes that the eIF-2 present in the 48 S preinitiation complex is transferred during the joining step to the 60 S subunit of an 80 S initiation complex or to a polyribosomal 60 S site, where it remains until it is released by the action of RF. It is not clear to what extent other components such as the joining factor, eIF-5, or events such as GTP hydrolysis, may be involved in mediating the transfer of eIF-2; nor can the possibility be dismissed that eIF-2 may have a function in elongation or termination. Alternatively, the 60 S ribosomal subunit may be acting as a matrix for the binding of eIF-2 and, possibly, other factors. Such a structure might enhance the efficiency by which terminating 80 S ribosomes are reutilized in the initiation cycle by increasing the local concentration of eIF-2 rather than requiring that ribosomal subunits interact with freely diffusible initiation factors. We emphasize that this hypothesis is preliminary and that further interpretation requires a quantitative analysis of the distribution of eIF-2.

IV. Heme-Regulated elF-Pa Kinase The eIF-2a kinases activated in the absence of heme (HRI) or by the addition of dsRNA (dsI) are different molecular species. HRI is also activated by GSSG, but this activation mechanism takes place in the presence of hemin and apparently proceeds through a different pathway. Both in v i m and in situ, HRI and dsI appear to phosphorylate the same single serine residue of eIF-2a (72). Purified HRI has been characterized as a dimer (- 150 kDa) of a single polypeptide (73, 74) with a sedimentation coefficient of 6.6 S in glycerol gradients (75). In SDS-PAGE, HRI migrates as a 80-90 kDa monomer. Activation of HRI both in vitro and in the lysate is accompanied by phosphorylation of HRI. In vifro, HRI is activated through an autokinase mechanism and is most active in the multiply phosphorylated state (73, 74, 76, 77). At least 3 phosphates per HRI subunit can be accommodated. HRI is most efficiently activated in vitro by treatment with N ethylmaleimide (NEM) (3-5 mM) which by interacting with free sulfhydryl groups permits an increase of up to 5 phosphate residues per HRI subunit introduced by autophosphorylation, with a concomitant increase in eIF-2a kinase activity (77). The enhanced activation produced by this chemical modification

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indicates that the maintenance of certain sulfhydryl groups in the reduced state plays a role in HRI activation. Recent studies suggest that internal sulfhydryl groups, which may not be accessible to NEM, are also involved in the mechanism of activation (unpublished data from this laboratory). The interaction of NEM with HRI sulfhydryl groups apparently stabilizes HRI in a molecular conformation that exposes additional autophosphorylation sites. However, it is not clear as yet how increased autophosphorylation enhances HRI activity. Several possibilities to be considered include ( a ) an increased stability of the active site; (b) increased affinity of HRI for eIF-2 or ATP; (c) an enhanced efficiency owing to some phenomenon associated with the increased charge; or (6) some combination of these. Nor is it clear whether HRI activity is a property of its monomeric or dimeric conformation. One attractive model includes an activation step in which the two subunits of the dimeric form phosphorylate each other, thus permitting the eIF-2a kinase activity to be expressed by the monomeric enzyme; however, this model has yet to be tested. The activation of HRI in v i m is blocked or decreased by treatment with hemin (5-20 k M ) which selectively binds to HRI (77), and is a principal physiological regulator of HRI activity in reticulocytes. Pretreatment of HRI with hemin diminishes its ability to autophosphorylate, to phosphorylate eIF-2a, and to inhibit protein synthesis in lysates. Hemin also partly diminishes the level of activity of NEM-treated HRI which retains a reduced capacity to bind hemin. In heme-deficient lysates, endogenous HRI is also activated by phosphorylation. It is not known, however, whether this phosphorylation occurs through an autokinase or heterokinase mechanism, although one study (78) suggests that an HRI-activating kinase is present in lysates. Nor is it clear if multiple phosphorylation of HRI occurs in situ. Both in vitro and in situ, the phosphorylation of HRI does not involve a CAMP-dependent protein kinase (79). Phosphoprotein analyses of protein synthesizing lysates reveal that compared to the phosphoprotein profile observed in uninhibited control lysates, heme-deficient lysates display only two additional proteins that are phosphorylated, HRI (85 kDa) and eIF-2a (38 kDa). The phosphate on eIF-2a turns over rapidly in inhibited lysates with a half-life of 2-3 minutes, due to the presence of a potent eIF-201 phosphatase activity, which has been tentatively identified as a type 1 protein phosphatase (80). The phosphate(s) on endogenous HRI also appear to turn over but at a much slower rate (30). Hence, the reversal of inhibition by the delayed addition of hemin (20 p M ) , MgGTP (2 mM), or CAMP (20 mM), all of which appear to act by blocking HRI activation and activity, permits a rapid dephosphorylation of endogenous eIF-2(aP) and a slower dephosphorylation of HRI . The restoration of protein synthesis in lysates inhibited by heme deficiency requires glucose-6-P as well as hemin (81). This requirement is readily demonstrated in lysates depleted of low molecular weight components by filtration in

12. REGULATION OF PROTEIN SYNTHESIS

37 1

dextran gels (81, 82). In gel-filtered lysates, both hemin (20 pM) and glucose-6P (50-500 pM) are required for optimal protein synthesis. The omission of either component permits partial HRI activation; when both components are omitted, HRI activation is maximal. Two unrelated functions of glucose-6-P contribute to the maintenance of protein synthesis: ( a ) the generation of NADPH, and (b) an effector or cofactor function at some step in initiation. Those sugars such as fructose-6-P, ribose-5-P, and sedoheptulose-7-P, which are readily converted to glucose-6-P by lysate enzymes fulfill both functions. 2-Deoxyglucose-6-P which can generate some NADPH owing to its oxidation by glucose-6-P dehydrogenase is as effective as glucose-6-P in both functions; fructose-l,6-diP and the triose intermediates of glycolysis are ineffective. The requirement for NADPH generation is associated with the maintenance of certain essential proteins in the sulfhydry1 form, a function which is mediated by the thioredoxin-thioredoxin reductase systems (83). In heme-deficient gel-filtered lysates, these proteins (but not eIF-2) are oxidized to the disulfide state (84, 85). In heme deficiency one or more of these disulfide proteins may be involved in HRI activation (83, 84). The effector function of glucose-6-P is unknown. Based on in vitro studies, no direct effect of glucose-6-P on purified HRI can be demonstrated. In situ studies in lysates suggest that the sugar phosphate regulates HRI activation indirectly, perhaps by acting as an allosteric effector on regulatory proteins, or by a stimulatory effect on an HRI phosphatase activity (78, 85). The generation of NADPH may also be necessary to maintain the large pool of reticulocyte glutathione (1-3 mM) in the reduced state (GSH), a function mediated by glutathione reductase. The oxidation of a small proportion of GSH to GSSG is sufficient to activate HRI and thereby inhibit protein synthesis. HRI activation is readily demonstrated in lysates by the addition of 20-200 p M GSSG (29, 81). The activation of HRI by GSSG in situ occurs by an indirect mechanism, since GSSG has little or no effect on HRI in vitro. The activation mechanism operates in the presence of heme, and apparently proceeds through a pathway that is different from that of heme deficiency. It is not clear if the disulfide protein products observed in heme deficiency are involved in the GSSG mechanism as well. The observation that HRI is activated by more than one pathway in lysates is supported by findings that indicate that certain physical stimuli such as pressure (74) and elevated temperature also give rise to HRI activation in situ.

V. dsRNA-Dependent elF-2a Kinase In contrast to HRI, which is present in the high-speed supernate (S loo), the dsRNA-activated eIF-2a kinase (dsl) is associated with the ribosomal complement (12, 28). Based on this and other criteria including molecular weight, sedimentation coefficient, immunological properties, and mechanism of activa-

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tion, HRI and dsI are different molecules. Although a rationale for the regulatory role of heme-regulated eIF-2a kinase seems self-evident, the function of dsI in reticulocytes is far less clear. Nonetheless, both activities produce inhibitions by the phosphorylation of eIF-2a; and both inhibitions are blocked or reversed by high concentrations of CAMP (20 mM), GTP (2 mM), and a variety of other purine derivatives, as well as stoichiometric levels of eIF-2, and catalytic levels of reversing factor (RF). dsl has been purified in both its latent and active states (28, 86). Both forms are 67 kDa single polypeptides with sedimentation coefficients of 3.7 S in glycerol gradients. Activation of latent dsI in v i m or in siru requires dsRNA (1-20 ng/ml) and ATP, and is accompanied by the dsRNAdependent phosphorylation of a polypeptide doublet of 70-72 kDa as monitored in SDS-PAGE. The two components represent different phosphorylated states of the same dsI polypeptide (87). Increased dsI activity correlates with increased phosphorylation and accumulation of the 72 kDa component. Kinetic data indicate that in the course of dsI activation the monophosphorylated 70 kDa component appears first, followed by the multiply phosphorylated 72 kDa component. The apparent shift-up is due to the slower migration of the multiply phosphorylated form in SDS-polyacrylamide gel electrophoresis (SDS-PAGE) (87). There is a rapid turnover of the phosphate(s) on dsI due to a type 2 protein phosphatase present in lysates which rapidly dephosphorylates the 72-kDa component producing an apparent shift down to the 70-kDa component with a concomitant loss in dsI activity (88, 89). An enigma of the activation mechanism is the paradoxical effect of dsRNA: while low levels of dsRNA (ng/ml) inhibit protein synthesis and activate dsI in vitro and in siru, high levels of dsRNA (pg/ml) are ineffective. This phenomenon has provided a useful tool in studies on the mechanism of dsl activation. In lysates, low levels of dsRNA (1-20 ng/ml) produce the characteristic biphasic kinetics of inhibition, and a phosphoprotein profile that differs from control profiles by the presence of two additional phosphorylated proteins, dsI (70-72 kDa) and eIF-2a (38 kDa). Since dsI activation in situ takes place in the presence of hemin, HRI (85 kDa) remains in the latent state, and is not involved in the inhibition. The dsRNA-induced phosphorylation pattern of lysates is similar to that observed in dsRNA-treated extracts of interferon-treated cells (90-92). Interferon induces two dsRNA-dependent activities, the (2’-5’A), oligo synthetase (92) and a dsRNA-dependent eIF-2a kinase (90, 91). As with reticulocyte dsI, the dsIs produced in 3T3 mouse fibroblasts, HeLa cells, and monkey kidney cells (CV-1) all require dsRNA and ATP for activation (unpublished data from this laboratory). Although the phosphoprotein profiles of dsI activation are similar for all of these dsI species, the phosphoprotein doublets associated with the various dsI species show slightly different patterns of migration in SDS-PAGE. These differences are probably attributable to differences in the distribution and/or number of phosphate moieties which are optimal for each dsI species. However,

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all of the latent forms of these dsl species migrate as 67-kDa polypeptides, and all are immunoprecipitated by antibodies prepared against reticulocyte dsI (unpublished data from this laboratory).

VI. Biological Significance of HRI and dsl The regulation of protein synthesis by HRI is not restricted to hemoglobin but is a general mechanism controlling the initiation of synthesis of soluble and membrane-bound proteins in reticulocytes (93, 94). This regulatory system is not limited to the reticulocytes of the rabbit. A translational inhibitor with properties similar to those of rabbit HRI has been isolated from human erythroid cells (95). This inhibitor is found predominantly in reticulocytes with little or no inhibitor associated with mature erythrocytes (96).Several lines of evidence indicate that the inhibitor formed in human reticulocytes is similar or identical to rabbit HRI: The inhibitory activity is characterized by activation during incubation in the absence of hemin, or by treatment with NEM: the inhibitor when added to a lysate produces biphasic kinetics of inhibition; reversal or prevention of inhibition occurs on addition of eIF-2; the phosphorylation of eIF-2a and the hemeregulated eIF-2a kinase activity are blocked by immune serum prepared to rabbit HR (96, 97). Although the regulation of initiation of protein synthesis by hemin has been most extensively studied in rabbit reticulocytes, there is some evidence that indicates that the initiation of protein synthesis can be regulated by heme and by eIF-2a phosphorylation in nonerythroid cells. Hemin has been shown to enhance protein synthesis in intact Krebs I1 ascites tumor cells (94), and to stimulate translation of mouse globin mRNA and encephalomyocarditis virus RNA added to extracts prepared from Krebs 11 ascites tumor cells and uninduced Friend erythroleukemic cells (98, 99). In other cell-free systems, heme has been shown to restore protein synthesis in extracts prepared from iron-deficient platelets (100), and hemin (25-200 pM) is required to maintain the activity of HeLa cell extracts in the initiation of protein synthesis (101). The latter effect of hemin is attributable to prevention of formation of an inhibitor of protein synthesis initiation and to protection of a ribosome-bound initiation factor (101). Whether these effects of heme are due to prevention of activation of an HRI-like eIF-2a kinase activity similar to that found in reticulocyte lysates remains to be determined. The rate of initiation of protein synthesis in Ehrlich ascites tumor cells that have been deprived of an essential amino acid or glucose is reduced by 60% as compared to fed control cells (102, 103). This reduction is accompanied by a 77% decrease in formation of (40 SaMet-tRNA,) initiation complexes both in vivo (102) and in vitro (103). The defect in 40 SaMet-tRNA, complex formation in nutrient-deprived cells can be overcome by the addition of exogenous eIF-2

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(103), an indication of impaired eIF-2 activity. Recent studies have indicated that a substantial portion of the a subunit of eIF-2 (eIF-2a) is phosphorylated in

Ehrlich ascites cells (68). The relationship, if any, of this phosphorylation of eIF-2a to an inhibitory factor previously isolated from these cells (104) as well as the relationship of these activities to nutrient starvation has yet to be elucidated. Several other biological systems exhibit translational control at the level of peptide-chain initiation by formation of inhibitors with properties similar to those of HRI. In resting lymphocytes the formation of (40 SaMet-tRNA,) initiation complexes is reduced compared to that in cells stimulated to proliferate with phytohaemagglutinin; the addition of eIF-2 to extracts prepared from resting lymphocytes can restore initiation complex formation (105). In addition, resting lymphocytes contain an inhibitor of peptide chain initiation that inhibits protein synthesis in reticulocyte lysates with many properties similar to those of HRI. The activity of this inhibitor is decreased in lymphocytes stimulated with phytohemagglutinin (106).Whether this inhibitor diminishes the activity of eIF-2 by phosphorylation is unknown. The formation of (40 S-Met-tRNA,) complexes has also been shown to be decreased in male rat ventral prostate gland following orchiectomy and adrenalectomy (107, 108).The decrease in initiation complex formation results from a decreased eIF-2 activity but this activity can be restored within 10-30 min after injection with dihydrotestosterone. These results suggest that in the hormoneresponsive prostate gland, an androgen-responsive factor may be involved in regulating the initiation of protein synthesis. It is tempting to speculate that the decrease in eIF-2 activity may be the result of phosphorylation due to activation of an androgen-responsive eIF-2a kinase. Inhibitors of initiation of protein synthesis with properties similar to those of reticulocyte HRI have been isolated and purified to various extents from a variety of nonerythroid cells using purification procedures previously described for reticulocyte HRI. These inhibitors have been prepared from Ehrlich ascites tumor cells (104), wheat germ extracts (109), Krebs ascites cells (110), and erythroid precursor cells from mouse spleen (110). The best characterized inhibitor of initiation has been isolated from perfused rat liver (111)and from freshly isolated hepatocytes depleted of heme with allylisopropylacetamide (112). The effect of the rat liver inhibitor on protein synthesis, its chromatographic behavior during isolation, the reversal of inhibition by eIF-2 and its ability to phosphorylate eIF-2a indicate that it is an HRI-like protein kinase. However, the physiological significance of the rat liver inhibitor and the other HRI-like inhibitors in the regulation of protein synthesis remains to be determined. It seems likely that the HRI system found in erythroid cells may be more general and operate in many nonerythroid cells. It is also possible that effectors other than heme may be involved in activation of these HRI-like protein kinases. It has been shown that addition of the reduced and alkylated NH,-terminal

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fragment of procollagen results in an inhibition of peptide chain initiation in reticulocyte lysates (113, 114). This inhibition is accompanied by phosphorylation profiles similar to those observed in lysates inhibited by HRI (114). At present it is unclear whether these phosphoproteins represent phosphorylated HRI or eIF-2a. The inhibition of initiation of protein synthesis by low levels of dsRNA (10, 115), is due to the activation of a dsRNA-dependent eIF-2a kinase, dsl, described in Section IV (12, 28). Although dsI has been studied extensively in rabbit reticulocytes it is not restricted to these cells. A similar dsRNA-dependent eIF-2a kinase has been observed in normal human reticulocytes but not in mature erythrocytes (96). Furthermore, the results of several studies indicate the presence of a similar dsRNA-dependent eIF-2a kinase activity which is inhibitory of protein synthesis in extracts of interferon-treated nonerythroid cells (91, 116). Interferons are glycoproteins (designated a,p and y) that elicit antiviral responses in cells (117), and a variety of biological responses in normal and neoplastic tissues and cells [for review, see Ref. (118)l. The antiviral properties of interferon directed against many viruses are attributable in part to induction of two cellular activities that inhibit protein synthesis ( I 19), ( a ) the (2’-5’A),synthetase which upon activation by dsRNA polymerizes ATP into (2’-5’A),,linked oligoadenylates (120) which activate a cellular endoribonuclease that degrades cellular and viral RNA in virus-infected and IFN-treated cells and in vitro (121, 122); and (b) the dsRNA-dependent eIF-2a kinase, dsI (91,116). The addition of dsRNA to extracts prepared from interferon-treated Ehrlich ascites tumor or L cells results in the phosphorylation and activation of dsI with a concomitant phosphorylation of eIF-2a (91, 92, 116). Moreover, the phosphorylation of dsI and eIF-2a occurs in vivo in cells treated with interferon and infected with viruses containing dsRNA genomes (119, 122-124). Thus it appears that dsI represents at least one mechanism by which various cells respond to viral invasion. The high endogenous levels of latent dsI in reticulocytes are unexplained. They may be the result of sensitization of the reticulocyte to circulating interferons in vivo or due to expression of dsI in an interferon-independent manner as a consequence of the terminal maturation process of the reticulocyte. Whatever the reason, it appears likely that the addition of dsRNA to reticulocyte lysates mimics the effect of virus infection in interferon-treated cells. In addition to their antiviral properties interferons have been reported to modulate cell growth and differentiation in a variety of cells (125-128). The mechanism of this action of interferon is not known; however, the interferon-induced and dsRNA-dependent (2’-5’A),-synthetase has been implicated in the regulation of growth and differentiation (128-131). Another study has demonstrated that mouse 3T3-F442A fibroblasts spontaneously produce and secrete interferon and exhibit a pattern of dsRNA-depen-

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dent phosphorylation of dsI that is related to various stages of growth and differentiation (132). The 3T3-F442A cells have normal fibroblast morphology in the growing state, but upon reaching confluence they undergo adipose differentiation with high frequency (133, 134). The dsRNA-dependent phosphorylation of dsI increases with cell growth until the cultures become confluent and protein synthesis is reduced. After confluence there is a rapid and marked decrease in phosphorylation of dsl and the cells begin to differentiate (132). The high level of dsI activity may be related to attainment of the resting state necessary for adipose differentiation. This dsI activity in turn is regulated by the amount of interferon produced and secreted by these cells. These studies lend support to the hypothesis of a physiological role for dsI in the regulation of growth and differentiation. Other studies raise the possibility that dsI may have regulatory functions independent of the interferon system. During the late stages of adenovirus infection the accumulation of a small RNA is required for maintaining efficient initiation of both cellular and viral protein synthesis (135). This RNA appears to prevent the phosphorylation of eIF-2a by inhibiting the activation of an eIF-2a kinase during adenovirus infection in HeLa cells (136). Although this eIF-2a kinase is not been definitively identified, it may prove to be dsI since high concentrations of dsRNA prevent its activation (135, 136). Another report suggests that the HeLa cell “host factor” which is required for the initiation of polio virus replication in virro is a phosphoprotein with many properties similar to those of dsI (137). Conclusive evidence relating to this identification will be awaited with interest.

VII. Guanine Nucleotide-Binding Proteins The RF-catalyzed exchange of GTP for GDP in the eIF-2.GDP complex is analogous to the Ts factor-catalyzed exchange of GTP for GDP in the TuGDP complex that is formed in prokaryote polypeptide elongation and to the GTPGDP exchange observed with similar factors in eukaryotic elongation. These analogies are part of a larger pattern of findings relating to guanine nucleotide-binding proteins. It is of particular interest that structural homologies are observed in these proteins. Transducin, a GTPase involved in light transduction in the vertebrate retinal rod cell, has structural homology with the stimulatory and inhibitory guanine nucleotide-binding proteins, Gs and Gi, of the hormone-sensitive adenylate cyclase systems; they each contain three subunits, the corresponding subunits have similar amino acid compositions (138), and the a-subunits contain sites for guanine nucleotide-binding and ADP-ribosylation (139-142). The a-subunit of transducin also has some homology at its COOH- and NH,-termini with similar

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regions of the ras oncogene products (143, 144). In other studies, the complete amino acid sequence of the transducin a-subunit has been deduced from the cDNA sequence and has revealed significant homologies of this protein with the prokaryotic elongation factors EF-Tu and EF-G, the prokaryotic initiation factor IF-2, and the rus proteins of man and yeast (145, 146). It will be of interest to determine whether eIF-2 shares in these homologies. In light of the effects of phosphorylation on the interaction of eIF-2 and RF, it should also be of interest to explore the possible role of phosphorylation in the protein-protein interaction of the various guanine nucleotide-binding proteins. In addition, the presence of ADP-ribosylation sites in the a-subunits of transducin, Gs, and Gi, as well as in eukaryotic elongation factor 2 (EF-2), and the presence of endogenous ADP-ribosyltransferase activity in rabbit reticulocytes and turkey erythrocytes (147, 148) raise interesting questions concerning the possible effects of ADP-ribosylation on the guanine nucleotide-bindingfunctions of these proteins and of eIF-2.

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15. 16. 17. 18. 19. 20. 21.

22.

Bruns, G. P., and London, I. M. (1965). BBRC 18, 236. Grayzel, A. I. P., Horchner, P., and London, I. M. (1966). PNAS 55, 650. Kramer, G., and Hardesty, B. (1980). Cell B i d . 4, 69. Jagus, R., Anderson, W. F., and Safer, B. (1981). Prog. NucleicAcidRes. Mol. Biol. 25, 127. Thomas, A. A. M., Benne, R., and Voorma, H. 0. (1981). FEBSLett. 128, 177. Kaempfer, R., Rosen, H., and DiSegni, G. (1984). In “Mechanisms of Protein Synthesis” (E. Bermek, ed.), p. 120. Springer-Verlag, Berlin and New York. Zucker, W. V., and Schulman, H. M. (1968). PNAS 59, 582. Rabinovitz, M., Freedman, M. L., Fisher, J. M., and Maxwell, C. R. (1969). CSHSQB 34, 567. Hunt, T., Vanderhoff, G., and London, I. M. (1972). JMB 66, 471. Ehrenfeld, E., and Hunt, T. (1971). PNAS 68, 1075. Kosower, N. S . , Vanderhoff, G. A,, and Kosower, E. M. (1972). BBA 272, 623. Farrell, P., Balkow, J., Hunt, T., Jackson, R. J., and Trachsel, H. (1977). Cell (Cambridge, Mass.) 11, 187. Legon, S., Jackson, R. J., and Hunt, T. (1973). Nature (London), New B i d . 230, 91. Dambrough, C., Legon, S., Hunt, T., and Jackson, R. J. (1973). JMB 76, 379. Pinphanichakam, P., Kramer, G., and Hardesty, B. (1976). EBRC 73, 625. Ranu, R. S., London, I. M., Das, A,, Dasgupta, A,, Majumdar, A., Ralston, R., Roy, R., and Gupta, N. K. (1978). PNAS 75, 745. de Haro, C., Datta, A,, and Ochoa, S . (1978). PNAS 75, 243. de Haro, C., and Ochoa, S. (1978). PNAS 75, 2713. Ranu, R. S . , and London, I. M. (1979). PNAS 76, 1079. de Haro, C., and Ochoa, S. (1979). PNAS 76, 1741. Das, A., Ralston, R. O., Grace, M., Roy, R., Ghosh-Dastidar, P., Das, H. K . , Yaghmai, B., Palmieri, S., and Gupta, N. K. (1979). PNAS 76, 5076. Grosfeld, H., and Ochoa, S. (1980). PNAS 77, 6526.

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23. Das, H. K., Das, A,, Gosh-Dastidas, P., Ralston, R. O., Yaghmai, B., Roy, R., and Gupta, N. K. (1981). JBC 256, 6491. 24. Levin, D. H., Ranu, R. S., Ernst, V., and London, I. M. (1976). PNAS 73, 31 12. 25. Kramer, G., Cimadevilla, M., and Hardesty, B. (1976). PNAS 73, 3078. 26. Ranu, R. S., and London, I. M. (1976). PNAS 73, 4349. 27. Gross, M., and Mendelewski, J. (1977). BBRC 74, 559. 28. Levin, D. H., and London, I. M. (1978). PNAS 75, 1121. 29. Ernst, V., Levin, D. H., and London, I. M. (1978). PNAS 75, 41 10. 30. Ernst, V., Levin, D. H., and London, I. M. (1979). PNAS 76, 21 18. 31. Cherbas, L., and London, I. M. (1976). PNAS 73, 3506. 32. Farrell, P. J., Hunt, T., and Jackson, R. J. (1978). EJB 89, 517. 33. Leroux, A,, and London, I. M. (1982). PNAS 79, 2147. 34. Menick, W. C. (1979). JBC 254, 3708. 35. Peterson, P. T., Merrick, W. C., and Safer, B. (1979). JBC 254, 2509. 36. Trachsel, H., and Staehlin, T. (1978). PNAS 75, 204. 37. Benne, R., and Hershey, J. W. B. (1978). JBC 253, 3078. 38. Clemens, M. J., Pain, V. M., Wong, S., and Henshaw, E. C. (1982). Nature (London) 296, 93. 39. Siekierka, J., Mauser, L., and Ochoa, S. (1982). PNAS 79, 2537. 40. Matts, R. L., Levin, D. H., and London, I. M. (1983). PNAS 80, 2559. 41. Panniers, R., and Henshaw, E. C. (1983). JBC 258, 7928. 42. Mehta, H. B., Woodley, C. L., and Wahba, A. J. (1983). JBC 258, 3438. 43. Walton, G. M., and Gill, G. N. (1975). BBA 390, 231. 44. Konieczny, A,, and Safer, B. (1983). JBC 258, 3402. 45. Walton, G. M., and Gill, G. N. (1976). BBA 418, 195. 46. Amesz, H., Goumans, H., Haubrich-Morree, T., Voorma, H. O., and Benne, P., (1979). EJB 98, 513. 47. Siekierka, J., Mitsui, K., and Ochoa, S. (1981). PNAS 78, 220. 48. Ralston, R. O., Das, A , , Dasgupta, A,, Roy, R., Palmieri, S., andGupta, N. K. (1978). PNAS 75, 4858. 49. Ralston, R. 0.. Das, A,, Grace, M., Das, H. K., and Gupta, N. K. (1979). PNAS 76, 5490. 50. Pain, V. M., and Clemens, M. J. (1983). Biochemistry 22, 726. 51. Safer, B. (1983). Cell (Cambridge, Mass.)33, 7. 52. Jagus, R., and Safer, B. (1981). JBC 256, 1317. 53. Safer, B., Jagus, R., Konieczny, A,, and Crouch, D. (1982). Dev. Biochem. 24, 31 I . 54. Siekierka, J., Manne, V., and Ochoa, S. (1984). PNAS 81, 352. 5 5 . Siekierka, J., Datta, A., Mauser, L., and Ochoa, S. (1982). JBC 257, 4162. 56. Siekierka, J., Manne, V., Mauser, L., and Ochoa, S. (1983). PNAS 80, 1232. 57. Salimans, M., Goumans, H., Amesz, H., Benne, R.,and Voorma, H. 0. (1984). EJB 145,91. 58. Thomas, N. S. B., Matts, R. L., Petryshyn, R., and London, I. M. (1984). PNAS 81, 6998. 59. Thomas, N. S. B., Matts, R. L., Levin, D. H., and London, I. M. (1985). JBC 260, 9860. 60. Goss, D. J., Parkhurst, L. J., Mehta, H. B., Woodley, C. L., and Wahba, A. J. (1984). JBC 259, 7374. 61. Manchester, K. L. (1985). FEBSLett. 182, 15. 62. Matts, R. L., Levin, D. H., and London, 1. M. (1986). PNAS 83, 1217. 63. Ochoa, S. (1983). ABB 223, 325. 64. Balkow, K., Hunt, T., and Jackson, R. J. (1975). BBRC 67, 366. 65. Ernst, V., Levin, D. H., Ranu, R. S., and London, 1. M. (1976). PNAS 73, 1 112. 66. Ranu, R. S. (1982). BBRC 109, 872. 67. Suzuki, H., and Mukouyama, E. B. (1985). J . Biochem. (Tokyo) 97, 1289.

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Matts, R. L., and London, I. M. (1984). JBC 259, 6708. Wong, S. T.,Mastropaolo, W., and Henshaw, E. C. (1982). JBC 257, 5231. De Benedetti, A,, and Baglioni, C. (1985). JBC 260, 3135. Gross, M., Redman, R., and Kaplansky, D. A. (1985). JBC 260, 9491. Emst, V., Levin, D. H., Leroux, A. L., and London, I. M. (1980). PNAS 77, 1286. Trachsel, H., Ranu, R. S., and London, I. M. (1978). PNAS 75, 3654. Hunt, T. (1979). Miami Winter Symp. 16, 321. Levin, D. H., Petryshyn R., and London, I. M. (1980). PNAS 77, 832. Gross, M., and Mendelewski, J. (1978). BBA 520, 650. Fagard, R., and London, I. M. (1981). PNAS 78, 866. Henderson, A. B., Miller, A. H., and Hardesty, B. (1979). PNAS 76, 2605. Levin, D. H., Emst, V., and London, I. M. (1979). JBC 254, 7935. Emst, V., Levin, D. H., Foulkes, J. G., and London, I. M. (1982). PNAS 79, 7096. Emst, V., Levin, D. H., and London, I. M. (1978). JBC 253, 7163. Michelson, A. M., Emst, V., Levin, D. H., and London, I. M. (1984). JBC 259, 8529. Hunt, T., Herbert, P., Campbell, E. A., Delidakis, C., and Jackson, R. J. (1983). EJB 131, 303. 84. Jackson, R. J., Herbert, P., Campbell, E. A,, and Hunt, T. (1983). EJB 131, 313. 85. Wallis, M. H., Kramer, G., and Hardesty, B. (1980). Biochemistry 19, 798. 86. Petryshyn, R., Levin, D. H., and London, I. M. (1980). BBRC 94, 1190. 87. Levin, D. H., Petryshyn, R., and London, I. M. (1981). JBC 256, 7638. 88. Petryshyn, R., Levin, D. H., and London, I. M. (1982). PNAS 79, 6512. 89. Foulkes, J. G., Emst, V., and Levin, D. H. (1983). JBC 258, 1439. 90. Zilberstein, A., Federman, P., Schulman, L., and Revel, M. (1976). FEBS Left. 68, 119. 91. Lebleu, B., Sen, G. C., Shaila, S., Cabrer, B., and Lengyel, P. (1976). PNAS 73, 3107. 92. Roberts, W. K., Hovanessian, A., Brown, R. E., Clemens, M. J., and Kerr, I. M. (1976). Nature (London) 264, 477. 93. Lodish, H. F., and Desalu, 0. (1973). JBC 248, 3520. 94. Beuzard, Y.,Rodvien, R., and London, I. M. (1973). PNAS 70, 1022. 95. Freedman, M. L., Gerighty, M., and Rosman, J. (1974). JBC 249, 7290. 96. Petryshyn, R., Rosa, F., Fagard, R., Levin, D., and London, I. M. (1984). BBRC 119, 891. 97. Petryshyn, R., Trachsel, H., and London, I. M. (1979). PNAS 76, 1575. 98. Mathews, M. B. (1972). BBA 272, 108. 99. Dabney, B. J., and Beaudet, A. L. (1978). JBC 253, 7124. 100. Freedman, M. L., and Karpatkin, S. (1973). BBRC 54, 475. 101. Weber, L. A., Feman, E. R., and Baglioni, C. (1975). Biochemistry 14, 5315. 102. Pain, V. M., and Henshaw, E. C. (1975). EJB 57, 335. 103. Pain, V. M., Lewis, J. A,, Huvos, P., Henshaw, E. C., and Clemens, M. J. (1980). JBC 255, 1486. 104. Clemens, M. J., Pain, V. M., Henshaw, E. C., and London, I. M. (1976). BBRC 72, 768. 105. Ahern, T., Sampson, J., and Kay, J. E. (1974). Nature (London) 248, 519. 106. Kay, J. E., Benzie, C. R., Dicker, P., and Lindahl-Kiessling, K. (1978). FEBSLert. 91, 40. 107. Liang, T., and Liao, S. (1975). PNAS 72, 706. 108. Liang, T., Castaneda, E., and Liao, S. (1977). JBC 252, 5692. 109. Ranu, R. S. (1980). BBRC 97, 1124. 110. Ranu, R. S. (1980). FEBSLett. 112, 211. 111. Delaunay, J., Ranu, R. S., Levin, D. H., Emst, V., and London, I. M. (1977). PNAS 74, 2264. 112. Fagard, R., and Guguen-Guillouzo, C. (1983). BBRC 114, 612. 113. Horlein, D., McPherson, J., Goh, S. H., and Bomstein, P. (1981). PNAS 78, 6163. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83.

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114. McPheison, J. M., Horlein, D., Abbott-Brown, D., and Bornstein, P. (1982). JBC257,8557. 115. Hunter, T., Hunt, T., Jackson, R. J., and Robertson, H. D. (1975). JBC 250, 409. 116. Zilberstein, A., Kimchi, A , , Schmidt, A,, and Revel, M. (1978). PNAS 75, 4734. 117. Stewart, W. E., I1 (1979). In “The Interferon System,” p. 421. Springer-Verlag, Berlin and New York. 118. Lengyel, P. (1982). Annu. Rev. Biochem. 51, 251. 119. Baglioni, C. (1979). Cell (Cambridge, Muss.) 17, 255. 120. Kerr, 1. M., and Brown, R. E. (1978). PNAS 75, 256. 121. Clemens, M .J., and Williams, B. R. G. (1978). Cell (Cambridge, Muss.) 13, 565. 122. Nilsen, T. W., Maroney, P. A,, and Baglioni, C. (1983). Mol. Cell. Biol. 3, 64. 123. Samuel, C. E., Duncan, R., Knitson, G. S., and Hershey, J. W. B. (1984). JBC 259, 13451. 124. Gupta, S. L., Holmes, S. L., and Mehra, L. (1982). Virology 120, 495. 125. Keay, S., and Grossberg, S . E. (1980). PNAS 77, 4099. 126. Lieberman, D., Voloch, Z . , Aviv, H., Nudel, U., and Revel, M. (1974). Mol. Biol. Rep. 1, 447. 127. Rossi, G. B., Dolei, A,, Cioe, L., Bernadetto, A , , Matarese, G. P., andBelardetti, F. (1977). PNAS 74, 2036. 128. Revel, M., Kimchi, A,, Shulman, L., Wolf, D., Merlin, G., Schmidt, A,, Friedrnan, M., Lapidot, Y., and Rapport, S. (1981). In ‘‘Cellular Responses to Molecular Modulators” (L. W. Mozes, J. Schultz, R. Werner, and W. A. Scott, eds.), p. 361. Academic Press, New York. 129. Stark, G. R., Dower, W. J., Schimke, R. T., Brown, R. E., and Kerr, I. M. (1979). Nature (London) 278, 47 1. 130. Krishnan, I., and Baglioni, C. (1980). PNAS 77, 6506. 131. Mechi, N., Affabris, E., Romea, G., Lebleu, B., and Rossi, G. B. (1984). JBC 259, 3261. 132. Petryshyn, R., Chen, J.-J., and London, I. M. (1984). JBC 259, 14736. 133. Green, H., and Kehinde, 0. (1974). Cell (Cambridge. Muss.) 1, 113. 134. Kuri-Harcuch, W., and Marsch-Moreno, M. (1983). J. Cell. Physiol. 114, 39. 135. Reichel, P. A , , Merrick, W. C., Siekierka, J., and Mathews, M. B. (1985). Nurure (London) 313, 196. 136. Siekierka, J . , Mariano, T. M . , Reichel, P. A,, and Mathews, M. B. (1985). PNAS 82, 1959. 137. Morrow, C. D., Gibbons, G. F., and Dasgupta, A. (1985). Cell (Cambridge, Muss.) 40,913. 138. Manning, D. R., and Gilman, A. G. (1983). JBC 258, 7059. 139. Cassel, D., and Pfeuffer, T. (1978). PNAS 75, 2669. 140. Katada, T., and Ui, M. (1982). JEC 259, 7210. 141. Van Dop, C., Yamanaka, G., Steinberg, F., Sekura, R. D., Manclark, C . R., Stryer, L., and Bourne, H. R. (1984). JBC 259, 23. 142. Abood, M. E., Hurley, J. B., Pappone, M.-C., Bourne, H. R., and Stryer, L. (1982). JBC 257, 10540. 143. Hurley, J. B., Simon, M. I., Teplow, D. B., Robishaw, J. D., and Gilman, A. G. (1984). Science 226, 860. 144. Taparowsky, E., Shimizu, K., Goldfarb, M., and Wigler, M. (1983). Ce/l(Cumbridge.Muss.) 34, 581. 145. Yatsunami, K., and Khorana, H. G. (1985). PNAS 82, 4316. 146. Medynski, D. C., Sullivan, K., Smith, D., Van Dop, C., Chang F.-H., Fung, B. K.-K., Seeburg, P.H., and Bourne, H. R. (1985). PNAS 82, 431 1. 147. Moss, J., and Vaughan, M. (1978). PNAS 75, 3621. 148. Sitikov, A. S., Davydova, E. K., and Ovchinnikov, L. P. (1984). FEBS Lett. 176, 261.

Regulation of Corctractile Activity JAMES R. SELLERS ROBERT S. ADELSTEIN Laboratory of Molecular Cardiology National Heart, Lung, and Blood Institute Beihesda, Maryland 20892

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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111.

IV. V.

VI.

Phosphorylation . . . . . . . . A. Effect of Phosphorylati .......... Activity of Myosin . . B. The Effect of Light Ch Assembly and on the Conformation of Smooth-Muscle Myosin . . . . . . . C. Myosin Phosphorylation in Smooth Muscle . . . . D. Phosphorylation of Myosin Kinases . . . . . . . . . . . . . . . Role of Phosphorylation in Mo Striated Muscle Proteins . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . . . . . . . . . . . . . A. Phosphorylation of Myosin ........... B. Phosphorylation of Troponi Phosphorylation-Dependent Regulatory Systems in Invertebrate Muscles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of Cytoplasmic Myosins . . . . . . . . . . . . . . A. Lower Eukaryotes . . . . . . . . . . . . . . . . . . . . B. Phosphorylation of Vertebrate Cytoplasmic C. Phosphorylation of Myosin by Protein Kinase C Summary . ......................................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , . , . . . . . , . . . 38 I THE ENZYMES.

Vol. XVIII

386 386 393 396 399 404 404 405 406 406 406 407 409 412 413

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JAMES R. SELLERS AND ROBERT S. ADELSTEIN

Introduction

All muscles contain filaments of actin and myosin that are required for contractile activity. Myosin and actin are not unique to well-differentiated muscle tissue but are components of virtually every eukaryotic cell where they may participate in a variety of contractile functions such as cytokinesis, phagocytosis, amoeboid motion, and perhaps organelle translocation (1-3). Actin filaments are extended polymers of monomeric actin, a globular protein with a molecular weight of about 42,000. Because of their relatively thin diameter, as compared to filaments of myosin, actin filaments are commonly referred to as “thin” filaments. A second protein usually associated with the thin filament is tropomyosin (M,= 68,000) which is present in a 1:7 molar ratio with actin. As Fig. 1 illustrates, thin filaments consist of a double helix of polymerized actin with end-to-end polymers of tropomyosin lying in the groove. In some muscle cells such as vertebrate skeletal, and cardiac muscle, a regulatory protein called troponin is also present in a stoichiometry of 1: 1 with tropomyosin (Fig. 1, blow up). In all muscle systems, actin filaments are anchored at one end. In striated muscles the anchor is a well-demarcated structure called the Z-line. In smooth-muscle and nonmuscle cells, the anchoring structures are less well defined. In smooth muscles they are thought to be the dense bodies, whereas in nonmuscle cells actin filaments can be anchored on the cell membrane as well as other intracellular organelles. The thick filaments, which are easily visualized in cardiac and skeletal muscle, are composed mainly of myosin (Fig. 1). The myosin molecule is a hexamer composed of two heavy chains (M,= 200,000) and two pairs of light chains (M,= 15,000-30,000) (Fig. 2). The carboxyterminal portion of the heavy chains forms a coiled-coil, alpha-helical, rodlike region of approximately 150 nm in length, which diverges near the aminoterminal ends of the two heavy chains to form two globular “head” regions, each of which is associated with one of each type of light chain. The MgATPbinding site and the actin-binding site are located in the globular region. In vitro at physiological ionic strengths or lower, myosin self-associates via the rod region to form bipolar thick filaments similar to those found in muscle (Fig. 1). The heads project away from the surface of the filament backbone. Contraction occurs when the thin filaments are pulled toward the center of the thick filaments by the ATP-dependent cyclical interaction of the myosin heads with the actin subunits (Fig. 1). In all muscles, contraction is initiated by an elevation of the intracellular concentration of free Ca2+. However, the site of action of Ca2+ varies from muscle to muscle. Three types of Ca2 -dependent regulatory systems have been well described in muscle: ( a ) actin-linked; (b) myosin-linked; and ( c ) phosphorylation-dependent myosin-linked regulation. Actin-linked systems require the presence of troponin which is composed of three polypeptide subunits: troponin-T (TN-T), troponin-1 (TN-I), and troponin-C (TN-C) (Fig. 1) [for a review, see Ref. ( 4 ) ] . +

Tropomyosin

I

\ \.

U

Thin Filament

A7tin

, I C T, Troponin Complex

/

/'

' Thick Filament

2

2 Line

Line

U

U

FIG. 1. Diagrammatic representation of myosin (thick) and actin (thin) filaments, in the relaxed and contracted state. The blow up shows a diagram of the actin filament together with tropomyosin and troponin. This latter complex is found in striated muscle (see text for details).

3 84

JAMES R. SELLERS AND ROBERT S. ADELSTEIN TAIL

HEAD N

20 kDa LC 17 kDa LC C C

FIG. 2. Diagrammatic representation of a myosin molecule showing the six polypeptide chains and the globular and helical domains.

TN-T is a tropomyosin-binding subunit, TN-I is an inhibitory subunit, and TN-C is a calcium-binding subunit which, upon binding of Ca2+, relieves the inhibition imposed by TN-I. This type of regulatory system is found in vertebrate striated muscle ( 4 ) and in some invertebrate muscle (5). Myosin-linked regulation, which operates via direct calcium binding to the myosin molecule, has been best characterized in molluscan muscles (6), where the myosin is inactive in the absence of bound calcium and becomes active when calcium is bound (6, 7). This regulation is mediated in part by one class of the light chains termed “regulatory” which are bound to each head (8, 9). Regulatory light chains also play a prominent role in phosphorylation-dependent myosin-linked systems. Here, however, the site of Ca2+ action is calmodulin, the ubiquitous calcium-binding protein, which binds to and activates myosin light chain kinase (2, 10). Myosin light chain kinase specifically phosphorylates the regulatory light chain of the myosin, thereby converting myosin to an active form which interacts with actin resulting in muscle contraction (2, 10). The myosin is returned to an inactive state by dephosphorylation. This type of regulation has been described for vertebrate smooth muscle (2, ZO), and for the striated muscle of Limulus, the horseshoe crab (IZ). Many muscles, for example Limulus, are dually regulated. In addition to the phosphorylation-dependent myosin-linked regulation previously described, Limulus also shows a functional actin-linked system with a well-characterized troponin system ( 1 2 ) .Calcium may also exert influences on contractile systems by binding to proteins other than troponin and calmodulin as discussed in Section ILC, 1. The Ca2 -dependent regulation of muscles can be studied at several different levels. Intact fibers and permeabilized “skinned” fibers can generate tension in a reversible manner and thus permit study of a fully intact system in which the contractile machinery is relatively undisturbed. In many systems myofibrils, which possess the basic sarcomeric structure, can be prepared for in vitro studies. +

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These have the advantage of being more amenable to biochemical manipulation and measurement than skinned fiber preparations. A further simplification involves the preparation of the various proteins in their purified form. For instance, actin and myosin from vertebrate skeletal muscle can be purified to near homogeneity. Myosin alone exhibits a MgATPase activity which can be activated by addition of pure actin. This ability of actin to activate the MgATPase activity of myosin is taken as an in vitro correlate of muscle contraction. A further observation can be made using the in vitro system; the activation of the MgATPase activity of vertebrate skeletal myosin by pure actin is not dependent on the presence of Ca2 ions. This system becomes Ca2 -dependent by the addition of the thin-filament regulatory proteins, troponin and tropomyosin. Thus, a Ca2 dependent regulatory system can be reconstituted from purified proteins. One marked disadvantage in working with myosin is that it forms filaments at physiological ionic strength that preclude the use of many spectroscopic techniques. In addition, actomyosin systems do not obey simple Michaelis-Menten kinetics, presumably due to the filamentous state. Fortunately, two different soluble subfragments which are enzymically active can be prepared by controlled proteolytic digestion of myosin using a variety of proteases. Subfragment-one ( S 1) is a single-headed subfragment consisting of one myosin head, and heavy meromyosin (HMM) is a two-headed subfragment containing the two heads of the parent myosin held together by the subfragment-two (S2) region (Fig. 3). Using these subfragments, the kinetics of ATP hydrolysis have been studied and a better understanding of the interaction of myosin with actin has been achieved (2, 13, 14). In this chapter we discuss how phosphorylation mediates or modulates the +

+

+

HMM LMM

Chyrnotrypsin

k

Papain

Rod

+

.+. s1

s1

FIG.3. Diagrammatic representation of the various subfragments of myosin. The S2 region is that part of the rod remaining as part of HMM.

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JAMES R. SELLERS AND ROBERT S. ADELSTEIN

interaction of actin and myosin in both muscle and nonmuscle systems. Although the major emphasis is on the phosphorylation of vertebrate smooth-muscle myosin, we also discuss phosphorylation of contractile proteins in other muscles as well as nonmuscle cells.

II. Regulation of Vertebrate Smooth-Muscle Myosin by Phosphorylation OF THE ACTIN-ACTIVATED A, EFFECTOF PHOSPHORYLATION MGATPASEACTIVITYOF MYOSIN.

1.

Regulation of the Actin-Activated MgATPase Activity of Smooth-Muscle Myosin by Reversible Phosphorylation

Myosin has been isolated from a variety of vertebrate smooth-muscle tissues of both avian and mammalian origins. In all cases the molecular weights of the subunits are remarkably similar with heavy chains of about 200,000 daltons and light chains of 20,000 and 17,000 daltons. The 20,000 dalton light chain can be phosphorylated by the calcium-calmodulin-dependent enzyme, myosin light chain kinase (see Volume XVII, Chapter 4).The site of phosphorylation in the 20,000-dalton light chain of turkey gizzard myosin is located near the amino terminus of the light chain at serine-19 (15). Since there is a 20,000-dalton regulatory light chain associated with each of the two heads of myosin, a total of 2 mol of phosphate can be incorporated per mol of myosin under usual conditions. With prolonged incubation time and high concentrations of myosin light chain kinase, a second mole of phosphate can be incorporated into each light chain (16-17a) and this is accompanied by a further 2- to 3-fold increase in the actin-activated MgATPase activity ( 1 7 , 17a). Phosphatases have been isolated from smooth muscles that dephosphorylate myosin (18-21) although it is not clear whether there is a specific myosin light chain phosphatase or how its activity is regulated. A phosphatase that binds strongly to myosin has been isolated from turkey gizzards (22). It is generally accepted that reversible phosphorylation regulates the actinactivated MgATPase activity of smooth-muscle myosin in vitro (23-27). An example of this is seen in Table I which shows the data from an experiment performed by mixing purified, well-characterized proteins to form a reversible regulatory system (27). Pure, unregulated actin used in this experiment was isolated from rabbit skeletal muscle although similar results were obtained with smooth-muscle actin. It can be seen that phosphorylation of myosin purified from turkey gizzards and human platelets is accompanied by a dramatic increase in the actin-activated MgATPase activity and that this activity can be returned to the original low basal

387

13. REGULATION OF CONTRACTILE ACTIVITY TABLE I CORRELATION OF THE ACTIN-ACTIVATED MGATPASEACTIVITY WITH PHOSPHATE INCORPORATION" Mg ATPase

Smooth-muscle myosin Unphosphorylated Phosphorylated Dephosphorylated Rephosphorylated Smooth-muscle HMM Unphosphory lated Phosphorylated Dephosphory lated Rephosphory lated Plaielet myosin Unphosphorylated Phosphorylated Dephosphory lated

K + -EDTA Minus actin ATPase (nmol/min/mg)

Phosphate incorporated (mol Pi/mol myosin)

Plus actin

0 I .9

<2 <2

250 250

2.0

4 51 5 46

0 1.9 0.1

10

<2 <2

1030

351 20

2.1

37 1

0.1

0

1.9 0.1

930

5 89 5

uData are from Ref. (27).

rate by dephosphorylation. Phosphorylation does not have a dramatic effect on the MgATPase activity in the absence of actin or on the nonphysiological highsalt ATPase activities measured in the presence of K+-EDTA and 0.5 M KCl. A similar experiment can be performed with HMM, prepared by chymotryptic digestion of phosphorylated myosin. It was noted that phosphorylation protects the regulatory light chains from chymotryptic digestion and allows isolation of a well-regulated heavy meromyosin (27). Again, Table I shows the same correlation between phosphorylation and high actin-activated MgATPase activities, and this correlation is maintained during dephosphorylation and rephosphorylation. The fact that HMM is regulated is of considerable importance since, as discussed in Section 11, phosphorylation also affects the filament assembly properties of smooth-muscle and nonmuscle myosin (28-30). 2. Light Chain Phosphorylation As a Derepressor What is the nature of the phosphorylation-dependentregulation? How does the light chain function? To properly answer these questions one would like to be able to reversibly dissociate the light chain from smooth-muscle myosin and

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JAMES R. SELLERS AND ROBERT S. ADELSTEIN

study the enzymic properties of myosin with and without bound regulatory light chains. In practice, this has not been done with smooth-muscle myosin, but it has been accomplished with scallop myosin which is regulated by a light chain that mediates calcium binding (7-9, 31). A number of studies by Szent-Gyorgyi and his colleagues have shown that the light chain acts as a repressor of the actinactivated MgATPase activity and that calcium-binding to the myosin at a site formed by the light chain and heavy chain overcomes this repression (8, 9, 31). Treatment of scallop myosin with EDTA removes the regulatory light chains and also removes the inhibition of the actin-activated MgATPase activity, giving rise to a high actin-activated MgATPase activity both in the presence and absence of calcium. Full calcium regulation can be restored by readdition of the light chains (31). That the regulatory light chain of smooth-muscle myosin may also act as a repressor is suggested by finding that brief trypsin treatment of aortic myosin, which preferentially digests the 20,000-dalton light chain, results in a partial loss of regulation. This loss was manifested by an increase in the actin-activated MgATPase activity of myosin (32). However, in this experiment readdition of smooth-muscle light chain was not attempted and thus there is no information as to whether the loss of regulation was reversible. 3 . Effect of Calcium on the MgATPase Activity of Smooth-Muscle Myosin

From the earliest reports that phosphorylation regulated the MgATPase activity of smooth-muscle myosin, there have been conflicting views as to whether calcium-binding by myosin also modulated the MgATPase activity (24, 25, 33). Chacko and his collaborators have shown that the actin-activated MgATPase of phosphorylated myosin from vas deferens (24) and pulmonary arteries (33) is further increased about twofold in the presence of calcium ions. The Ca2+sensitivity observed with the arterial myosin was correlated with Ca2 -binding to myosin (33). Further examination reveals that the Ca2 -sensitivity occurred over a rather narrow range of Mg2+ concentration (between 1-3 mM free Mg2+) (33). This latter point is important since many of the studies using myosin from other smooth muscles which failed to observe the Ca2 -dependency were conducted at higher Mg2+ ion concentration (3-8 mM free Mg2+) (25). The calcium-dependency of phosphorylated gizzard smooth-muscle myosin has been reexamined at varying Mg2 concentrations (34-36). Surprisingly, under some conditions (free Mg2+ concentration less than 1 mM using gizzard actin and tropomyosin), the actin-activated MgATPase activity of phosphorylated gizzard myosin was almost totally dependent upon the presence of calcium (3436). This calcium-sensitivity was also lost upon increasing the Mg2 concentration. Calcium ions activated HMM to a lesser degree than intact myosin suggesting that Ca2+ may also be affecting myosin filament formation (37). +

+

+

+

+

389

13. REGULATION OF CONTRACTILE ACTIVITY

In summary, with all vertebrate smooth muscles, myosin phosphorylation appears to be essential for actin-activation of the MgATPase activity. Under certain conditions however, usually at low concentrations of free Mg2 , the activity of phosphorylated myosin is further increased by addition of calcium. The exact Mg2 concentrations at which the calcium-sensitivity is observed may depend on the source of myosin. Thus, the physiological relevance of these in vitro findings depends upon the intracellular free Mg2 concentration in smooth muscles. One report suggests that the free Mg2+ concentration in smooth muscles is rather low (in the mM range), and thus that Ca2+ may indeed play a modulating role by binding to myosin (38). +

+

+

4.

Cooperativity between the Two Heads of Myosin

The fact that a myosin molecule has two active sites, each located on a discrete head, raises the question as to whether the two heads can interact. There are two points to be considered: First, what are the kinetics of phosphorylation of the two heads? Are they phosphorylated randomly with equal rates or is there order or cooperativity in the kinetics, such that the two heads are phosphorylated at different rates? Second, is each head activated independently upon phosphorylation or is it necessary that both heads be phosphorylated before activation of either head is possible? One way to study both of these points is to examine the correlation between the actin-activated MgATPase activity and the extent of phosphorylation. This approach was developed by Chantler et al. (39) for the actin-activation of scallop myosin by calcium and has been extended to the case of smooth muscle by Persechini and Hartshorne (40)and other investigators (414 4 ) . For a detailed analysis of the method the reader is referred to Sellers et al. (42). The correlation between phosphorylation of gizzard smooth-muscle myosin (in filamentous form) and the MgATPase activity is nonlinear such that little activation of the MgATPase is seen in the presence of actin until about 50% of maximal phosphorylation is obtained (41-44). This is diagnostic of a case where both heads of a molecule must be phosphorylated before the MgATPase of either head can be activated by actin and where the phosphorylation of the two heads is ordered such that at 50% phosphorylation most of the myosin is monophosphorylated. With the soluble subfragment, HMM, a hyperbolic correlation curve is obtained which indicates that, like myosin, both heads of HMM must be phosphorylated for activation of either head (41, 4 2 ) . However, with HMM the phosphorylation of the two heads appears to occur randomly and at equal rates (41, 42). That the two heads of myosin in filaments are phosphorylated in an ordered manner while the two heads of HMM are phosphorylated randomly has been confirmed by direct examination of the kinetics of phosphorylation of the two

390

JAMES R. SELLERS AND ROBERT S. ADELSTEIN

molecules (42). Sellers et al. (42) showed that this difference in phosphorylation patterns was due to polymerization of myosin into filaments by examining phosphorylation kinetics of myosin in 0.6 M KCl where it exists in a soluble twoheaded monomeric form. Here the two heads phosphorylate randomly with equal rates indicating that the two heads of myosin are inherently indistinguishable by this criterion. Polymerization of the myosin into filaments affects the heads such that they now can be phosphorylated at different rates by myosin light chain kinase. This ordered phosphorylation could either be due to negative cooperative interactions between the two heads or simply to the fact that the two heads are in asymmetric environments and are being phosphorylated randomly but with markedly unequal rates. It is not possible to distinguish between these two mechanisms kinetically. It is interesting in this regard that Ikebe and Ogihara (45)found that phosphorylated and unphosphorylated myosin filaments had different appearances in the electron microscope. The unphosphorylated filaments showed a periodic banding pattern as if the myosin heads were being held in a more highly ordered state. Some preparations of myosin filaments phosphorylate randomly [Refs. (45a, 45b),and J. R. Sellers, unpublished data] indicating that kinetics of phosphorylation may depend upon how the filaments were formed. Is the requirement that both heads be phosphorylated for activation a common property of all smooth-muscle myosins? Chacko and co-workers found a linear coorelation between phosphorylation and MgATPase activity in myosins from bovine stomach and porcine pulmonary arteries (33, 46). Their experimental approach was somewhat different from the reports previously mentioned and the assays for actin-activation of myosin MgATPase activity were conducted in the presence of tropomyosin. Ikebe et al. (41) reported that tropomyosin had no effect on the shape of the gizzard myosin correlation curve, but Merkel et al. (43) reported that tropomyosin did result in a more linear curve. It is also interesting to note that Wagner et al. (47) find a linear relationship between the MgATPase activity and phosphorylation of nonmuscle myosin from thymus, even in the absence of tropomyosin. A linear correlation has ambiguous interpretations. It could arise either because the two heads are activating independently upon phosporylation, or because the phosphorylation is occurring in a positively cooperative fashion (i.e., the phosphorylation of one head increases the rate of phosphorylation of the second head). Therefore, one cannot determine whether interactions between the two heads are required for activation in all myosins. It is interesting, however, to speculate on the mechanism of the apparent cooperative head-head interactions observed with gizzard myosin. The phosphorylation site on the regulatory light chain is at serine- 19 which is close to the amino terminus (15).Results from cross-linking studies (48,49)and from immunoelectron microscopic localization of the light chains (50, 51) from other myosins, indicate that the regulatory light chains are located toward the proximal or

39 1

13. REGULATION OF CONTRACTILE ACTIVITY

“neck” region of S l and may, in fact, overlap into the S2 region. This is in agreement with the observation that regulatory light chains can rebind to a 26,000-dalton carboxy-terminal tryptic peptide of smooth-muscle S I which would be contiguous with S2 in the intact molecule (52). Therefore, all six polypeptides of myosin are in close proximity in this neck region, and it is likely that this is the essential area for the observed cooperativity. This may explain why S1 is not regulated, even if it is prepared such that it retains the regulatory light chain (53),whereas single-headed myosin prepared by mild papain digestion is regulated (54). Similar observations have been made with S1, HMM, and single-headed myosin from the scallop (55). 5 . Which Kinetic Step Is Regulated by Phosphorylation?

The extent of regulation by phosphorylation can be estimated by comparing the V,,, of the actin-activated MgATPase activity of phosphorylated HMM (1.8 s- I ) and unphosphorylated HMM (0.075 s- I ) (56). The actual extent of regulation may be larger than this since it is possible that some of the unphosphorylated HMM is damaged and behaves kinetically like phosphorylated HMM. It is likely that phosphorylation increases the actin-activated MgATPase activity of myosin by changing a single, or perhaps a few, kinetic rate constants that occur as intermediates in the MgATPase cycle. To examine this, it is necessary to draw a simple kinetic scheme for the hydrolysis of MgATP by actomyosin (Scheme I). The basis for this scheme has its foundation in a large number of studies made primarily on the skeletal-muscle actomyosin systems, which have been extensively reviewed (13, 14). Taylor and his colleagues have more closely examined the kinetics of smooth-muscle acto-S 1 and confirmed the general applicability of such a scheme (57-59). I

7

5

-

9

very slow

M

M.ATP*.

M.ADP.Pi

M.ADP

M

SCHEME I

Briefly, the pathway for hydrolysis of ATP can be described as follows: In the absence of nucleotide all of the actin and myosin will be complexed as actomyosin (AM) due to the extremely tight binding constant present under these conditions (60). Upon binding of ATP, actomyosin can (a) hydrolyze ATP directly; or (b) dissociate to form actin and myosin (M) which then hydrolyzes ATP. The degree of dissociation depends on the ionic strength and the actin

392

JAMES R. SELLERS AND ROBERT S. ADELSTEIN TABLE I1 OF

ACTIN-BINDING AND MGATPASEACTIVITY UNPHOSPHORYLATED A N D PHOSPHORYLATED HMM AT 300 pA4 ACT IN^ MgATPase activity

HMM

Fraction bound

Unphosphorylated Phosphorylated

0.6 0.9

(s-1)

0.05 1.75

aData are from Ref. (56).

concentration (61). Since k7 is known to be very slow, any myosin that dissociates will recombine with actin (step 11) and will lose its products in a sequential manner in steps 8 and 10, yielding the initial actomyosin complex. The actual pathway is likely to be more complex than the simple scheme shown and, in particular, there may be another step between AM-ADP-Piand AM.ADP as has been shown to exist in the kinetic cycle of rabbit skeletal muscle (13). There may also be more steps involved in the binding of ATP prior to hydrolysis (58, 59). Although it is not known which of these steps is regulated by phosphorylation, the available data do allow one to rule out several plausible models; for instance, one which proposes that the unphosphorylated light chain either sterically or allosterically blocks the actin-binding sites on myosin. The model can be directly tested by measuring the binding of phosphorylated and unphosphorylated HMM to actin in the presence of ATP. Such measurements reveal only a fourfold difference in the binding constants (56). This difference is not sufficient to account for the extent of regulation, as can be seen in Table 11. At 300 pkf actin the majority of both phosphorylated and unphosphorylated HMM is bound to actin yet the actin-activated MgATPase activity of unphosphorylated HMM is 35-fold less than that of the phosphorylated HMM. Therefore, steps 4 and 11 (Scheme I) cannot be the regulated steps. Ikebe et al. found a somewhat greater difference between the binding constants of phosphorylated and unphosphorylated HMM (62). It is also unlikely that the actual hydrolysis of ATP (i.e., steps 5 and 6) is the major point of regulation. The rate of ATP hydrolysis for both phosphorylated and unphosphorylated HMM alone is about 100 s - which is greater than 1000fold faster than the measured V,,, for the actin-activated MgATPase activity of unphosphorylated HMM (56). While the binding of actin to S1 has been shown to lower the rate constant for this step, it is unlikely that it could have such a large effect (57, 58). That steps 5 and 6 are not regulated implies that steps 1 and 2 (i.e., the binding of ATP to AM) are also not regulated. The rate of ADP release from acto-S1 (step 10) has been shown to be fast (15 s - l ) and, thus is not likely to be the major regulated step (57). Therefore, by

393

13. REGULATION OF CONTRACTILE ACTIVITY

elimination it appears that the most likely step at which regulation occurs is the release of Pi from AMeADP-P, or some step (not shown in Scheme I) that occurs between the hydrolysis and the release of Pi. Experimental results provide evidence for this. Using a transient kinetic technique (63) it was shown that the release of Pi from unphosphorylated smooth-muscle HMM alone (step 7) was 0.002 s - l ) . Similarly, it was determined indirectly from a very slow ( k turbidimetric assay that the release of Pi from unphosphorylated HMM in the presence of actin (step 8) was also very slow ( k 0.002 s- I ) and that this rate constant was not increased by increasing the actin concentration (63).Therefore, phosphorylation appears to activate the MgATPase activity of avian smoothmuscle HMM by specifically increasing the rate of Pi release from AM-ADP-Pi (step 8) by a factor of about 1000. In contrast, Wagner and Vu reported that phosphorylation regulates aortic myosin by increasing the binding of actin to myosin while having only a small effect on the V,,, of the actin-activated MgATPase activity (63a).

-

-

B. THEEFFECTOF LIGHTCHAINPHOSPHORYLATION ON THICK FILAMENT ASSEMBLY AND ON THE CONFORMATION OF SMOOTH-MUSCLE MYOSIN It was shown by Suzuki et al. (28) that phosphorylation of the regulatory light chain affected the assembly of smooth-muscle myosin. If filaments are formed by dialysis from high ionic strength to a solution of 150 mM KCl, both phosphorylated and unphosphorylated myosin form thick filaments. Addition of MgATP to these filaments reveals a major difference, however, between the two. MgATP dissociates filaments of unphosphorylated myosin but not those of phosphorylated myosin. Furthermore, the amount of ATP required for this dissociation is small, approximately stoichiometric with the myosin heads. Phosphorylated myosin filaments remain mostly assembled even in millimolar concentrations of MgATP. Similar observations were also made for myosin from nonmuscle sources (29). Examination by analytical ultracentrifugation of MgATP dissociated, unphosphorylated, smooth-muscle myosin reveals that it has a sedimentation coefficient of about 10 S instead of 6 S which is characteristic of myosin monomers in 0.5 M KCl (28, 30). At first, it was proposed that this represents a myosin dimer (28),but later work using light-scattering and sedimentation-equilibrium ultracentrifugation showed that both 6 S myosin and 10 S myosin had the same molecular weight of approximately 500,000 (30, 64). Electron microscopic examination of single-rotary shadowed myosin molecules under the dissociating conditions shows that the myosin has adopted a folded “hairpin-like” structure in which the tail folds back between the heads (see Fig. 4) (65-67). This reduction in length is consistent with the 10 S sedimentation coefficient. There are two very flexible regions in the myosin rod portion that permit this unusual

394

JAMES R. SELLERS AND ROBERT S. ADELSTEIN

FIG.4. Electron micrographs of rotary shadowed myosin molecules illustrating the 10 S (A) and 6 S (B) conformation [adapted from Ref. (65)].

13. REGULATION OF CONTRACTILE ACTIVITY

395

folding to occur. They appear to be located about 50 nm and 100 nm from the head-tail junction (66). Subsequent studies by a number of laboratories have shown that the effects of phosphorylation on the conformation and polymerization of smooth-muscle myosin is complex. Phosphorylated myosin can also adapt the 10 S conformation and unphosphorylated myosin can assume a 6 S conformation depending on the ionic conditions (65, 68). At intermediate ionic strengths (0.2-0.3 M KCl) smooth-muscle myosin exists in a 6 S-10 S equilibrium where phosphorylation favors the 6 S state. The presence of folded antiparallel myosin dimers has also been observed both directly by electron microscopy and hydrodynamically (65, 69). These structures may be an intermediate in the assembly of small oligomers and thick filaments. Therefore, it appears that, in vitro, smooth-muscle myosin exists in a complex equilibrium between thick filaments, short oligomers, 10 S myosin monomers, 6 S myosin monomers, and dimers which are affected by many variables including the state of phosphorylation, pH, ionic strength, choice of cation, and myosin concentration (70). Although HMM cannot form a folded structure it also undergoes a smaller change in sedimentation coefficient when phosphorylated (71). Ikebe, Hartshorne, and co-workers have proposed that the 6 S conformation of myosin is the active state whereas the 10 S conformation is the inactive state and that phosphorylation of the myosin per se is not necessarily sufficient for activation since under appropriate conditions one can have 10 S phosphorylated myosin, which they term inactive (53,68). These ideas are based upon skinned fiber experiments, changes in the MgATPase activity of myosin in the absence of actin and changes in the K+-EDTA ATPase activity (68, 72, 72a). The fiber work showed that tension could be generated without phosphorylation if the MgCl, concentration was greatly increased (i.e., to 6-20 mM) (72a). They showed that unphosphorylated myosin under these conditions sedimented as a 6 S species. However, it is unlikely that these relatively large shifts in pH, ionic strength, or MgCl, concentrations occur in living muscle. Whether phosphorylation-dependent regulation of thick filament assembly is of physiological relevance is not certain. Somlyo et al. (73) have presented data that demonstrate the presence of thick filaments in rapidly frozen resting vascular smooth muscle where the myosin is largely unphosphorylated, but Cande et al. (74) in experiments with glycerinated gizzard muscle preparations have shown that some disassembly of the filaments can occur under relaxing conditions. In nonmuscle cells it has been difficult to visualize myosin filaments under any conditions; however, the immunofluorescent staining of cells using antimyosin antibodies shows different patterns at different stages of the cell cycle (75). Thus, it has been suggested that the MgATP-induced disassembly of unphosphorylated myosin filaments might facilitate their transport to an intracellular site where thick filaments are required for a contractile event.

396

JAMES R. SELLERS AND ROBERT S. ADELSTEIN

While myosin filament assembly that is regulated by phosphorylation may not occur in vivo in smooth muscle, it is possible that an analogous change is occurring in the conformation of the head and that this affects the properties of the filament. In this regard, it is interesting that Ikebe and Ogihara (45) visualized two types of filaments in high Mg2 concentrations and ATP, depending upon whether the myosin was phosphorylated. Phosphorylated myosin filaments were typically “fuzzy” in appearance with no regular structure. Conversely, dephosphorylated myosin filaments showed a cross-striation pattern with a 13.4 nm repeat. They suggest that this repeat arises from a regular array of myosin heads oriented parallel to the backbone. This structure may serve to lock down the myosin heads and, perhaps, reduce their ability to interact with actin, thereby contributing to a turned-off state. Cross and Sobieszek have examined the kinetics of phosphorylation of both 6 S and 10 S myosin and find that the 10 S species is a poor substrate for myosin light chain kinase and imply that, therefore, the 10 S conformation is unlikely to represent a “relaxed” state of myosin in vivo (752). +

C. MYOSINPHOSPHORYLATION IN SMOOTH MUSCLE

1. Studies with Intact and Skinned Muscle Fibers There is ample evidence from a number of smooth-muscle tissues that myosin phosphorylation is required for muscle contraction [see Kamm and Stull for a review, Ref. (76)]. Resting muscle typically contains a low level of phosphorylation which rises, upon stimulation of the muscle to a maximal value of 0.5 to 0.8 mol P,/mol 20-kDa light chain, depending upon the type of stimulation, extracellular calcium levels, and tissue type (77-83). This is accompanied by a rise in tension. Similar experiments can be carried out with skinned smooth-muscle preparations with the same result; increasing the Ca2 concentration results in phosphorylation of the myosin light chains and an increase in tension (84-89). The phosphorylation of myosin by myosin light chain kinase is both necessary and sufficient for contractions in such skinned fiber preparations as has been shown by a number of methods: 1. Calmodulin antagonists such as trifluoperazine block contraction and myosin phosphorylation (87). 2. Fibers containing thiophosphorylated myosin, which is resistent to phosphatases, remain contracted even when Ca2+ is removed (86, 88). 3. Addition of a Ca2 -insensitive proteolytic fragment of myosin light chain kinase results in a Ca2 -independent contraction (89). 4. In some preparations there is a good correlation between the extent of myosin phosphorylation and steady-state tension development whether the myosin phosphorylation level is achieved by phosphorylation or by dephosphorylation of previously phosphorylated myosin (86). +

+

+

13. REGULATION OF CONTRACTILE ACTIVITY

397

During a sustained contraction there appears to be an uncoupling of the relationship between phosphorylation and tension. In some muscle preparations, particularly those derived from arterial smooth muscle, tension is maintained for long periods of time even though myosin phosphorylation has returned to near resting levels (79-82, 90-92). In these muscles phosphorylation correlates quite well with the shortening velocity of the muscle. In other words, the velocity of shortening initially increases following a stimulation and gradually returns to resting levels at the same rate as does the phosphorylation of myosin. Under these conditions tension is presumably supported primarily by slowly cycling, dephosphorylated cross-bridges. There is also a concomitant drop in energy consumption (80, 93, 94). Murphy and his colleages termed this state “latch” in analogy with the molluscan “catch” state (79). It is a state where contraction can be maintained with low expenditure of energy. The mechanism by which the latch state is maintained is not known. Aksoy et a f . have postulated that there are two calcium-dependent regulatory systems operating in smooth muscle (90). The first is the calcium-calmodulin-dependent myosin light chain kinase which phosphorylates myosin. It is proposed that the myosin light chain kinase is more sensitive to calcium than the second system. When a stimulus is applied to the muscle the sarcoplasmic calcium levels increase and both calcium-dependent systems are activated. This results in myosin phosphorylation and tension development. After a few minutes the free calcium levels are lowered below the threshold for activation of myosin light chain kinase but remain above the threshold for the putative second system. This results in dephosphorylation of myosin with a concomitant drop in shortening velocity. Now the tension is maintained primarily by the slowly cycling, dephosphorylated cross-bridges. When the sarcoplasmic calcium is further lowered below the threshold for the second system, the muscle relaxes since now neither phosphorylated nor dephosphorylated cross-bridges can support tension. The nature of the putative second calcium-dependent regulator is not known. As discussed previously, smooth-muscle myosin itself can bind calcium, but it is unlikely that its calcium affinity is strong enough to function as the second regulator. Other regulatory systems have been proposed for smooth muscle which operate at the level of the thin filament such as leiotonin (95) and caldesmon (96, 97). Caldesmon is a calmodulin-binding protein present in a 1:26 molar ratio with actin which can reversibly regulate the MgATPase activity of myosin in v i m in a Ca2+ dependent manner ( 9 7 ~ ) . The latch hypothesis is not universally accepted. Using tuenia coli muscle, Siegman et al. (94) describe a state in muscle contraction where tension can be maintained with low energy utilization and where the shortening velocity is low. However, a pronounced decline in myosin phosphorylation levels is not seen when this occurs. Based on energetic measurements they also suggest that latch bridges do not occur and suggest that the reduced shortening velocity is instead

398

JAMES R . SELLERS AND ROBERT S. ADELSTEIN

due to an intrinsic Ca2 -dependent modulation of the shortening velocity of the entire myosin population (94).Several other smooth-muscle fiber types maintain high phosphorylation levels along with tension throughout the stimulus. An example of this is the methacholine-induced contraction of canine tracheal muscle (83).In this system a good correlation between myosin phosphorylation and steady-state tension was made, however shortening velocity was not measured. Gerthoffer and Murphy (81)argue that latch does occur in rabbit tracheal muscle but that the Ca2 concentrations required for tension and myosin phosphorylation are more nearly the same, making a latch state more difficult to observe experimentally. Moreover, Kamm and Stull also see a latch state using bovine tracheal muscle that has been electrically stimulated (92).This method of stimulation produces a much faster contractile response than those elicited pharmacologically, indicating that diffusion of agonist may be limiting in some muscle preparations. Skinned fiber preparations are also proving useful in elucidating the mechanism of contraction. Two such skinned fiber preparations from gizzard and uterus have demonstrated a close correlation between tension and phosphorylation (85-87). It is possible to obtain Ca2+-insensitive contractions in such preparations by two different techniques that have already been discussed. The first is to use a proteolytic fragment of myosin light chain kinase which does not require Ca2+ or calmodulin for activity (89).The second is to thiophosphorylate the myosin in the fibers (86, 87). These experiments demonstrate that the myosin light chain kinase phosphorylating system is the only regulatory system operating in the skinned fibers. There are skinned fiber preparations, however, that have manifested latch-like behavior. In one skinned arterial smooth-muscle preparation, a marked dissociation between the Ca2 -dependence of steady-state tension and light chain phosphorylation was observed provided that the experiment was conducted by decreasing the calcium concentration of contracted muscle from l o p 5 M free calcium (84).This was interpreted as evidence that a muscle can enter latch only from an active state. Hoar et al. showed that a latch-like state could be produced by dephosphorylating a skinned chicken gizzard muscle preparation in the presence of calcium by the addition of an exogenous phosphatase (98). Under these conditions, the fibers continued to maintain tension even though the myosin was not phosphorylated. A skinned preparation of gizzard muscle prepared by Barsotti et al. (36) demonstrates that Ca2+ can directly modulate the unloaded shortening velocity at a constant level of phosphorylation. This is supportive of the data of Siegman et al. (94) from their intact taenia coli preparations and is consistent with skinned fiber experiments from rat portal vein (99). The differences between the various skinned fiber preparations that show latch and those that do not, is not entirely clear. It is possible that the differences may +

+

+

13. REGULATION OF CONTRACTILE ACTIVITY

399

be due to loss of a diffusable protein or proteins. In both cases, however, phosphorylation is a prerequisite for muscle contraction. The latch state is only observed when one begins to dephosphorylate actively contracting muscle in the presence of calcium. It is clear that in order for the processes involved in smooth-muscle contraction to be understood, the nature of the second proposed calcium-dependent regulatory system must be described and the factors modulating shortening velocity must be determined.

2. Studies with an in Vitro Motility Assay Another system may prove useful for investigating shortening velocity. Sheetz, Spudich, and co-workers have developed an in vitro motility assay whereby the movement of myosin-coated beads (0.5 pm in diameter) can be measured on an actin substratum derived from microdissection of the algae, Nitellu (100, 101). From numerous studies it has been concluded that this system represents an in vitro model of unloaded shortening velocity in the muscle. Studies with smooth-muscle myosin reveal that rapid bead velocity is strongly dependent upon phosphorylation of myosin and that this rapid velocity can be reversed by dephosphorylation (Fig. 5 ) (102). It was observed that with fully phosphorylated myosin, the velocity was essentially “all or none.” That is, above 10 pg/ml phosphorylated myosin added to beads there was a constant velocity of movement and below this concentration there was no movement. However, if mixtures of fully phosphorylated and fully unphosphorylated myosin are added at a constant total concentration to the beads, intermediate velocities are obtained which decrease as the percentage of unphosphorylated myosin increases (102). This implies that the unphosphorylated myosin is retarding the movement of the phosphorylated myosin, presumably by virtue of its lower cycling rate. This may be one means by which graded shortening velocities are obtained in systems where the phosphorylation levels decline during tension maintenance. It will be of interest to test the effect of Ca2+ concentration on the velocity of bead movement in this in vitro system. OF MYOSINLIGHTCHAINKINASEBY D. PHOSPHORYLATION

PROTEIN KINASES The enzyme myosin light chain kinase can serve as a substrate for a number of different kinases, including CAMP-dependent protein kinase (103-105), cGMPdependent protein kinase ( 1 0 9 , and protein kinase C (106, 107). Although there are a number of important similarities among myosin light chain kinases isolated from vertebrate striated muscle, smooth muscle, and nonmuscle cells, major

400

JAMES R. SELLERS AND ROBERT S. ADELSTEIN 0.2

5 -> 0.1 0

Unphos

Phos Phos Phos Phos Dephos Dephos Dephos Rephos Rephos Dephos

FIG. 5. Effect of reversible phosphorylation-dephosphorylation of smooth-muscle myosin on bead movement. Unphosphorylated myosin was phosphorylated, dephosphorylated, and rephosphorylated as described by Sellers ef al. (27). After each manipulation a sample was mixed with beads and its motility assayed [from Ref. (102)].

differences exist, including their ability to serve as substrates for other kinases (see Volume XVII, Chapter 4). Myosin light chain kinase from smooth-muscle and nonmuscle cells can be rapidly and stoichiometrically phosphorylated in vitro by a number of different protein kinases which, in some cases, have a marked effect on myosin light chain kinase enzymic activity in vitro. Skeletal and cardiac muscle myosin light chain kinases are relatively poor substrates for CAMP-dependent protein kinase, and phosphorylation of these kinases does not alter their activity (108, 109). Studies carried out in vitro have shown the following general pattern of phosphorylation. When calmodulin is bound to smooth-muscle myosin light chain kinase, one mole of phosphate can be incorporated per mole of enzyme. Exhaustive tryptic digestion of the denatured myosin light chain kinase, which has previously been phosphorylated when calmodulin is bound, results in the liberation of one phosphorylated peptide which can be identified following separation by two-dimensional peptide mapping (105, 106). When calmodulin is not bound to myosin light chain kinase, CAMP-dependent protein kinase can incorporate 2 mol of phosphate into myosin light chain kinase.

13. REGULATION OF CONTRACTILE ACTIVITY

40 1

Peptide mapping of an exhaustive tryptic digest of the denatured diphosphorylated myosin light chain kinase reveals two phosphorylated peptides. One of these phosphopeptides comigrates with the peptide that is phosphorylated when calmodulin is bound to myosin light chain kinase. The additional site that is phosphorylated is presumably blocked when calmodulin is bound to myosin light chain kinase. Interestingly, inspection of the 2-dimensional tryptic peptide maps suggests that the same phosphorylated tryptic peptides are found in myosin light chain kinases isolated from turkey gizzard, bovine trachea, and human platelets, which have molecular weights of 130,000, 150,000, and 90,000, respectively (105,106).This implies that the sites phosphorylated by CAMP-dependent protein kinase are conserved, despite a large variation in the size and origin of the myosin light chain kinase molecules. In vitro, the effect of phosphorylating myosin light chain kinase is only observed when 2 mol of phosphate are incorporated by CAMP-dependent protein kinase (103-107).Phosphorylation of the site available when calmodulin is bound to myosin light chain kinase has no effect on enzymic activity. Conversely, phosphorylation of both sites results in a decrease in the apparent ability of the kinase to bind calmodulin and hence in the enzymic activity at a given calmodulin concentration. A 10- to 15-fold decrease in the amount of calmodulin required to achieve 50% activation of myosin light chain kinase has been reported for the turkey gizzard enzyme (103).Somewhat smaller effects have been reported for the tracheal and platelet enzyme (105). The decreased ability of dephosphorylated myosin light chain kinase to bind calmodulin can be reversed, using a purified phosphatase from smooth muscle (21).Both sites can be rapidly dephosphorylated when calmodulin is not bound. However, when calmodulin is bound to the diphosphorylated enzyme, dephosphorylation is specific for the site that is blocked when calmodulin is bound to the unphosphorylated enzyme (20).Evidence has been published suggesting that both phosphorylated sites ccntribute to the weakening of calmodulin binding to myosin light chain kinase (106). The amino acid sequences of both phosphorylated sites have been determined (110,110a).The site phosphorylated when calmodulin is bound is very similar to other sites phosphorylated by CAMP-dependent protein kinase in containing the sequence -Arg-Lys-X-Ser- (110).Structural studies both at the peptide and at the cDNA level suggest that both phosphorylated sites are located carboxy terminal to the catalytic site and the calmodulin-binding site, respectively (111, 1 11a). In addition to being phosphorylated by CAMP-dependent protein kinase, smooth-muscle myosin light chain kinase can also serve as substrate for cGMPdependent protein kinase (103,as well as protein kinase C (106,107).Bovine tracheal myosin light chain kinase cannot be phosphorylated by cGMP-dependent protein kinase when calmodulin is bound. When calmodulin is not bound 1

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mol of phosphate is incorporated into the same site that is phosphorylated by the CAMP-dependent enzyme when calmodulin is bound. As might be expected, phosphorylation by cGMP-dependent protein kinase has no effect on tracheal smooth-muscle myosin light chain kinase activity. Phosphorylation of human platelet myosin light chain kinase by cGMP-dependent protein kinase yields somewhat different results. In the presence of bound calmodulin, 1 mol of phosphate is incorporated into the same site that is phosphorylated by CAMPdependent protein kinase, but when calmodulin is not bound an additional mole of phosphate is incorporated into a new site that is not the same as that phosphorylated by CAMP-dependent protein kinase when calmodulin is not bound. However, even following the introduction of 2 mol of phosphate there is a relatively small decrease in the ability of platelet myosin light chain kinase to bind calmodulin compared to that seen following phosphorylation by CAMPdependent kinase (105). Two different laboratories reported the phosphorylation of smooth-muscle myosin light chain kinase by protein kinase C (106, 107). Although both agree that protein kinase C can incorporate 2 mol of phosphate into myosin light chain kinase when calmodulin is not bound and that diphosphorylation of myosin light chain kinase weakens calmodulin binding, there is disagreement on whether one or both of the sites phosphorylated by protein kinase C are different from those phosphorylated by CAMP-dependent protein kinase (106, 107). The in vitro demonstration that phosphorylation of myosin light chain kinase by CAMP-dependent protein kinase can weaken calmodulin-binding and thereby decrease its ability to phosphorylate myosin suggests that this may be a mechanism by which this kinase can be regulated in vivo. Such a mechanism might explain, at least in part, why certain hormones are able to relax smooth muscle. To date, experiments demonstrating a role for this mechanism in vivo have not been conclusive. Using skinned gizzard smooth muscle fibers, Kerrick and Hoar (112) demonstrated that addition of the catalytic subunit of CAMP-dependent protein kinase inhibited the development of Ca2 -activated tension. Moreover, following a maximal Ca2 -activated contraction, addition of both the regulatory and catalytic subunits of CAMP-dependent protein kinase resulted in a slow relaxation of the fibers and addition of 10-4MCAMP produced a faster relaxation. This relaxation was not seen when the skinned fiber bundle was contracted irreversibly using ATP-$3, suggesting that the relaxation that occurred following the addition of ATP (but not ATP-yS) was due to dephosphorylation of myosin. This dephosphorylation, in turn, might have resulted from a relative decrease in the activity of myosin light chain kinase due to CAMP-dependent phosphorylation of myosin light chain kinase. Whether myosin light chain kinase was in fact phosphorylated during these experiments was not determined. In experiments carried out with intact canine tracheal muscles, de Lanerolle et al. (113) showed that a +

+

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methacholine-induced contraction could be relaxed by forskolkin, which induced a rise in intracellular CAMP. An antibody raised to gizzard myosin light chain kinase was used to precipitate the tracheal enzyme and the amount of phosphate incorporated into tracheal myosin light chain kinase was quantitated. The increase in cAMP was accompanied by an increase in phosphate incorporation into myosin light chain kinase to a maximum of 1.9 mol phosphate/mol of myosin kinase. These studies imply that myosin light chain kinase is a substrate for CAMP-dependent protein kinase in intact smooth muscle. They also demonstrate that the enzyme can be phosphorylated even after a methacholine-induced contraction, a time when calmodulin is most likely bound to much of myosin light chain kinase. However, there remains some uncertainty as to the location of the phosphorylated sites, since the enzyme was not subjected to 2-dimensional peptide analysis in order to be sure that phosphorylation was confined to the sites phosphorylated by CAMP-dependent protein kinase in vitro. There are a number of compelling reasons to believe that cAMP may also cause relaxation of smooth muscle through other mechanisms. A rise in cAMP has been associated with a decrease in intracellular Ca2 in smooth muscle (114) and this in conjunction with the mechanism previously discussed might explain the ability of this ligand to relax smooth muscle. This type of mechanism might explain the ability of isoproterenol to relax smooth muscle which has undergone a sustained contraction and is in the latch state (see Section II,C,l), a condition during which contraction is maintained but light chain phosphorylation is substantially decreased (76). Furthermore, Miller et al. (115) treated strips of tracheal smooth muscle with isoproterenol and then assayed myosin light chain kinase activity at 4 and 100 pA4 Ca2+ . They saw no difference in the Ca2 activation curve between untreated and treated strips, although a control using purified myosin light chain kinase that had been phosphorylated in vitro did show decreased activity under the same assay conditions. (For further discussion of this aspect of myosin light chain kinase phosphorylation, see Volume XVII, Chapter 4). In summary, the ability of CAMP-dependent protein kinase to phosphorylate myosin light chain kinase at two sites and alter the binding of calmodulin has been demonstrated for enzymes isolated from a number of different smoothmuscle cells as well as platelets. Whether this constitutes a mechanism by which cAMP acts to regulate the contractile activity in muscle and nonmuscle cells remains to be demonstrated conclusively. Present evidence indicates that under certain circumstances it might play a role, but other potential effects of CAMP, such as decreasing the concentration of intracellular Ca2 may be of equal or of even greater importance. The finding that protein kinase C can also phosphorylate myosin light chain kinase isolated from gizzard smooth muscle has only been demonstrated in vitro. It will be necessary to show that this phosphorylation can take place in vivo before its significance can be assessed. +

+

+

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111.

Role of Phosphorylation in Modulating Contractile Activity of Striated Muscle Proteins

A.

PHOSPHORYLATION OF MYOSIN

Phosphorylation of myosin was first described for the 18,500-dalton light chain of rabbit skeletal-muscle myosin (116). However, actin-activation of skeletal-muscle myosin MgATPase activity differs from smooth muscle in that phosphorylation of myosin is not required for activation to occur. Indeed, the initial reports found no difference between phosphorylated and unphosphorylated skeletal-muscle myosin (117). Pemrick was the first to report an effect of phosphorylating the 18,500-dalton light chain of skeletal-muscle myosin (118). She found a I .5-to 2-fold increase in the actin-activated MgATPase activity at low concentrations of Ca2 , and low ionic strength. A steady-state kinetic analysis of the system showed that phosphorylation decreased the Kappof actin for myosin with no apparent effect on V,,,. In another study, Persechini and Stull observed a 2.4-fold decrease in the K , for actin, from about 6 phi’ to 2.5 pA4, with no significant change in V,,, in response to phosphorylation (119). This result obtained using skeletal-muscle myosin contrasts to the findings of Sellers et al. using turkey gizzard smooth-muscle HMM (56).The latter study found a 4fold decrease in the K , of phosphorylated myosin for actin and a 25-fold increase in V,,,. A number of studies, using intact fast-twitch skeletal muscle, have demonstrated that myosin light chain phosphorylation has little effect on the development of maximum tension (120-122) or the unloaded speed of muscle shortening (121). On the other hand, Stull and his co-workers using an in situ preparation have shown that phosphorylation of the 18,500-dalton light chain of fast skeletalmuscle myosin can occur at physiologically relevant contractile stimuli (i.e., with respect to frequency and duration) (123, 124). Thus, electrical stimulation (5 Hz for 20 s) of the rabbit plantaris muscle produced an increase in phosphorylation from 0.17 to 0.45 mol of phosphate/mol of myosin light chain. This increase in phosphate content was accompanied by a 1.6-fold increase in maximal isometric twitch tension (123). Similar results have been reported for other mammalian fast-twitch muscles. In contrast, tetanic muscle contraction resulted in no increase in isometric twitch tension and only a small increase in myosin light chain phosphorylation in rabbit and rat slow-twitch muscle (123, 124). This lack of increase in phosphorylation in the case of slow skeletal muscle is in contrast to that observed by Westwood et al. (125). Persechini et al. studied the effect of phosphorylation on isometric tension using skinned fibers from rabbit skeletal muscle (126). At low concentrations of Ca2+ (0.6 phi’),there was a 50% increase in isometric tension which correlated with an increase in phosphorylation of the 18,500-dalton light chain. This effect +

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was not seen at Ca2+ concentrations required for the development of maximal isometric tension. This finding has been related to an earlier observation that showed a relation between skeletal-muscle phosphorylation and potentiation of isometric twitch tension (127). Cardiac-muscle myosin can be phosphorylated by a myosin light chain kinase that has been isolated from bovine hearts (109). This phosphorylation has not been shown to alter the actin-activated MgATPase activity of myosin. Although it is tempting to reason that cardiac-muscle myosin may show an effect similar to that reported for skeletal muscle (i.e., that phosphorylation of myosin may cause a modest alteration in the K,,, of myosin for actin at low concentrations of Ca2+) no definitive studies along these lines have been carried out. In summary, the most complete data available relate to skeletal fast muscle. Using fast-twitch skeletal muscle from a number of different sources, an increase in phosphorylation of the 18,500-dalton light chain has been demonstrated following stimulation. This increased phosphorylation has been correlated with an increase in isometric twitch tension at concentrations of Ca2 below those required for the development of maximal isometric tension. Thus, phosphorylation of fast-twitch-skeletal-muscle myosin appears to modulate muscle contraction under certain circumstances. Definitive physiological and biochemical studies relating vertebrate slow-twitch muscle and cardiac-muscle phosphorylation to changes in contractile activity are not available. +

OF TROPONIN AND TROPOMYOSIN B . PHOSPHORYLATION

Troponin I, troponin T, and tropomyosin have all been shown to contain phosphate in vivo and can be phosphorylated in vitro (128-131). However, the only reversible phosphorylation that has been shown to result in a potentially significant alteration in contractile activity is that found in cardiac troponin I. Cardiac-muscle troponin I differs from its skeletal muscle counterpart in containing an additional 26 amino acids at the amino terminus. In this region, at position 20, there is a serine residue that is rapidly phosphorylated by CAMP-dependent protein kinase (129, 132). Studies with intact cardiac tissues show that epinephrine can induce phosphorylation at this site (133). Robertson et al. (134) showed that cardiac troponin complexes that were phosphorylated at serine 20 of troponin I released Ca2+ more rapidly than unphosphorylated troponin complexes. It is possible, therefore, that this phosphorylation might explain the increased rate of relaxation seen in cardiac muscles following treatment with epinephrine. What has not yet been determined is whether troponin I phosphorylation is the limiting step in determining the increased rate of cardiac relaxation or whether an increase in Ca2+ uptake by the sarcoplasmic reticulum might be of primary importance. In any event, phosphorylation of cardiac-muscle troponin I is at present the only example of tro-

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ponin phosphorylation shown to alter the binding of Ca2 to the troponin complex, with the potential to alter contractile activity. +

IV. Phosphorylation-Dependent Regulatory Systems in Invertebrate Muscles

The existence of a myosin-linked regulatory system was first shown in molluscan muscle where Ca2+ binding to the myosin directly regulates its activity (8). The light chains of scallop striated myosin are not phosphorylated (135). There are other invertebrate myosins where a phosphorylation-dependentregulatory system has been described. Myosin from Limulus, the horseshoe crab, is an example of such a system. Phosphorylation of the regulatory light chains of Limulus myosin markedly increases the actin-activated MgATPase activity and the calcium-calmodulin-dependent myosin light chain kinase responsible for this phosphorylation has been purified (11). There is also a functional troponintropomyosin system in Limulus making it a dually regulated system (12). Based on experiments with skinned Limulus muscle fibers, Kerrick and Bolles (136) suggest that phosphorylation of myosin does not play a major role in Limulus muscle contraction. More work will be required in order to resolve this difference. There have been reports of phosphorylation of the myosin from locust muscle (137). Paramyosin, a protein found at the core of many invertebrate thick filaments, is also phosphorylated but the role of this phosphorylation and the enzyme responsible for catalyzing it are not known (138, 139).

V.

Regulation of Cytoplasmic Myosins

A.

LOWEREUKARYOTES

Myosin that is structurally similar to that from muscle sources has been purified and characterized from a number of lower eukoryotic organisms including Acanthamoeba castellanii (140, 141), Dictyostelium discoideum (142), and Physarum polycephalum (143, 144). Each of these myosins has two heavy chains and four light chains and can form bipolar filaments via self-association of the tail portion. It has been known for a number of years that Acanthamoeba also has a pair of low-molecular-weight myosin isozymes (termed myosin IA and IB), which resemble myosin S1 (see Fig. 3) in that they have a single heavy chain of about 125,000 daltons and one or two light chains (145, 146). Myosin I does not form filaments. Recently, a similar myosin has been described in Dictyostelium (147).

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The actin-activated MgATPase activity of myosin I is considerably activated by phosphorylation of the heavy chain at its aminoterminal end (148). This reaction is catalyzed by a kinase which has been purified (149). The function that this myosin plays in the cell is not known, although it can bind to and cause the movement of beads in the Nitella-based in vitro motility system described in Section II,C,2 (150). Phosphorylation also regulates the activity of the larger Acanthamoeba myosin I1 although in this case phosphorylation markedly decreases the actin-activated MgATPase activity (151, 152). The mechanism by which this occurs has not been elucidated, but it is known that dephosphorylation of these myosins favors filament assembly (151-153). However, work with Acanthamoeba myosin I1 indicates that filament formation per se is not sufficient to account for actinactivation of the myosin’s MgATPase activity (153). The site of the phosphorylation in Acanthamoeba is a region located near the carboxy terminus of the “tail” of the myosin (151). A maximum of six serines (three on each chain) can be phosphorylated by a partially purified kinase. These sites have been sequenced (154). Thus, the phosphorylation sites are located at the opposite end of the molecule from the active sites. If these sites are digested away by treatment with chymotrypsin which removes about 9000 daltons from the carboxy terminal end, the myosin can no longer form filaments and is inactive (155). Kuznicki et al. showed that there was an interesting intermolecular regulation of this myosin. They found that phosphorylation effects could be transmitted through the filament to affect the conformation of the filament as a whole (156). Dictyosteliurn myosin seems to be regulated in a manner similar to Acanthumoeba myosin (142) although one of the light chains of Dictyostelium can also be phosphorylated. The role of this phosphorylation has not been elucidated. An antibody against this light chain strongly inhibits motility of myosincoated beads in the Nitella based in vitro motility assay, suggesting that it may be involved in regulation (157).

B.

PHOSPHORYLATION OF VERTEBRATECYTOPLASMIC MYOSIN

1.

In Vitro Studies

Myosin has been found in virtually every cell type including erythrocytes where it is present in very low amounts (158, 159). The best-characterized cytoplasmic myosins from vertebrate sources are from platelets (160, 161), macrophages (162), thymus (47, 163), lymphocytes (164), and intestinal epithelial brush border (165, 166). For each of these myosins, the actin-activated MgATPase activity is increased by phosphorylation. It is not known whether the mechanism of the phosphorylation-dependent regulation is similar to that described above for gizzard smooth-muscle myosin although Wagner et al. (47)

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have shown that there are potentially important differences between thymus myosin and gizzard smooth-muscle myosin. Thymus myosin may be regulated in a manner similar to that described for aorta myosin (47, 63a). Evidence that the heavy chain of vertebrate nonmuscle myosin can be phosphorylated in vivo has been published for myosin isolated from a number of sources including macrophages (167),lymphocytes (164),fibroblasts (168),retina (169), and the brain (170). Myosin purified from a murine myeloid leukemia cell line was shown to be phosphorylated on the heavy chain and preliminary evidence was presented suggesting dephosphorylation of the heavy chain was necessary before light chain phosphorylation could cause an increase in the actinactivated MgATPase activity of myosin (171).Trotter has reported that a kinase isolated from brain can phosphorylate macrophage myosin on the carboxy-terminal portion of the heavy chain at the same site that is phosphorylated in vivo (172).

2. Studies with Intact Cells The functions ascribed to cellular myosins are numerous and varied. Although some of the evidence for actomyosin playing a role in cell function is convincing, much of it is circumstantial. Some of the cellular processes attributed to the contractile proteins include cytokinesis, karyokinesis, organelle translocation, endocytosis, secretion, cell shape change, capping, cell locomotion, and contractility of microvilli (1-3). As an example, the role of phosphorylation in platelet function is discussed in more detail. Numerous stimulants, such as ADP, thrombin, or the cationophore, A 23187, can stimulate intact platelets, leading to a sequence of well-defined responses [for review, see Ref. (173)l.The stimulated platelets undergo a rapid change in cell shape from a smooth disk-like structure to irregular spheres bearing filopodia and pseudopodia. This is followed by aggregation and then granule secretion. Numerous studies have found a rapid increase in myosin phosphorylation and an increase in the amount of myosin associated with the triton-insoluble cytoskeleton following platelet activation (174-1 76). Nachmias et al. (175) took advantage of the fact that chilling of platelets ellicits a slow change in shape with a time course that is easily studied. Their results show that myosin becomes associated with the triton-insoluble cytoskeleton as the shape change occurs. It was also observed that myosin becomes phosphorylated during the shape change. The shape change, myosin association with the cytoskeleton, and myosin phosphorylation were all reversible upon rewarming the platelets (175). Several other studies have shown that activation of platelets is associated with a net phosphorylation of the myosin pool and an increase in myosin associated with cytoskeleton (177, 178). Treatment of platelets with prostaglandin D, or forskolin inhibited the phosphorylation of

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myosin and prevented the formation of cytoskeletons (179). These agents are known to raise CAMP levels, and their negative effects on myosin phosphorylation are similar to those seen in smooth-muscle preparations (113). ADP can induce the shape change response without concomitant secretion or aggregation (176). It was shown that under these conditions the phosphorylation level of the myosin rapidly increased to about 1 mol/mol myosin (r1/2 = 1.5 sec) from basal levels (0.2 mol/mol) while the shape change took slightly longer ( t l 1 2 = 2.5 s). The myosin phosphorylation remained elevated for 10-20 seconds and gradually returned to baseline within 5 min while the platelets retained their changed shape. This is reminiscent of the situation seen during the latch state in intact smooth muscle (90). It will be very interesting to determine whether similar mechanisms are operating in the two systems. Together these studies indicate that the change in cell shape may be due to an actomyosin system since the phosphorylation of no other protein correlated with the response. Platelet secretion, however, seems to require the phosphorylation of a protein of about 40,000 daltons perhaps in addition to myosin phosphorylation (180182). Nishizuka (182) found that treatment of platelets with the phorbol ester TPA (12-0-tetradecanoylphorbol- 13-acetate) resulted in phosphorylation of the 40,000-dalton polypeptide with only slight myosin light chain phosphorylation. Under these conditions he observed a very slow secretory response. Based on these data he suggested that secretion may require only phosphorylation of the 40,000-dalton protein alone. However, rapid secretion always seems to be accompanied by both myosin and 40,000-dalton protein phosphorylation (180, 181). The 40,000-dalton protein has recently been reported to be an inositol trisphosphate 5’-phosphomonoesterase (181). OF MYOSINBY PROTEIN KINASEC C. PHOSPHORYLATION

Myosin light chain kinase is not the only enzyme that catalyzes phosphorylation of myosin. In this section we consider enzymes that have been shown to phosphorylate myosin on the 20,000-dalton light chain, as does myosin light chain kinase. There is no evidence that the 17,000-dalton light chain of myosin can be phosphorylated in vivo or in vitro. In considering phosphorylation of the 20,000-dalton light chain of myosin it is important to emphasize that phosphorylation of the isolated 20,000-dalton light chain does not necessarily imply that this light chain can be phosphorylated when it is bound to the heavy chain of myosin. Two notable examples of kinases that phosphorylate the isolated 20,000-dalton light chain but not intact myosin, are CAMP-dependent protein kinase (183) and the tyrosine kinase associated with epidermal growth factor (184). Although synthesis of myosin light chains occurs independently of heavy chain synthesis, there is no known function for the

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isolated myosin light chains and thus the physiological significance of these phosphorylations is in doubt. The calcium-activated, phospholipid-dependent kinase, protein kinase C has been shown to phosphorylate smooth-muscle myosin (185) and platelet myosin in vitro (186). Evidence has also been published suggesting that myosin can be phosphorylated by protein kinase C in intact platelets (286, 187). The site phosphorylated by protein kinase C is located on the 20,000-dalton light chain and in the case of smooth-muscle myosin is known to be a threonine residue (185). Two-dimensional peptide mapping of the 20,000-dalton light chain isolated from both purified platelet and smooth-muscle myosin has been carried out following phosphorylation by myosin light chain kinase and protein kinase C. Two different tryptic phosphopeptides, one due to protein kinase C phosphorylation and the second due to myosin light chain kinase phosphorylation, have been identified. Recent work has shown that protein kinase C phosphorylates two sites on the 20,000 Da light chain of smooth muscle myosin. Peptide mapping and sequence analysis show that one site is threonine-9 and the other either serine-1 or -2. Interestingly, phosphorylation of smooth-muscle myosin or HMM by protein kinase C has a different effect on the actin-activated MgATPase activity than does phosphorylation by myosin light chain kinase. When 2 mol of phosphate are incorporated into unphosphorylated myosin (i.e., 1 mol/light chain) by protein kinase C, the actin-activated MgATPase activity remains very low, in contrast to the effect of phosphorylating myosin by myosin light chain kinase. Moreover, myosin that has been phosphorylated by protein kinase C is a much poorer substrate for myosin light chain kinase than is myosin that is unphosphorylated. This is due to a weakening of the apparent affinity of the phosphorylated myosin for myosin light chain kinase from 4.25 pikf to 37.0 pikf (185). When myosin is first phosphorylated by myosin light chain kinase and this is followed by protein kinase C phosphorylation, 4 mol of phosphate can be incorporated/myosin molecule (i.e., 2 mo1/20,000-dalton light chain; one at a serine residue and one at threonine residue). The actin-activated MgATPase activity of this myosin is about one-half that of myosin which has been phosphorylated only by myosin light chain kinase (188). Studies carried out with turkey gizzard smooth-muscle HMM indicate that the decrease in actin-activated MgATPase activity is due to a decrease in the ability of myosin, which has been phosphorylated at both sites, to bind to actin (185). These in vitro studies suggest that protein kinase C phosphorylation of myosin should act either to inhibit the contractile response of unphosphorylated myosin or to decrease the activation of myosin by myosin light chain kinase. This does not imply that protein kinase C will terminate contractile activity, in fact its effect may be to prolong contractile activity albeit at a reduced amplitude. Thus, protein kinase C may decrease, but not abolish,,the rate of cross-bridge turnover.

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41 1

Experiments using intact human platelets have been interpreted in this manner (187). As previously discussed, if intact human platelets are treated with thrombin after being equilibrated with 32Pi,there is an increase in 32P label at the site on myosin light chain phosphorylated by myosin light chain kinase. Under the conditions described by Naka et af. (186) phosphorylation of myosin is mostly confined to this site and is associated with the rapid release of granules. However, when intact platelets are treated with TPA, an agent that is known to activate protein kinase C, the 20,000-dalton light chain of myosin has been shown to be phosphorylated at two sites: the site phosphorylated in vitro by protein kinase C and that phosphorylated by myosin light chain kinase. By 5 min after addition of TPA a majority of the phosphate is located in the protein kinase C site and the platelet release reaction is delayed. The distribution between the protein kinase C site and the site phosphorylated by myosin light chain kinase can be altered by treating platelets with H-7 (1-(5-isoquinolinesulfonyl)-2-methylpiperazine)an agent that has been shown to inhibit protein kinase C (and cyclic nucleotidedependent protein kinases) more effectively than calmodulin-dependent kinases (187, 189). These experiments are of interest because they show that in intact platelets, myosin is a substrate for protein kinase C, and they raise the possibility that this might also be true for other nonmuscle cells as well as smooth muscle cells. However, they also raise at least three important questions. 1. Why is the treatment of the intact platelets associated with phosphorylation at the myosin light chain kinase site? Unpublished observations have indicated that TPA cannot directly activate myosin light chain kinase (190), and it seems likely that the phosphorylation may be due to an increase in calcium, perhaps from intracellular stores. 2. How does an enzyme that is thought to be active only when bound to the membrane fraction gain acess to myosin filaments? 3. What is the physiological effect of phosphorylating myosin at the protein kinase C site? The answers to the last two questions are unknown, but data from preliminary experiments suggest that at least one fraction of platelet myosin may be associated with the membrane (191). With respect to the last question, work by Hidaka and his colleagues indicates that phosphorylation of platelet myosin by protein kinase C appears to be associated with a decrease in the rate at which platelets release their granules, compared to phosphorylation at the myosin light chain kinase site alone (187).The implication of these studies, as well as the in vitro work previously outlined, is that phosphorylation at the myosin light chain kinase site on the 20,000-dalton light chain is required for contractile activity,

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and that phosphorylation by protein kinase C will modify this activity, resulting in a decreased, but perhaps prolonged contractile response. In summary, the 20,000-dalton light chains of platelet and smooth-muscle myosins are substrates for protein kinase C . Studies conducted in vitro with smooth-muscle myosin and heavy meromyosin suggest that phosphorylation by protein kinase C alters the effects of phosphorylation by myosin light chain kinase. The significance of these studies with respect to the alteration of contractile activity of platelets and smooth muscle in vivo remains to be demonstrated. The exact function of myosin heavy chain phosphorylation is still under study. It is of interest that the effects of phosphorylating the 20,000-dalton light chain of myosin by protein kinase C as well as phosphorylating the carboxy terminus of the myosin heavy chain appear to modulate the effect of phosphorylating myosin by myosin light chain kinase. As such, they may constitute different mechanisms for modulating contractile activity in vertebrate cells.

VI. Summary Since 1980 there have been a number of advances in our understanding of how phosphorylation regulates and modulates contractile activity in vertebrate muscle and nonmuscle cells. Using purified proteins from avian gizzard, the most intensively studied smooth muscle to date, it is possible to reconstitute a reversible myosin phosphorylating system in vitro. Moreover, it has been learned that phosphorylation of both heads of myosin is required for actin-activation of the MgATPase activity and that the kinetic step that is regulated by phosphorylation is most likely phosphate release. The major conformational change that myosin is capable of undergoing following phosphorylation has been described in detail and an in vivo role for this conformational change (or a variant of it) is being actively sought in smooth-muscle and nonmuscle cells. How much of the mechanism for the regulation of avian myosin will be applicable to other vertebrate smooth-muscle and cytoplasmic systems remains to be seen. Evidence is already accumulating that there are important variations on the avian theme. Experiments documenting a major decrease in the amount of myosin phosphorylated following a sustained contraction provide good evidence for at least one other regulatory system that also requires calcium. How this system acts to modulate contractile activity and how it interacts with the myosin phosphorylating system is under study. Major progress has also been made in our understanding of how phosphorylation of vertebrate fast skeletal-muscle myosin acts to modulate contractile activity. One of the most exciting aspects of this work involves the development of techniques to study phosphorylation of live muscles in situ, in a number of

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species, including humans. Similar kinds of experiments should help to uncover the role of cardiac-muscle myosin phosphorylation in vertebrate muscle. Perhaps the greatest remaining challenge is to understand the role of myosin phosphorylation in regulating contractile activity in vertebrate nonmuscle cells. Although it is clear that there are some important similarities between phosphorylation and regulation of cytoplasmic and smooth-muscle myosins, there are most likely important differences too. Of particular importance in nonmuscle cells is the role of myosin phosphorylation in regulating filament assembly and the putative role of myosin heavy chain phosphorylation and protein kinase C phosphorylation in modulating contractile activity. How phosphorylation acts to regulate cytoplasmic contractile activity and the biological consequences of this activity remain an exciting area for investigation.

ACKNOWLEDGMENTS The authors wish to acknowledge all their colleagues who generously shared their results and hence contributed to this chapter. We also thank our editorial assistants, Imogene Surrey and Theresa Wilson. Mary Anne Conti kindly helped by reading and commenting on the manuscript.

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14

Protein Phosphorylation in Prokaryotes and Single-Celled Eukaryotes HOWARD V . RICKENBERG**t BEN H. LEICHTLING? *Department of Biochemisrry, Biophysics, and Generics The University of Colorado Health Sciences Center Denver, Colorado 80262 fDepartment of Molecular and Cellular Biology National Jewish Center for Immunology and Respiratory Medicine Denver, Colorado 80206

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Protein Phosphorylation in Prokaryotes .............................. A. Protein Kinases Encoded by Bacteriophages .......................

B. Intrinsic Bacterial Protein Kinases and Phosphoproteins . . . . . . . . . . . . . C. Conclusions on the Occurrence and Function of Protein Phosphorylation in Prokaryotes 111. Protein Phosphorylation in Single-Celled Eukaryotes . . . . . . . . . . . . . . . . . . . A. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Yeast . . . . .......................... . . . . . . . . . . . . . . . .

.......................................... D. Mucor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Blastocladiella F. Miscellaneous Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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G. Slime Molds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . H . Protozoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. General Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1.

440 447 450 451

Introduction

The covalent modification of proteins by phosphorylation constitutes a major mechanism of regulation. It was first recognized as such in the context of the hormonal control of the metabolism of glycogen in mammalian tissues. A CAMP-dependent protein kinase catalyzes the phosphorylation of a phosphorylase kinase, which in turn, by ATP-dependent phosphorylation, converts enzymically inactive phosphorylase b to active phosphorylase a . Ultimate regulatory control is exercized, depending on the tissue, by the hormones epinephrine and glucagon which activate membranal adenylate cyclase, leading to an increase in cellular CAMPand the activation of the CAMP-dependent protein kinase. A large body of information on the phosphorylation of proteins has accumulated since those germinal observations were made some thirty years ago. The proteins subject to phosphorylation and dephosphorylation include enzymes and structural proteins, particularly membranal proteins with a dynamic function such as the components of ion channels. It is now clear that the phosphorylations are catalyzed by several classes of protein kinases differing in their modes of regulation as well as in the proteins and amino acids phosphorylated; dephosphorylation is also catalyzed by different phosphoprotein phosphatases. The fact that many protein kinases are ultimately controlled by hormones, growth factors, and the antiviral, hormone-like interferons, suggests a role for protein phosphorylation in intercellular, integrative signal transmission in the multicellular organism as well as a role within the individual cell (I). In prokaryotic bacteria and unicellular eukaryotes, unlike in metazoa, the single cell constitutes the normal viable unit of life. Admittedly, the mating-type substances of certain fungi and aggregation in the slime molds bring about cellcell interaction. But these instances are marginal to a predominantly singlecelled mode of life. Be that as it may, there is a growing body of evidence for the occurrence of protein phosphorylation and protein kinases in both prokaryotes and single-celled eukaryotes. In view of what has been said earlier about the intercellular, integrative role of protein phosphorylation, questions arise as to the regulation of the activity of protein kinases and the function of protein phosphorylation in single-celled organisms. It may well be that an attempt to answer these questions will bring new insights into the evolutionary origins of protein phosphorylation as a mechanism of regulation.

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II. Protein Phosphorylation in Prokaryotes A.

PROTEIN KINASESENCODED BY BACTERIOPHAGES

An early claim (2) of the occurrence in Escherichia coli of a CAMP-stimulated protein kinase, able to catalyze the phosphorylation of calf thymus histone, was not confirmed; in retrospect it appears that the activity measured may have been that of a polyphosphate kinase that required a basic protein for maximal activity ( 3 ) .In 1973 Rahmsdorf et al. (4), however, found that several ribosomal, as well as nonribosomal, proteins became phosphorylated after infection of E. coli with phage T7. The protein that catalyzed the phosphorylation of the bacterial proteins was encoded by a phage gene transcribed early (4 min) after infection; uv irradiation of the phage blocked the appearance of the protein kinase activity, irradiation of the bacterial host did not ( 5 ) . The phage-encoded protein kinase also catalyzed the phosphorylation of histone; the activity of the enzyme was independent of cyclic nucleotides, irrespective of the substrate employed. The protein kinase encoded by phage T7 was purified approximately 5,000-fold from the ribosomal wash of infected bacteria. Unlike the protein kinases encoded by E. coli, discovered later (see below), the phage protein kinase catalyzed the phosphorylation of a number of exogenous basic proteins. Egg white lysozyme was the best substrate and was used in affinity chromatography for the purification of the enzyme. The optimal pH for the kinase reaction was 6.9-7.1 and the enzyme was maximally active at low ionic strength and with 15 mM magnesium. Its molecular weight is approximately 37,000. There is evidence that the primary function of the phage-coded protein kinase is the rapid arrest of the synthesis of host proteins. I n vivo as many as 40-50 proteins became phosphorylated after infection of E . coli with phage T7. Among these were several ribosomal proteins and the p’, the p-, and possibly the usubunit of the bacterial RNA polymerase; p’ was strongly and p weakly phosphorylated; u contained only traces of label. A protein of molecular weight 26,000 and unknown function, coisolated with the RNA polymerase, was also phosphorylated in vivo. Purified T7 protein kinase catalyzed the in vitro phosphorylation of E. coli RNA polymerase; the p’- and p-, but neither u- nor asubunits, were phosphorylated in virro; the bulk of the phosphate was incorporated into threonine and only traces appeared in serine. The core polymerase was a less efficient acceptor than the holoenzyme and DNA stimulated the phosphorylation of the holoenzyme significantly. I n vivo, the protein kinase was inactivated rapidly after its activity had reached a maximum. A similar phagecoded protein kinase was found after infection of E . coli with phage T3. The shutoff of the synthesis of host proteins coincided with the appearance of phagecoded protein kinase and thus the phosphorylation of the bacterial RNA poly-

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merase and of several ribosomal proteins would seem a likely mechanism for the inhibition of the synthesis of bacterial proteins (6, 7). The phosphorylation of bacterial proteins has been noted following the infection of the dimorphic bacterium, Caulobacter crescentus, by the small lytic phage, c$Cd 1 (8). Approximately 40 phosphorylated proteins were detected in bacterial extracts 10 min after infection, and additional proteins were phosphorylated if the infection was permitted to continue for 30 min. The addition of either rifampin or chloramphenicol to the medium prior to infection with phage completely blocked the phage-specific phosphorylation, suggesting that the relevant protein kinase was encoded by the phage. The occurrence of a kinase activity specific for the formation of protein acylphosphates in Caulobacter had been established earlier (see Section II,B ,1). The presumptive phage-encoded protein kinase catalyzed the phosphorylation of the P’-subunit of the host RNA polymerase as well as that of about eight other DNA-binding proteins, isolated on a DNA-cellulose column and not found in uninfected cells. Phosphoproteins were found also in the ribosomes and the outer and inner membrane fractions. All but two of the phosphoproteins found in phage-infected bacteria were sensitive to alkaline phosphatase. The phosphorylated amino acid of the P’-subunit of the RNA polymerase was primarily phosphoserine. It might be pointed out parenthetically that different bacteriophages may employ different strategies for shutting off bacterial protein synthesis; thus E . coli phage T4 codes for an enzyme that catalyzes the ADP-ribosylation of the a-subunit of the bacterial RNA polymerase (9).

B.

INTRINSIC BACTERIALPROTEINKINASES AND PHOSPHOPROTEINS

1.

General

Earlier observations bearing on the profound significance of protein phosphorylation in higher eukaryotes suggested the occurrence of a similar mechanism of control in bacteria. Several preliminary reports (10, 11) indicated the existence of protein phosphorylation catalyzed by bacterial protein kinases; however these preliminary reports were not followed up and in retrospect it is difficult to evaluate the validity of the original claims. Furthermore there are several types of phosphate transfer reactions in which either an acyl rather than an ester linkage is formed or in which the phosphate donor is not a nucleoside triphosphate; these reactions are not considered in depth here. For example, an unusual enzyme was observed in the bacterium C . crescentus. The enzyme catalyzed the transfer of the y-phosphate of ATP to added basic proteins with the formation of acylphosphates; the physiological function of the reaction is not clear (12). ATP is not the only phosphate donor in the phosphorylation of bacterial proteins. A

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case in point are the carrier proteins of the phosphoenolpyruvate-sugar phosphotransferase systems (PTS). They catalyze the concomitant transport and phosphorylation of a number of sugars. The PTS system has been reviewed extensively (13-15) and is not discussed here, particularly insofar as the phosphoproteins formed are in their function similar to the transiently phosphorylated enzymes that catalyze phosphotransfer reactions and hence are beyond the scope of this review. Reports, however, on the ATP-dependent phosphorylation of one of the components of the PTS transport system by a protein kinase, stimulated by fructose 1,6-diphosphate, are relevant and are discussed in Section II,B,3. Definitive proof for the occurrence of protein phosphorylation and protein kinases in coliform bacteria was obtained in 1978 when Wang and Koshland (16) found that when Salmonella typhimurium was pulse-labeled with 32Piand SDSsolubilized bacterial contents were analyzed by SDS-polyacrylamide gel electrophoiesis, at least 4 phosphate-labeled proteins were found; they had molecular weights of 88,000, 53,000,48,000 and 45,000, respectively. Labeling occurred also when de novo protein synthesis was inhibited with chloramphenicol. The same 4 proteins, as well as an additional one of molecular weight 63,000 were labeled in vitro. Both phosphoserine and phosphothreonine were found after acid hydrolysis of the proteins phophorylated either in vivo or in vitro. The phosphorylating kinase was separated from at least one of the substrate proteins by column chromatography. Histone, casein, and phosvitin were not phosphorylated by the extracts from Salmonella. These preliminary findings were extended (17); at least 4 protein kinases were identified on the basis of chromatographic behavior, substrate specificity, and sensitivity to inhibitors. A 100,000 g supernatant fraction was obtained from S. typhimurium, incubated in vitro with [-p3*P]ATP and fractionated by column chromatography. Fractions were analyzed by SDS-gel electrophoresis and the 32P-labeled proteins were identified. The same supernate was further fractionated to separate the substrates and protein kinases and then, after chromatography, individual fractions were incubated with [Y-~~P]ATP. The kinases and substrates were then identified by recombination of the fractions in various permutations and specific kinases associated with the phosphorylation of specific proteins. The distinct kinases were characterized also on the basis of the differential inhibitory effects on phosphorylation of pyrophosphate, AMP, and an unidentified, heat-stable, low-molecular-weight (<5000) inhibitor, found in crude bacterial extracts. A comparison of phosphoproteins labeled in vivo and in vitro by two-dimensional gel electrophoresis showed that of 24 proteins labeled in vitro at least 10 were also labeled in vivo. Pulse-chase experiments showed that phosphorylation was reversible and that certain phosphate groups turned over more rapidly than others. Indirect evidence indicated the occurrence of at least 2 protein phosphatases in S. typhimurium. Similar observations were made by Manai and Cozzone (18)who, working with E. coli, found a number of phosphoproteins in the soluble fraction and

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distinct phosphoproteins associated with a crude ribosomal preparation and the ribosomal salt wash, respectively; phosphoserine as well as phosphothreonine were observed. Subsequently they found (19) a major, acidic ribosomal protein that contained phosphotyrosine. Between 92 and 98% of the cytoplasmic protein phosphate was in phosphoserine and the rest in phosphothreonine, whereas phosphotyrosine accounted for nearly all the ribosomal protein phosphorylation. The fact that in vitro protein phosphorylation occurred in both separated cytoplasmic and ribosomal fractions and that different amino acids were phosphorylated, suggested to the authors the occurrence of discrete protein kinases. An effect of the source of carbon on which the bacteria had been cultivated was noted, with a higher degree of phosphorylation observed in bacteria grown on succinate or acetate (relatively poor sources of carbon for most strains of E . coli) than in bacteria grown with glycerol or glucose. Enami and Ishihama (20) found that in E . coli the phase of bacterial growth (i.e., exponential, stationary, and transitional) as well as the duration of in vivo labeling affected the pattern of phosphoproteins quantitatively and qualitatively. Bacterial extracts obtained from late exponential phase cultures labeled for two hours contained, for example, at least 27 phosphoproteins; a total of more than 40 species of phosphoproteins were found in E . coli. Two of the phosphoproteins identified after prolonged in vivo labeling were the p- and p’-subunits of the RNA polymerase. The authors (20) purified the protein kinase(s) on the basis of the in vitro phosphorylation by bacterial extracts; 90,000- and 100,000-dalton extract proteins were used as in vitro substrates. It appears that the 100,000-dalton protein is itself a protein kinase with autophosphorylating activity; the self-phosphorylating activity was dependent on Mn2+; [y-32P]GTPalso served as phosphate donor. The identity of the 100,000-dalton protein has not been established. A second protein kinase, isolated on the basis of its ability to catalyze the phosphorylation of a 90,000-dalton E . coli protein, has a molecular weight of 120,000 and is either a heterodimer of polypeptides of 66,000 and 61,000 daltons, respectively, or a homodimer of one of these polypeptides. The 90,000-dalton substrate protein, despite certain similarities in electrophoretic behavior, does not seem to be the a-subunit of RNA polymerase, insofar as it neither bound to the polymerase core nor cross-reacted with antibody against the a-subunit. Nonetheless, preliminary findings, cited by Enami and Ishihama, suggest that the a-subunit of the E . coli polymerase may in fact be phosphorylated. This observation and evidence for the phosphorylation of the p- and P’-subunits of the RNA polymerase (see Section II,A) in noninfected as well as in phage-infected bacteria suggests that phosphorylation of proteins in E. coli may be of regulatory significance. An earlier finding discussed in the following section demonstrates that this is indeed the case in one specific instance.

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2. tsocitrate Dehydrogenase Garnak and Reeves ( 2 1 ) found that an up to 80% decrease in NADP+-linked isocitrate dehydrogenase activity, which occurs in E. coli when the bacteria adapt.to the utilization of acetate as sole source of carbon, was paralleled by the phosphorylation of the enzyme. [During growth on acetate the tricarboxylic acid cycle and the glyoxylate bypass must function simultaneously; the inhibition of the activity of isocitrate dehydrogenase permits the preferential utilization of isocitrate via the glyoxylate cycle ( 2 2 ) . ]Garnak and Reeves labeled a culture of E . coli with 32 Pi after the addition of acetate to the medium from which glycerol or glucose had been exhausted. Three labeled proteins were found in the extracts including one of apparent subunit molecular weight 5 1,000 which was identical with isocitrate dehydrogenase by a number of criteria. Acid hydrolysis of purified isocitrate dehydrogenase yielded phosphoserine as the major phosphoamino acid. The amino acid sequence of the phosphorylation site of the E. coli isocitrate dehydrogenase is -Ser(P)-Leu-Asn-Val-Leu-Arg (23).The activity of the isocitrate dehydrogenase of S. typhimurium is also regulated by reversible phosphorylation (24). LaPorte and Koshland (25) made the very interesting observation that in E. coli the isocitrate dehydrogenase kinase and phosphatase activities are associated with the same protein. The enzyme was purified by ammonium sulfate fractionation, chromatography on DEAE-Sephacel and affinity chromatography on isocitrate dehydrogenase immobilized on Sepharose. The molecular weight of the isocitrate dehydrogenase kinase, as deduced from its mobility upon SDS-polyacrylamide gel electrophoresis, was 66,000. The isocitrate dehydrogenase-phosphatase activity could not be resolved from the kinase activity by the ion exchange, gel filtration, and affinity column purification procedures. A mutant of E . coli defective in kinase activity was simultaneously devoid of phosphatase activity and the gene encoding the enzyme maps in the glyoxylate bypass operon; the gene which codes for the isocitrate dehydrogenase kinase-phosphatase has been cloned (26). It appears then that isocitrate dehydrogenase kinase-phosphatase, a key enzyme in the regulation of the flow of carbon through the tricarboxylic acid cycle and the glyoxylate bypass, is regulated at both the level of its synthesis and of its activity. The reversible phosphorylation of isocitrate dehydrogenase as an example of “zero-order ultrasensitivity,” when the substrate saturates the converter enzyme, is discussed by LaPorte et al. (27). The relationship, if any, between the isocitrate dehydrogenase kinase and the protein kinase purified by Enami and Ishihama (20) from E . coli remains to be explored.

3 . Transport of Sugars

A protein kinase that may play a role in linking the metabolism to the transport of sugars has been isolated from Streptococcus pyogenes (28, 29) and from

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Streptococcus faecalis (30). This kinase catalyzes the ATP-dependent phosphorylation of a serine residue of HPr; HPr is one of the components of the bacterial phosphoenolpyruvate-sugar phosphotransferase system (15). The phosphoryl carrier function of HPr involves its transient phosphorylation at the N-1 position of a single histidyl residue and is catalyzed by enzyme 1 of the phosphotransferase system with phosphoenolpyruvate as the phosphoryl donor. It appears furthermore that the phosphoenolpyruvate-dependentphosphorylation of the histidyl residue and the ATP-dependent phosphorylation of the seryl residue interfere with each other. Both the S . pyogenes and the S.faecalis ATP protein kinases catalyze the phosphorylation of HPrs of several other grampositive bacteria, but not of E . coli. The kinase obtained from S. pyogenes has a molecular weight of about 60,000 and that of S. faecalis of 65,000. Both enzymes appear to be natively membranal but are readily solubilized, in the case of S. pyogenes by 1.5 M MgCl,. The enzymes require Mn2 or Mg2 for activity and are inhibited by high salt concentrations. Furthermore their activities appear to be regulated by inhibition by phosphate and stimulation by fructose 1,6diphosphate, a potent allosteric activator of the kinases. The intracellular concentration of phosphate in bacteria is about 50 mM and, in the case of Streptococcus lactis drops to 4 mM when nongrowing bacteria are exposed to glucose (28); the concentration of fructose 1,6-diphosphate increases under these conditions. It appears possible then that fructose 1,6-diphosphate, by its stimulation of the ATP-dependent phosphorylation of HPr and the consequent inhibition of the phosphoryl transfer function of this key component of the phosphoenolpyruvatesugar transferase system, acts as a feedback inhibitor of sugar transport. +

+

4. Bacterial Photosynthesis A role of protein phosphorylation in the regulation of bacterial photosynthesis is indicated by experiments with the extreme halophile, Halobacterium halobium, which employs retinal-containing protein pigments for the electrogenic transport of monovalent protons across the membrane, and R . rubrum, a nonsulfur purple bacterium in which bacteriochlorophyll serves as light-harvesting pigment. Spudich and Stoeckenius (30a) and Spudich and Spudich (31) found several phosphoproteins in H . halobium. Three of these phosphoproteins of molecular weights of 110,000, 83,000, and 62,000 were rapidly dephosphorylated when the bacteria were illuminated and rephosphorylated in the dark. The first of these proteins is membranal, the other two are soluble. The amino acids phosphorylated appear to be serine and/or threonine for the 110,000- and 83,000-dalton proteins; the identity of the phosphoamino acids in the 62,000dalton protein was not reported. Inhibitor studies ( M a )indicated that one of three known retinal pigments mediated the light-induced dephosphorylation of the proteins. Spudich and Spudich (31) have shown that bacteriorhodopsin mediates the photosensitivity of the three phosphoproteins and that the light-induced ejec-

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tion of protons, effected by bacteriorhodopsin, brought about the dephosphorylation. That the dephosphorylation of the three Halobacterium proteins was indeed regulated by the proton motive force was shown by the following experiments. The proton ionophore CCCP blocked the light-induced dephosphorylation and, if added after illumination, reversed the effect of light by inducing rephosphorylation. Furthermore, a “pH jump” experiment (32), in which sudden acidification of the cell suspension caused a transient increase in the proton motive force, led to the same conclusion. The cellular concentration of ATP, affected by illumination, did not appear to play a regulatory role in the light-sensitive phosphorylation of the proteins. The identity of neither the phosphoproteins nor of the kinase and phosphatase appears to be known. The authors point out, however, that there is an analogy between the light-induced and bacteriorhodopsin-mediated protein dephosphorylation described here and the control of the phosphorylation of chloroplast lightharvesting pigment-protein complex polypeptides by the redox state of plastoquinone. The phosphorylation of these polypeptides has been proposed as a regulatory mechanism by which photosynthetic systems adapt to changing wavelengths of light (33). Protein phosphorylation in R. rubrum was studied both in virro (34) and in vivo (35).The incubation of cell-free extracts of bacteria, grown anaerobically in the light, with [ Y - ~ ~ P ] A Tgave P rise to at least six phosphoproteins; three of these were found in the intracytoplasmic membrane vesicles and two in the soluble fraction when phosphorylation was carried out in the resolved fractions; the sixth phosphoprotein apparently was not phosphorylated in the resolved fractions. Phosphoserine, phosphothreonine, and phosphotyrosine were found in the ratio of 0.32:O.16:0.52 in the membrane vesicle phosphoproteins and in the ratio of 0.66:0.25:0.09 in the phosphorylated soluble fraction. The phosphoryl moieties turned over rapidly; CAMP had no effect on phosphorylation; casein, histones, and phosvitin were not phosphorylated. Interestingly, one of the phosphoproteins, of a molecular weight of 13,000 appeared to be identical (overlapping in HPLC analysis) with the chlorophyll antenna apoprotein of the R. rubrum chromatophores. In the in vivo studies (35)with R. rubrum, grown anaerobically in the light, phosphotyrosine was the major, if not exclusive, phosphoamino acid found in the proteins of the cell-free extract, the membranes, and the soluble fraction. When cell-free extracts were incubated with [Y-~~PIATP, incorporation occurred into endogenous substrates as well as into three synthetic peptides and Val5 angiotensin 11, all containing tyrosine as the only potential phosphate acceptor. The nature of the apparent discrepancy between the in vitro findings, where serine and threonine as well as tyrosine residues were phosphorylated, and the in vivo observations, where phosphorylation was largely confined to tyrosyl residues, is not clear. The possibility that the extent of the phosphorylation of the light-harvesting

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protein 1 of R . rubrum plays a regulatory role in photosynthesis is suggested by the finding of increased phosphorylation of the protein under conditions leading to cooperativity among the photosynthetic units (i.e., in the light and in the presence of Mg2+) (36). ON THE OCCURRENCE AND FUNCTION OF C. CONCLUSIONS PROTEINPHOSPHORYLATION IN PROKARYOTES

Evidently, the conclusion that protein phosphorylation did not occur in prokaryotes was premature and based on the observation that bacterial protein kinases apparently do not catalyze the phosphorylation of exogenous proteins such as histones, casein, phosvitin, etc., which are employed traditionally in the assay of eukaryotic protein kinases. This restricted substrate specificity of prokaryotic protein kinases is striking, but how general this phenomenon is, is not clear. The kinase encoded by the coliphage T7, for example, catalyzes the phosphorylation of egg-white lysozyme, histone H2A, and protamine (37). Few generalizations can be made at this point about either the functions or the regulation of prokaryotic protein kinases. There are several exceptions to this negative conclusion; the phosphorylation of the isocitrate dehydrogenase of E . coli and S . typhimuriurn plays a major role in the regulation of carbohydrate and energy metabolism. Furthermore, both the synthesis and the activity of the kinase that catalyzes the phosphorylation of the isocitrate dehydrogenase are controlled by intermediates of carbohydrate metabolism. Other examples where a regulatory, metabolic function seems likely, if not proven, are the phosphorylation of HPr, one of the components of the bacterial phosphoenolpyruvate-sugar phosphotransferase system, and the light-induced phosphorylation and dephosphorylation of proteins in two different genera of photosynthetic bacteria. The examples cited bear on a role of protein phosphorylation in the regulation of bacterial carbohydrate and energy metabolism; a similar role of the phosphorylation of enzymes catalyzing reactions of carbohydrate metabolism was first demonstrated in higher eukaryotes. Despite the occurrence of the phosphorylation of the RNA polymerase, there is no direct evidence that the phosphorylation of proteins regulates protein synthesis in prokaryotes. Cyclic AMP, by a mechanism independent of protein kinases, and involving a CAMP-binding, DNA-binding protein controls the transcription of some genes in certain bacteria [see Ref. (38) for review]. Furthermore, there is no conclusive evidence for the occurrence of CAMP-dependent protein kinases in bacteria [see, however Ref. (11)]. By contrast, in higher eukaryotes protein phosphorylation plays a central role in the regulation of protein synthesis at the level of transcription as well as posttranscription. The juxtaposition of these two facts might lead one to the conclusion that there is here a fundamental difference between eukaryotes and prokaryotes insofar as the

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synthesis of certain proteins is regulated by kinase-catalyzed phosphorylation of proteins in eukaryotes but not in prokaryotes. Such a conclusion, however, would be premature for at least two reasons. On the one hand, the number of species of prokaryotes examined for the occurrence of a CAMP-dependent protein kinase is quite small and hence evidence is at best fragmentary. On the other hand, the fact that in both uninfected and phage-infected bacteria one or more subunits of the RNA polymerase are susceptible to phosphorylation in vivo as well as in vitro is at least compatible with a role of protein phosphorylation in the regulation of transcription, irrespective of the mechanism by which the relevant kinase is controlled. It is evident that further studies, in breadth as well as in depth, are essential if one is to arrive at an understanding of the role of protein phosphorylation in the regulation of prokaryotic protein synthesis. In this context, it would seem that two directions might be particularly rewarding. One would be an investigation of a possible occurrence of protein phosphorylation during transitions (such as in heat shock) or in development (such as during sporulation in bacilli), and the other, of relevance to our understnading of the evolution of regulatory systems, would be a quest for CAMP-binding proteins, CAMP-dependent protein kinases, tyrosine kinases, and Ca2 -regulated kinases in the Archaebacteria, as well as in hitherto unexplored genera of Eubacteria. +

111.

Protein Phosphorylation in Single-Celled Eukaryotes

A.

INTRODUCTION

The literature on the occurrence of protein phosphorylation in the single-celled eukaryotes, particularly in the fungi, is voluminous and this survey is meant to be illustrative rather than exhaustive. The emphasis is on function, or possible function, of protein phosphorylation in the individual, free-living cell. In the multicellular organism the phosphorylation of proteins, catalyzed by a variety of kinases, is the physical basis for the transmission of signals between the environment and the biosynthetic as well as the replicative machinery of the cell. The environment of the individual cell in the metazoon is constituted by other cells and the plasma and hence the external signal is a hormone or growth factor. Hormones and growth factors that act at the cell surface frequently exert their effects by initiating cascades of protein phosphorylation. In certain cases the cell surface receptor itself is a protein kinase, particularly a tyrosine kinase; in other cases occupancy of the receptor by a hormone stimulates the synthesis of a second messenger which then activates a protein kinase. In single-celled organisms the primary stimulus for protein phosphorylation is clearly intracellular and in many cases relates to the nutritional state of the cell.

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Interestingly, however, there exist intrinsically, single-celled eukaryotes, of which the cellular slime molds are the most intensively studied example, where the phosphorylation of proteins does occur in response to extracellular signals. Typically, compounds such as cAMP and folic acid, which in the case of the cellular slime molds act as external signals and cause the phosphorylation of cellular proteins, are also intermediates or byproducts of metabolism. The protein kinases of the single-celled eukaryotes resemble, at least superficially, those observed in multicellular organisms; there are kinases that catalyze the phosphorylation of acidic proteins such as casein and other kinases which act on basic proteins such as histones and protamines. Cyclic AMP-dependent protein kinases have been found in all single-celled eukaryotes examined and there is evidence for the presence of tyrosine kinases. One would predict the occurrence of protein kinase C and of calmodulin-activated kinases also, although to our knowledge such have not been reported. The functions of the phosphorylation of proteins in single-celled eukaryotes have been identified in a few cases, and in the following sections we concentrate on those organisms where a physiological role can be assigned to protein phosphorylation. We deal in large measure with CAMP-dependent protein kinases since more is known about this class of enzymes in the lower eukaryotes than about the other protein kinases; the reason for this is that mutants defective in one or another component of the cAMP system have been isolated and, furthermore, frequently the cellular content of CAMP, and hence the activities of CAMP-dependent protein kinases, is controlled metabolically and can be manipulated by the experimenter. We consider first the ascomycetous yeasts since more is known about the physiological role of cAMP in yeasts than in other fungi.

B. YEAST 1.

Protein Kinases

Takai et al. (39) purified a CAMP-dependent protein kinase about 100-fold from Saccharomyces cerevisiae. The enzyme catalyzed the phosphorylation of calf thymus histone and salmon sperm protamine. The apparent K,,, for cAMP M. Seryl and, to a lesser extent, threonyl residues were phoswas 2 X phorylated. Catalytic and regulatory subunits were separated by electrofocusing; interestingly, the catalytic and regulatory subunits of yeast and rat liver CAMPdependent protein kinase were cross-reactive in the sense that the regulatory subunit obtained from one organism inhibited the catalytic subunit of the CAMPdependent protein kinases from the other organism in a manner reversible by CAMP.In retrospect it appears that the yeast subunits, as isolated by the authors, had undergone partial proteolytic degradation. Hixson and Krebs (40)purified the regulatory subunit of the CAMP-dependent protein kinase of S. cerevisiae to homogeneity. The molecular weight of the native dimer was 95,000 and that of the monomer, as determined by SDS-gel

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electrophoresis, was 50,000, indicating a dimer of presumably identical subunits. This work confirmed the finding (39) that the protein obtained from yeast inhibited in a manner reversible by cAMP the activity of the catalytic subunit of purified mammalian CAMP-dependent protein kinases as well as of partially purified homolgous, yeast catalytic subunit. Evidently the domains required for the interaction between the regulatory and catalytic subunits have been conserved over a considerable evolutionary distance. Antibody to yeast regulatory subunit, however, did not cross-react with either type I or type I1 mammalian regulatory subunit. The yeast regulatory subunit, like the mammalian R,, subunit is subject to autophosphorylation (40). Sy and Roselle (41) found that the regulatory subunit of the CAMP-dependent protein kinase of the yeast Kluyveromyces fragilis was multiply phosphorylated and that the phosphorylations were catalyzed by the catalytic subunit. Three distinct sites of phosphorylation were identified in contrast to the one site phosphorylated in the mammalian type I1 regulatory subunit. Evidence was presented suggesting that the extent of phosphorylation of the yeast regulatory subunit varied with the metabolic state of the cells. The authors also point out (42) that the regulatory subunits of the CAMP-dependent protein kinases of different species of yeast may fall into two major size classes, with one having a molecular weight of approximately 50,000 and the other about 60,000. Other investigators reported on the occurrence of CAMP-dependent and CAMP-independent protein kinases in yeast. Gasior and his collaborators (43, 44) found several protein kinases in S. cerevisiae, including one which was associated with the ribosomes and catalyzed the phosphorylation of ribosomal proteins; it was not stimulated by CAMP. A CAMP-dependent protein kinase was found in the 150,000 g supernate and catalyzed the phosphorylation of histones. Becker-Ursic and Davies (45) also isolated protein kinases from S. cerevisiae ribosomes and found that a large number of ribosomal proteins were phosphorylated in vivo as well as in vitro; in some cases the identity of the proteins phosphorylated in vitro with those phosphorylated in vivo was established. Lerch et al. (46) purified from S. cerevisiae a CAMP-independent protein kinase about 17,000-fold; the monomer had a molecular weight of approximately 42,000 and catalyzed the phosphorylation of acidic proteins such as phosvitin and caseins, but not of protamine or histones; no natural substrates were identified. There is preliminary evidence (47) for the occurrence of phosphotyrosine in both exponentially growing and stationary phase S. cerevisiae; a membrane preparation catalyzed the phosphorylation of tyrosine residues of endogenous yeast proteins and of added casein. 2 . Cyclic AMP-Dependent Protein Kinase in Mitosis and Meiosis Studies in the laboratory of Matsumoto and his colleagues (48, 49) with mutants of S. cerevisiae, selected on the basis of a requirement for cAMP for growth, led to insights into the physiological functions of cAMP and CAMP-

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dependent phosphorylation in yeast. The relevant mutants fell into three classes: one class was defective in adenylate cyclase activity (48);in a second class the regulatory subunit of the CAMP-dependent protein kinase had a K, value for cAMP that was 10-fold higher than that of the wild type and there was an alteration in the isoelectric points of the enzyme (49); a third class led to the phenotypic suppression of the consequences of both the adenylate cyclase defect and of the abnormality of the regulatory subunit with reduced affinity for cAMP (i.e., the double mutants grew in the absence of CAMP). This third mutant had less than one-tenth of the normal regulatory subunit activity; the CAMP-dependent protein kinase had been converted into a CAMP-independent protein kinase (48). The unsuppressed mutants were arrested in the G I phase of growth in a nutrient medium, indicating a role for the CAMP-dependent protein kinase in the transition from G I to S , (48).By contrast, the initiation of meiosis in S. cerevisiae diploids, triggered normally by nutritional limitation, appears to be inhibited by cAMP and an active CAMP-dependent protein kinase insofar as the adenylate cyclase-negative mutant and the mutant having a CAMP-dependent protein kinase regulatory subunit with low affinity for cAMP initiated meiosis even in nutrient media (50). In brief then, it appears that in S. cerevisiae, an active CAMP-dependent protein kinase, is required for mitosis and growth whereas entry into meiosis occurs when the CAMP-dependent protein kinase is inactive. A significant number of yeast mutants, defective in the initiation of the cell cycle and in the entry into meiosis, have been isolated and it may well be that their analysis will lead to the identification of substrates of the protein kinase and the function of such substrates in mitosis and meiosis, not only in yeast, but in eukaryotes in general. 3.

Cyclic AMP-Dependent Protein Kinase in the Regulation of Enzyme Synthesis

Studies (51) with the same adenylate cyclase and CAMP-dependent protein kinase regulatory subunit mutants described in the previous section indicate that an active CAMP-dependent protein kinase is required for the synthesis of the repressible acid phosphatase in S. cerevisiae. Nothing is known about the underlying mechanism; the authors speculate that the phosphorylation of a chromosomal protein may be involved. There is evidence for the phosphorylation of yeast RNA polymerases I, 11 and 111. Bell et al. (52) found that several subunits of the three polymerases, including the shared 24,000-molecular-weight subunit, were phosphorylated on seryl and threonyl residues, when S. cerevisiae was grown in the presence of radioactive phosphate. The phosphorylation of the same subnits was catalyzed in vitro by a cyclic AMP-independent protein kinase which copurified with polymerase I through several steps of purification. No effect of in vitro phosphorylation on the activities of the RNA polymerases was detected

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when yeast DNA was employed as template in an in vitro transcription assay; however, as the authors point out, their assay may not have been sufficiently incisive to detect a possible effect of phosphorylation on transcription.

4. The Regulation of Enzyme Activity by Phosphorylation There is now evidence for the regulation of the activities of a number of yeast enzymes by phosphorylation-dephosphorylation.One of the intensively studied examples is that of trehalase. Trehalose (a-D-glucopyranosyl-a-D-glucopyranoside) is a major storage compound in vegetative cells and spores of fungi and the regulation of its metabolism has been reviewed (53).In general the disaccharide is formed under suboptimal conditions of growth and broken down during the reinitiation of intensive growth or the germination of fungal spores. The hydrolysis of trehalose is catalyzed by trehalases which fall into two general groups. Certain fungi have heat-stable trehalases with an acid pH optimum; these trehalases are not subject to regulation by phosphorylation. Other fungi, including S. cerevisiae, have trehalases with low heat stability and a neutral pH optimum; these latter trehalases are activated by CAMP-dependent phosphorylation. Strong evidence that in S. cerevisiae the conversion of inactive trehalase to the active form of the enzyme is brought about by phosphorylation, catalyzed by a CAMP-dependent protein kinase, was obtained by Uno et al. (54) who employed the mutants, abnormal in the adenylate cyclase and in the regulatory subunit, respectively, of the CAMP-dependent protein kinase, described earlier. Briefly, trehalase activity in crude extracts prepared from wild-type yeasts was activated by preincubation with cAMP and ATP; if ATP was replaced with the nonhydrolyzable ATP analog AMP-PNP (imido ATP), activation did not occur. Trehalase activity in a mutant with a defective CAMP-dependent protein kinase regulatory subunit (kinase active even in absence of CAMP) was higher than in the wild-type strain and not activated further by preincubation with cAMP and ATP. A double mutant with high levels of adenylate cyclase and defective cAMP phosphodiesterase, and hence high levels of CAMP, also had a high level of trehalase activity. In experiments designed to demonstrate the in vitro activation of trehalase, inactive trehalase was separated from the CAMP-dependent protein kinase by column chromatography and active trehalase was obtained after recombination of the fractions and incubation with cAMP and ATP. The K , value for cAMP in the phosphorylation of inactive trehalase was 2 x lo-* M . Activated trehalase was inactivated by treatment with alkaline phosphatase. Another example of a yeast enzyme activated by phosphorylation is that of 6phosphofructo-2-kinase which catalyzes the synthesis of fructose 2,6-biphosphate, a positive effector of 6-phosphofructo-1-kinase, from fructose 6-phosphate and which is activated by a CAMP-dependent protein kinase (55). The phosphorylated enzyme catalyzes the reaction with a 2-fold lower K,,, for fruct-

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ose 6-phosphate and a 4.3-fold higher V,,, than the unphosphorylated enzyme. Interestingly, the situation in yeast is the converse of that in liver where phosphorylation of the 6-phosphofructo-2-kinaseleads to inactivation of the enzyme (56). Gluconeogenesis in yeast appears to be controlled by the phosphorylation of several enzymes. The addition of glucose or metabolically related sugars to yeast growing on acetate or ethanol causes the time-dependent loss of activity of gluconeogenic enzymes, including fructose- 1,6-biphosphatase, phosphoenolpyruvate carboxykinase, and cytoplasmic malate dehydrogenase (57, 58). There is evidence, at least in the case of fructose-l,6-biphosphatase,for a biphasic mechanism of inactivation; the reversible inactivation by phosphorylation precedes irreversible degradation by proteolysis. The addition of glucose and metabolically related sugars to cultures of yeast growing on ethanol or acetate brings about a transient increase in CAMPand there is evidence that a CAMP-dependent protein kinase catalyzes the phosphorylation of fructose- 1,6-biphosphatase. Gancedo et al. (59) found that the incubation of purified fructose-l,6-biphosphatase with bovine heart CAMP-dependent protein kinase led to a 50% loss of fructose- 1,6-biphosphatase activity; the phosphorylation of the enzyme was greatly stimulated by fructose 2,6-biphosphate with a maximal effect at 5 phi fructose 2,6-biphosphate. The rate of phosphorylation in either the absence or presence of fructose 2,6-biphosphate closely paralleled the rate of inactivation of fructose- 1,6-biphosphatase activity. These in vitro observations are similar to findings made in vivo where the addition of glucose to a yeast culture also brought about an approximately 50% loss of activity and where the stoichiometry of the phosphorylation was the same as in the in vitro experiments (60).Furthermore, the addition of glucose to the yeasts in culture brought about a sharp increase in the cellular concentration of fructose 2,6-biphosphate. Experiments (61) in which the mutants, defective in adenylate cyclase and in the regulatory subunit of the CAMP-dependent protein kinase, were employed led to the conclusion that the CAMP-dependent protein kinase catalyzed the inactivating phosphorylation of fructose- 1,6-biphosphatase. The role of the CAMP-dependent protein kinase in the phosphorylation of fructose-1,6-biphosphatase was demonstrated also by Pohlig et al. (62) who found, in addition to stimulation of the phosphorylation by fructose 2,6-biphosphate, an inhibitory effect by 1 mM 5'-AMP. Recent experiments (63) in which purified yeast CAMP-dependent protein kinase was employed for the phosphorylation of purified yeast fructose- 1,6-biphosphatase showed that maximal phosphorylation was accompanied by 60% loss of enzyme activity. Serine was the only amino acid phosphorylated. The unphosphorylated and phosphorylated fructose- 1,6biphosphatases differed in their pH optima and the ratios of activities in the presence of Mg2+ and Mn2+, respectively. The affinities for fructose 2,6biphosphate and 5'-AMP were not affected; the two compounds stimulated the

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phosphorylation of fructose- 1,6-biphosphatase and inhibited the activity of the gluconeogenic enzyme. The inactivation of another enzyme by phosphorylation has been studied in some detail. Hemmings (64, 65) found that starvation of the yeast, Candida utilis, for glutamate, which brings about the inactivation of the catabolic, NADdependent glutamate dehydrogenase, was paralleled by the phosphorylation of the enzyme. The enzyme has a molecular weight of approximately 460,000 and consists of four identical subunits of a molecular weight of approximately 116,000. The two forms of the enzyme were purified to homogeneity; phosphorylation occurred on a serine residue and approximately I mol of phosphate was incorporated per mol subunit. The phosphorylated enzyme had 8- to 10-fold less activity than the unphosphorylated species, a higher K,,,for glutamate, and a different pH optimum. The phosphorylated enzyme was dephosphorylated and reactivated by incubation with a crude yeast extract and the phosphoprotein phosphatase which catalyzed the dephosphorylation was isolated-the first isolation of a phosphoprotein phosphatase from a simple eukaryote (66). The phosphoprotein phosphatase had a molecular weight of approximately 40,000 and catalyzed the dephosphorylation of phosphocasein, phosphohistone, and (more effectively) phosphokemptide. Its preference for phosphokemptide and ability to dephosphorylate the phospho form of glutamate dehydrogenase suggests that the phosphorylation of glutamate dehydrogenase may be catalyzed by a CAMPdependent protein kinase. Limited proteolysis with trypsin of the inactive phosphoglutamate dehydrogenase yielded fragments of molecular weights of 64,500 and 48,000 and activation of the enzyme, followed by loss of activity. Only the 48,000-dalton fragment was labeled with 32P. Uno et al. (67) working with S. cerevisiae, including the mutants, defective in adenylate cyclase and in the regulatory subunit of the CAMP-dependent protein kinase, found that in vitro the phosphorylation and inactivation of the NAD-dependent glutamate dehydrogenase could be catalyzed by both CAMP-dependent and CAMP-independent protein kinase.

C . NEUROSPORA The search for protein kinases in Neurospora was stimulated by the observation that adenylate cyclase-negative mutants of the ascomycete were abnormal in the control of elongation and in the formation of conidia (68, 69) and the assumption that the effect of cAMP might be mediated by a CAMP-dependent protein kinase. Judewicz and Torres (70) found that cAMP increased the phosphorylation of chromatinal nonhistone proteins and that phosphorylated chromatin preparations were more effective than nonphosphorylated chromatin as templates for a-amanitin-sensitive transcription. Three protein kinase activities were observed in 105,000 g supernatants of Neurospora crassa (71).One of the

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kinases was CAMP-dependent and inhibitible by an endogenous heat-stable inhibitor; the other two kinases were CAMP-independent. Powers and Pall (72) found two protein kinases in N. crassa; one was CAMP-dependent, the other CAMP-independent. The CAMP-dependent protein kinase was inhibitible by the bovine heat-stable protein inhibitor, specific for the catalytic subunit of CAMPdependent protein kinases (73); histone H2B was the best substrate for the CAMP-dependent protein kinase. The regulatory subunit of the CAMP-dependent protein kinase of Neurospora crassa was purified 2000-fold; it was a dimer with an estimated subunit molecular weight of 47,000. The Kd for cAMP was 300 nM and there were two CAMP-binding sites per monomer; 8-bromo cAMP bound selectively to the site from which cAMP dissociated slowly as is the case for the mammalian subunits. The regulatory subunit of the Neurospora CAMP-dependent protein kinase was quite similar to the mammalian protein kinase R, regulatory subunit as well as to the regulatory subunit of the yeast CAMP-dependent protein kinase; as in the case of yeast and Dictyostelium (see Section IILG), there is evidence for the occurrence of only one type of regulatory subunit in each species (74).

D. Mucof? 1 . Protein Kinases and Phosphoprotein Phosphatases

Protein kinases have been studied also in certain coenocytic, dimorphic fungi such as Mucor and Blastocladiella for much the same reason as in yeast (i.e., it appears that cyclic nucleotides play a role, causal or otherwise, in the morphological transitions and it is assumed that protein kinases mediate the effects). Certain species of Mucor, a phycomycete, are dimorphic; that is, depending upon environmental conditions, their growth is either yeast-like, with spherical cells which reproduce by budding, or filamentous (75, 76). A possible involvement of cAMP in the transition between the two types of growth was suggested by the finding that a 4-fold decrease in the concentration of cellular cAMP preceded the transition from the yeast-like to the mycelial form of growth (77) and the observation that the addition of cAMP to Mucor rouxii spores led to the formation of budding spherical cells instead of filaments (78). The CAMP-dependent protein kinases of M . rouxii has been studied intensively by S. Passeron and her collaborators (79, SO). It appears that the CAMPdependent protein kinase of M . rouxii has a molecular weight of 242,000 and that it has the tetrameric structure R,C,. The catalytic subunits have a molecular weight of approximately 41,000 while the regulatory subunits have the unusually high molecular weight of 75,000; frictional coefficients of 1.55 and 1.62 for the holoenzyme and the regulatory dimer, respectively, were determined (81).The Mucor CAMP-dependent protein kinase is unusual in not being physically dissociated by cAMP alone, although cAMP fully activates the enzyme; a combina-

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tion of cAMP with either histone, protamine, or NaCl is required for the physical dissociation of the enzyme (79). The fact that cAMP binds to the CAMP-dependent protein kinase holoenzyme without dissociating it, permitted the demonstration that twice as much cAMP binds to the dissociated subunits as to the holoenzyme (82); possibly the catalytic subunits shield one type of CAMP-binding site. Two phosphoprotein phosphatases have been isolated from Mucor rouxii (83). The two enzymes are found in the soluble fraction of the mycelium, and differ in their molecular weights, divalent cation requirements, and substrate specificities; both enzymes are strongly inhibited by inorganic phosphate. Phosphoprotein phosphatase I has a molecular weight of 64,000, a sedimentation coefficient of 3.2 S, depends on the presence of divalent cations for activity, and is slightly inhibited by NaF and ATP; its preferred test substrate is P-histone. Phosphoprotein phosphatase-2 catalyzes the dephosphorylation of mammalian phosphorylase a; its molecular weight is 43,000 and sedimentation coefficient 3.0 S; its activity does not require exogenous divalent metal ions and it is strongly inhibited by NaF and ATP. Two heat-stable inhibitors of the phosphoprotein phosphatases have also been isolated from M . rouxii (84). 2. Regulation of Enzyme Activity by Phosphorylation in Mucor The trehalase of M . rouxii can be activated in vitro by CAMP-dependent phosphorylation, much as the yeast trehalase is activated (see Section III,B,4), and there is evidence that the activity of the Mucor trehalase is regulated by phosphorylation-dephosphorylation also in vivo (85). It appears that the activity of the cAMP phosphodiesterase of Mucor rouxii is regulated by phosphorylation-dephosphorylation (86, 87); two forms of low K , cAMP phosphodiesterase were resolved from mycelial extracts of M . rouxii; one form, PDE I, could be activated either reversibly by phosphorylation or, irreversibly, by proteolysis; the other form, PDE 11, was unresponsive to activation. Endogenous proteolysis or controlled treatment with trypsin converted PDE I to PDE 11. Either phosphorylation or proteolysis led to an approximately 15-fold increase in the V,,, of the enzyme without a significant change in the K,,,. The two forms of the cAMP phosphodiesterase were partially purified and it could be shown that the cAMP phosphodiesterase can be phosphorylated and activated by a CAMP-dependent protein kinase in vitro. Furthermore, the domain containing the phosphorylation site(s) was within the domain susceptible to proteolytic activation (88). It may well be that the reversible phosphorylation of the Mucor cAMP phosphodiesterase is of regulatory significance; it appears that PDE I is the only form of the enzyme in germlings of Mucor, a mixture of the two forms is found during logarithmic growth and PDE I1 is the only form in mycelia from the stationary phase of growth.

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E. BLASTOCLADIEUA Blastocladiella emersonii is an aquatic phycomycete with a cell cycle in the course of which sessile, multinucleate vegetative cells differentiate and cleave into motile uninucleate zoospores; the zoospores, in turn, germinate and give rise to vegetative cells [reviewed in Ref. (89)l. 1. Protein Kinases

Much as in the case of Mucor, a role of cAMP and cGMP in the transition between the morphological stages of the life cycle has been invoked (90) and hence protein kinases, particularly CAMP-dependent protein kinases, have been studied. Brochetto-Braga et al. (91) found a CAMP-dependent protein kinase in B. emersonii zoospores which is developmentally regulated insofar as its activity is low in vegetative cells and increases during sporulation (92). As in Mucor and Dictyostelium (see Section III,G), and unlike in higher eukaryotes, only one form of the CAMP-dependent protein kinase was observed in B . emersonii. The molecular weight of the holoenzyme was approximately 200,000; the catalytic subunit monomers had a molecular weight of 40,000 and the regulatory subunit monomer of 58,000. The chromatographic behavior of the holoenzyme, the effect of NaCl on the reassociation of the subunits, and the fact that the regulatory subunit was subject to autophosphorylation were similar to the behavior of the mammalian type I1 protein kinase. The phosphate group of the Blastocladiella protein kinase regulatory subunit was removed very rapidly by an endogenous phosphoprotein phosphatase in the presence of cAMP and the authors suggested (93) that the very rapid dephosphorylation of the fungal CAMP-dependent protein kinase regulatory subunit may reflect high sensitivity to in vivo regulatory signals.

2 . Phosphorylation of Ribosomal Proteins The occurrence of CAMP-independent protein kinases in B . emersonii has been studied, particularly in relation to the phosphorylation of a 40 S ribosomal subunit protein with a molecular weight of 32,000; the protein corresponds probably to the S6 mammalian ribosomal protein (94). (It will be recalled that multiple phosphorylation of S6 precedes the transition from quiescence to the early G, phase of the cell cycle in mammalian cells) (95). Zoospores of B . emersonii contain a complete, but nonoperative system for protein synthesis, which includes untranslated mRNA, stored in the nuclear cap, a membranebound structure which contains the zoospore ribosomes. After the onset of germination of the zoospores, the nuclear cap disintegrates, the ribosomes are dispersed in the cytoplasm, polysomes are formed, and protein synthesis is initiated. Bonato et al. (94) found two cyclic nucleotide-independent protein kinases that could be extracted from the nuclear cap ribosomes and catalyzed the phosphorylation of a single protein of the Blastocladiella 40 S ribosomal subunit

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(i.e., the protein of a molecular weight of 32,000). The two kinases were tightly associated with the ribosomes, but were extracted with 0.5 M KCl; they could be separated on DEAE cellulose; both catalyzed the phosphorylation of casein, but differed with respect to the optimal Mg2+ and KCl concentrations for their activities. The 32,000-dalton, ribosome-derived substrate migrated on 2-dimensional polyacrylamide gels in a manner similar to that of the mammalian ribosomal protein S6 and was found to be phosphorylated also after growth of the organism in the presence of 32Pi. Work by Bonato et al. (96)suggests that, although in vitro the Blustocladiella ernersonii S6 protein can be phosphorylated by the two ribosome-derived, CAMP-independent protein kinases as well as by the soluble, CAMP-dependent protein kinase, the latter enzyme may be primarily responsible for the phosphorylation of S6 in vivo. The phosphopeptide maps obtained after the tryptic hydrolysis of S6 phosphorylated in vitro by the two CAMP-independent protein kinases were quite different from those found after in vivo phosphorylation or after the in vitro phosphorylation catalyzed by the CAMP-dependent protein kinase. One major phosphopeptide observed after in vitro phosphorylation by the CAMP-dependent protein kinase was identical with one of the phosphopeptides found in both the zoospores and germlings of B. ernersonii. The authors suggest that, unlike in the case of vertebrates, only the CAMP-dependent protein kinase may be capable of catalyzing S6 phosphorylation in vivo (96). Germination of the zoospores in a nutrient medium was paralleled by an increase in S6 phosphorylation. After two hours of growth, the state of phosphorylation returned to the basal, zoospore level. When germination of the zoospores was induced in a nonnutrient buffer, containing K + ions as inducer, there was no increase in S6 phosphorylation; this suggests that S6 phosphorylation in this case was dependent on the presence of exogenous nutrients rather than causally related to germination per se. F. MISCELLANEOUS FUNGI The occurrence of several protein kinases in the basidiomycete Coprinus rnucrorhizus has been reported (97); one of the protein kinases was stimulated by CAMP,two were inhibited, and one was CAMP-independent. The exposure of C. rnucrorhizus to certain antibiotics such as nystatin, amphotericin B, polymixin B, and tyrocidin and to depolarizing agents led to a transient, approximately 2fold increase in endogenous CAMP, to the activation of CAMP-dependent protein kinase activity, and to the stimulation of the phosphorylation of at least three proteins (98). The occurrence of a CAMP-dependent protein kinase in the basidiomycete Ustilugo rnuydis has also been described (99);the enzyme catalyzes the phosphorylation of kemptide and protamine but apparently not of histones. Analysis by sucrose gradient sedimentation indicated an apparent mo-

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lecular weight of 72,000 for the holoenzyme and molecular weights of approximately 35,000 each for both the catalytic and regulatory subunits. Clearly more experiments with a highly purified preparation of the enzyme are required before conclusions regarding the polymeric structure of this protein kinase can be drawn.

G . SLIMEMOLDS The slime molds fall into two major groups, the syncytial “true” slime molds and the pseudoplasmodial cellular slime molds. Both groups are characterized by the occurrence of a single-celled, amoeboid, feeding, ‘‘protozoon-like” stage and a plasmodia1 or multicellular, nonfeeding, “fungus-like” stage in their life cycles. The slime molds, in view of their transitional position between uni- and multicellularity and their protozoon and fungal affinities, have been studied intensively. The most commonly investigated species of the true slime molds is Physarurn polycephalum; DNA synthesis and nuclear division are naturally synchronous in the plasmodia of this organism and hence are highly suitable for a number of relevant investigations. In this context a possible role for the phosphorylation and dephosphorylation of histone HI in the cell cycle has been mooted. The most commonly studied species of the cellular, pseudoplasmodial slime molds is Dictyostelium discoideurn. In this organism cAMP plays a dual role in development; cAMP is the extracellular, chemotactic agent which in starving amoebae brings about aggregation which precedes the formation of a fruiting body composed of moribund stalk cells and viable spores. Cyclic AMP acts also as “second messenger” in the sense that extracellular cAMP stimulates the membranal adenylate cyclase of the organism and affects the synthesis of a number of developmentally regulated proteins. It is assumed that intracellular cAMP mediates some of the effects of the extracellular, “hormone-like” CAMP. Hence many studies have focused on the functions of the CAMP-dependent protein kinase in D. discoideurn in motility and development. 1. Physarurn Polycephalum The Phosphorylation of Histone H I . There is evidence that histone H1 stabilizes the nucleosome and that it is located in the region of the entry and exit points of the DNA (100). Bradbury and his colleagues (101) showed that there was a large increase in the phosphate content of the Physarurn lysine-rich H I histone during the progression from late G , to mitosis. They measured the phosphorylating activity of nuclear extracts of P. polycephalurn isolated at defined stages of the mitotic cycle, using calf thymus histone H1 as substrate. They found that the phosphorylating activity increased steadily through the mitotic

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cycle from a minimum near metaphase to a maximum in late G, phase and then fell rapidly to a minimum near metaphase. The high phosphate content in late G , could be explained by an increase in phosphorylating activity rather than by an increase in available substrate since an excess of added calf thymus H1 histone was employed in the in vitro measurements. The authors suggested that, in fact, the phosphorylation of histone H 1 might initiate the condensation of the chromosome (102). Fischer and Laemmli (103) confirmed the finding of Bradbury et al. and observed that phosphorylation of histone HI weakened its affinity for both native and single-stranded DNA cellulose. The complexity of the process of phosphorylation of the Physarum H1 histone is underlined by the recent findings of Mueller et al. (104) who observed that at metaphase the HI molecules contained between 20 and 24 phosphates. Immediately after metaphase HI underwent rapid dephosphorylation to 9- 16 phosphates. During progression into the S phase, the newly synthesized H1 was phosphorylated and merged with the old dephosphorylated H 1. By the end of S phase all H 1 molecules underwent phosphate turnover and higher states of phosphorylation accompanied progression into G,; by prophase all of the H1 molecules contained 15-24 phosphates. The authors conclude that the phosphorylation of H1 does play an important role in the process of chromosome condensation. They point out also that, consistent with the fact that the Physarum H 1 histone has a higher molecular weight than its mammalian counterpart, there is a much larger number of phosphorylation sites in the Physarum than in the mammalian HI histone. Another recent study (103, also dealing with the phosphorylation of Physarum histone H1 during the mitotic cycle, led to the interesting observation that methylation of the E-NH, groups of several of the HI lysines preceded phosphorylation. The authors confirmed the earlier findings that the Physarum HI histone could accept more than 20 phosphates per molecule and that it was superphosphorylated in mitosis and not totally dephosphorylated in the following cycle. Most of the phosphorylation sites were located in the COOH-terminal portion of the molecule. The overall findings are consistent with the hypothesis that the newly synthesized H 1 associated with DNA prior to methylation and phosphorylation and that methylation and phosphorylation, in that order, occur in the course of the maturation of the chromatin. The authors speculate that the multiple conformational classes of Physarum HI histone may be functionally equivalent to the true sequence variants of mammalian H1 histone. Chambers et al. (106) addressed themselves to the identification of the kinases responsible for the phosphorylation of the Physarum HI histone. Upon DEAEcellulose chromatography, they found three major peaks of activity; the protein kinase activities were further characterized on the basis of their sensitivities to the heat-stable proteinaceous inhibitor of the mammalian CAMP-dependent protein kinase and the sites phosphorylated in calf and Physarum HI histones in vitro. The activities of two of the fractions were inhibited by an excess of the

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specific proteinaceous inhibitor; the activities of neither of these two fractions were stimulated by CAMP, which was not unexpected, since they were isolated from nuclei where CAMP-dependent protein kinases are in the dissociated state (107).Of the two fractions, one catalyzed the phosphorylation of primarily serine-37 in calf H1; the second fraction, possibly a mixture of kinases, catalyzed the phosphorylation of multiple sites, mainly in the N-terminal half of calf HI ; it differed from the first fraction with respect to the sites phosphorylated in Physarum H1 (106).The third fraction, unaffected by the inhibitor, catalyzed the phosphorylation of multiple sites in both the N- and C-terminal halves of calf H I . These sites included those phosphorylated by the mammalian growth-associated histone kinase and thus the third kinase may be analogous to the mammalian growth-associated kinase (108). There is evidence (109)for the occurrence in nuclei of P . polycephalum of a histidine kinase that catalyzes the transfer of phosphate from [ Y - ~ ~ P ] A TtoP histidine-75 of calf thymus histone H4 with the formation of 1-phosphohistidine. It is not known, whether phosphohistidine is found in vivo (other than as an intermediate in phosphotransfer reactions). The occurrence in P . polycephalum of a protein kinase, activated by polyamines, has been described (110); the kinase catalyzed the phosphorylation of a nonhistone, acidic nucleolar protein of 70,000 molecular weight. The substrate in its phosphorylated form apparently stimulated the transcription of DNA coding for ribosomal RNA (111);the ribosomal DNA genes in Physarum are located in the nucleolus on satellite DNA. Furthermore, it was suggested that the 70,000molecular-weight protein, in its unphosphorylated form, had ornithine decarboxylase activity and that it was inactivated by phosphorylation (112).Other work (113)casts doubt on this interpretation; the authors purified the ornithine decarboxylase of P . polycephalum considerably and found that both the active and inactive forms had a molecular weight of 52,000 and argue that the protein, apparently phosphorylated by the polyamine-dependent protein kinase, was not ornithine decarboxylase. 2 . Dictyostelium Discoideum The central role of cAMP in the development of D . discoideum was first recognized when the cyclic nucleotide was identified as the chemoattractant responsible for the transition from unicellularity to multicellularity in this cellular slime mold (114).Cyclic AMP binds to cell-surface receptors and activates a membranal adenylate cyclase; the CAMP-stimulated synthesis of cAMP then forms the basis of a relay system by which chemotactic stimuli are propagated. At the same time cAMP is required for cell differentiation. There is evidence that protein phosphorylation is involved in both processes. a. Phosphorylation of Myosin. The occurrence of the phosphorylation of myosin heavy chains in D . discoideum, regulated in response to the chemotactic

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stimulation of the amoebae by CAMP, was demonstrated by Malchow et al. (115). Myosin heavy chains were dephosphorylated in stimulated cells (116) and kinase activity, associated with an actomyosin-containing membrane preparation, was inhibited by CA2+ in the presence of endogenous calmodulin (115). Chemotactic stimulation by cAMP led to an enhanced influx of Ca2+ into the cells. It appears then that the chemotactic response of D . discoideum to cAMP during aggregation is mediated by the dephosphorylation of the myosin heavy chains and that this dephosphorylation results from by an inhibition of myosin heavy chain kinase activity by Ca2 -calmodulin. One may assume the occurrence of a highly active myosin heavy chain phosphatase. It was shown earlier that the phosphorylation of the Dictyostelium myosin heavy chains resulted in a decrease of the actin-activated MgATPase activity and inhibition of the formation of filaments (117). Maruta et al. (118) partially purified two myosin heavy chain (MHC) kinases from D . discoideum. MHC kinases I1 was found in aggregation-competent, but not in growing, amoebae; the enzyme had an apparent molecular weight of 70,000; it was inactivated by Ca-calmodulin. Nonphosphorylated myosin had a 2- to 3-fold higher MgATPase activity than did myosin maximally phosphorylated by MHC kinase 11. Threonine was the only amino acid phosphorylated in Dictyostelium myosin heavy chains and there were at least two phosphorylatable threonine residues per myosin heavy chain. The two phosphorylated sites on the myosin heavy chain were localized by the use of monoclonal antibody (119). The binding sites of the monoclonal antibodies had been determined earlier by electron microscopic mapping (120). Both phosphorylation sites were within the terminal third of the myosin tail. The authors point out that the location of the phosphorylation sites distal from the myosin heads and close to the polymerization sites suggests an effect of phosphorylation on the state of assembly of the myosin and, in fact, phosphorylated myosin is less polymerized than the nonphosphorylated molecule; the disassembled myosin may have less actin-activated ATPase activity than the assembled molecule. The heavy chains of myosin are phosphorylated also in another single-celled eukaryote, Acanthamoeba castellanii (121). A . castellanii contains at least three different myosin isoenzymes; the phosphorylation of myosin 1A and 1B stimulates actomyosin MgATPase activities whereas the phosphorylation of myosin I1 inhibits actomyosin MgATPase activity (122). By contrast, in vertebrate nonmuscle cells light chain myosin is phosphorylated by a calmodulin-dependent light chain kinase; this phosphorylation induces the assembly of myosin filaments and is correlated with enhanced actin-activated myosin MgATPase activity (123). +

b. Cyclic AMP-Dependent Protein Phosphorylation and Development. Suggestive evidence that CAMP, and hence presumably the phosphorylation of proteins, plays a central role in cellular differentiation and development in Dictyostelium came from several sources. For example, Gerisch et al. (124) found

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that the exposure of stationary phase amoebae to pulses of cAMP led to the premature synthesis of the developmentally regulated ‘‘contact sites A” required for the end-to-end concatenation of aggregating amoebae (contact sites A are constituted of a glycolipophosphoprotein with sulfated carbohydrate residues) (125). Darmon and his colleagues (126) and Juliani and Klein (127) found that certain nonaggregating mutants, blocked at an early stage of development, could be rescued phenotypically (i.e., they formed normal fruiting bodies), if exposed to pulses of CAMP. It is clear that in D . discoideum cAMP is required not only for the aggregation of the amoebae but also for normal development; the assumption is generally made that an increase in intracellular CAMP, brought about by the stimulation of the adenylate cyclase by extracellular CAMP, mediates, via a CAMP-dependent protein kinase, some of the developmental effects of the extracellular CAMP. This is a reasonable assumption, but, it should be stressed, formal proof of its correctness awaits the isolation of the appropriate mutants. The occurrence of a CAMP-dependent protein kinase (128-131) as well as of CAMP-independent protein kinases [e.g., Refs. (132, 133)] has been described. The CAMP-dependent protein kinase isolated from 100,000 g supernates of D . discoideum resembles the analogous enzyme of vertebrate origin and, as in the case of yeast (39, 40), the domains on the catalytic and regulatory subunits required for their interaction have been conserved to the extent that the Dictyostelium and bovine regulatory subunits inhibit the activity of the heterologous catalytic subunits in a mnner reversible by cAMP (134, 135). The regulatory subunit of the Dictyostelium CAMP-dependent protein kinase has been purified and antibody against it has been produced (128, 129). The Kd for cAMP is similar to that of the mammalian regulatory subunits, but, as in the case of the regulatory subunit of the yeast CAMP-dependent protein kinase (40), there appears to be only one CAMP-binding site per monomer (136) with relative affinities for several analogs of CAMP, similar to that of the mammalian regulatory subunits (131). The apparent molecular weight of the Dictyostelium regulatory subunit, as determined by SDS-gel electrophoresis, is 41,000 (128) (i.e., smaller than that of any other characterized regulatory subunits). That the 41,000-dalton entity was not a proteolytic fragment was demonstrated by two approaches; Western blots of extracts obtained by the lysis of the amoebae directly in SDS yielded the regulatory subunit having the same apparent molecular weight (129); the in vifro translation of mRNA gave rise to a protein of molecular weight 41,000 which was recognized by antibody prepared against the Dictyostelium regulatory subunit (137). It is not clear whether the apparent difference in size between the Dictyostelium and mammalian regulatory subunits results from a real difference in molecular weights or reflects an absence from the Dictyostelium regulatory subunits of the peculiar conformation which causes the mammalian regulatory subunits to migrate on SDS-gels with an apparent molecular weight of 8000 higher than the actual molecular weight as based on amino

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acid composition. The CAMP-dependent protein kinase regulatory subunits of other lower eukaryotes migrate at the higher molecular weights characteristic of the mammalian subunits. The CAMP-dependent protein kinase of Dictyostelium, like that of other lower eukaryotes, appears to have only one type of regulatory subunit, when analyzed on l-D SDS-gels; however, upon 2-D gel electrophoresis two proteins of identical molecular weight and slightly different p1 were observed consistently (128). The relationship, if any, between these two forms of the Dictyostelium regulatory subunit and the occurrence of the two regulatory subunits in the mammalian CAMP-dependent protein kinase remains to be established. Dictyostelium discoideum represents a very deep branch of eukaryotic evolution (138) and a comparison of the amino acid sequence of its regulatory subunit with those of the vertebrate CAMP-dependent protein kinase regulatory subunits as well as the catabolite gene activator (CAMP-binding) protein of the prokaryotic E . coli might well be of interest. The catalytic subunit of the Dictyostelium CAMP-dependent protein kinase has not been purified to homogeneity; its marked preference for the synthetic peptide substrate, Kemptide, its Kd for ATP, and its inhibitibility by the specific proteinaceous inhibitor (isolated from rabbit skeletal muscle) indicate strong similarity of the respective domains of the catalytic subunit with their mammalian counterparts (128, 129, 139). The subunit structure of the D . discoideum holoenzyme is a matter of debate; for example, it has been reported to be a dimer of molecular weight 82,00088,000 (129), a tetramer of molecular weight 166,000-185,000 (128), and a tetramer of molecular weight 270,000 (131). An association of, at least of some of the enzyme with the cytoskeleton has not been ruled out, but the CAMPdependent protein kinase of Dictyostelium is generally found in the high-speed supernatant fraction of extracts. The observation (139) that there was a coordinate 4- to 6-fold increase in the cellular concentration of the two subunits of the CAMP-dependent protein kinase during early development in D . discoideum, at a stage prior to the onset of prespore and prestalk cell-specific mRNA synthesis and, furthermore, that prespore cells had an approximately 5-fold higher concentration of the enzyme than did prestalk cells (140) suggests that the CAMP-dependent protein kinase plays a role in development. The increase in the activity of the regulatory subunit and, presumably, of the catalytic subunit represented de novo synthesis (139). The increase in CAMP-dependent protein kinase occurs prior to the appearance of known spore-specific proteins and the high level of CAMP-dependent protein kinase persists after the experimental disaggregation of the multicellular “slug” to a single-cell suspension, under conditions where CAMP is required for the maintenance of cell-type-specific mRNAS. These facts are also compatible with a role of the CAMP-dependent protein kinase in development (139). Finally, there is evidence for the translocation of a fraction of the two subunits from the

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cytoplasm into the nucleus; the fraction found in the nucleus increases significantly during development (141). As was found earlier in the case of mammalian cells (107), CAMP stimulates translocation into the nucleus. An analysis of nuclear substrates of the CAMP-dependent protein kinase is under way in the authors’ laboratory. Preliminary results indicate that some substrates are phosphorylated primarily at certain stages of development and that in some cases they are phosphorylated in only one or the other of the two types of cells. A very basic nuclear protein of 38,000 molecular weight occurred in both prespore and prestalk cells; it was phosphorylated in vivo as well as in vitro and the addition of purified catalytic subunit to nuclear extracts greatly enhanced its phosphorylation. c. Cyclic AMP-tndependent Protein Phosphorylation. The developmental regulation of the phosphorylation of a protein from the D . discoideum 40 S ribosomal subunit, tentatively identified as S6, has been reported (142). The protein was minimally phosphorylated in growing amoebae, became maximally phosphorylated at the time of aggregation, and returned to the low level of phosphorylation when the fruiting body was formed. The state of phosphorylation of S6 correlated with the stage of development rather than starvation since the starvation of a mutant, blocked prior to aggregation, did not lead to S6 phosphorylation. The protein kinase which catalyzed S6 phosphorylation has not been identified. Rahmsdorf and Pai (132) found four major soluble and one membrane-associated protein kinases; all these enzymes had high affinity for casein and were CAMP- and cGMP-independent. The membranal protein kinase catalyzed the phosphorylation of endogenous membrane proteins in addition to casein and histone IIA. Parish et al. (143) also noted the occurrence of a membranal protein kinase in D . discoideum and found that it catalyzed the phosphorylation of a number of membranal and soluble proteins including myosin heavy chains and discoidin [discoidins are Dictyostelium lectins; discoidin I has apparently a “fibronectin-like” function in that it promotes the attachment, spreading, and ordered migration of the amoebae during morphogenesis (244)l. They found that intact cells could utilize ATP as phosphate donor. Other workers (145) also claimed that a membranal CAMP-independent protein kinase could use extracellular ATP for the phosphorylation of membranal proteins. The possibility, however, that the buffers in which the amoebae were suspended caused the permeabilization of the cells to ATP is not excluded. Renart et al. (133) purified a nuclear, CAMP-independent protein kinase approximately 400-fold from D . discoideum. The kinase, which had an apparent molecular weight of 38,000, could utilize either ATP or GTP as phosphate donor and catalyzed the phosphorylation of casein and phosphovitin and, to a lesser degree, of histones; when casein served as substrate the phosphate was found in

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both threonine and in serine. Spermine, spermidine, and polylysine stimulated the phosphorylation of casein; spermine was the most effective and at a concentration of 2 mM stimulated severalfold; spermine was inhibitory, however, when phosphovitin was the substrate. The activity of the kinase was sensitive to inhibition by low concentrations of heparin irrespective of the substrate. The purified protein kinase was a substrate for its autophosphorylation; several endogenous, nuclear proteins were also phosphorylated by the enzyme. The occurrence of a number of cytoplasmic and membranal phosphoproteins in D. discoideum has been reported (e.g., 146-149); their phosphorylation is catalyzed in some cases by CAMP-dependent, in others by CAMP-independent protein kinases; it is not clear at present to what extent the phosphorylation of these proteins plays a direct role in development and they are therefore not further discussed. There is one case, however, where the physiological relevance of the phosphorylation of a membranal protein may be close to elucidation. ) that the affinity-labeled cell surface cAMP receptor Recent work ( 1 4 9 ~ 6shows of D. discoideum migrated as a doublet of proteins with molecular weights of 40,000 and 43,000 when submitted to SDS-gel electrophoresis. The relative intensities of the two bands of the doublet alternated with the same frequency as the spontaneous oscillations of cAMP synthesis in the cells of origin. The oscillations reflect the alternation between activity and inactivity (adaptation, refractoriness) of the cAMP receptor in resonse to extracellular CAMP. It had been observed earlier (146) that the exposure of the Dictyostelium amoebae to cAMP induced the rapid and reversible phosphorylation of a protein, provisionally identified as the cAMP receptor, and that this was paralleled by a change of the apparent molecular weight of the receptor from 45,000 to 47,000. It appears ) to the active (unlikely that the two bands of the doublet ( 1 4 9 ~correspond phosphorylated) and the inactive (phosphorylated) forms of the cAMP receptor respectively. The behavior of the cAMP receptor in Dictyostelium would be quite analogous then to that of a number of vertebrate hormone receptors where phosphorylation regulates the interaction of the receptor with its proximal cellular respondent. The data presented in the preceding sections are compatible with a role of phosphorylation in the cellular differentiation of D . discoideum. A function of nuclear phosphoproteins in the regulation of transcription seems likely; an autonomous role of the regulatory subunit of the CAMP-dependent protein, analogous perhaps to that of the CAMP-binding, DNA-binding protein of prokaryotes, is not ruled out.

H. PROTOZOA The occurrence of protein kinases and of the phosphorylation of proteins in several classes of protozoa has been described. Most of these reports deal with

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organisms such as Paramecium and Tetrahymena, favorite subjects of study of the developmental biologist, or with Trypanosoma and Plasmodium, important pathogens where the emphasis is on the quest for key enzymes, sufficiently different from those of the host, to make them potential targets for therapeutic attack. 1. Trypanosoma The trypanosomes represent an even deeper branch of eukaryotic evolution than, for example, D . discoideum (150) and in this context the inability of investigators to find a CAMP-dependent protein kinase in any one of several species of trypanosomes examined may be of interest; admittedly, the possibility that this apparent lack of the occurrence of CAMP-dependent protein kinases may have a trivial basis such as, for example, exceptional sensitivity of the relevant domains of the enzyme to proteolysis, is not ruled out. Three protein kinases from Trypanosoma gambiense, the causative agent of sleeping sickness, were partially purified (151); their respective molecular weights were 37,000, 95,000, and 200,000. One of the enzymes catalyzed the phosphorylation of phosvitin and to a lesser extent of histones; the other two enzymes acted primarily on histones and protamine; CAMP and cGMP had no effect on their activities, nor did two CAMP-binding proteins, isolated from the same organism (152). A similar situation obtains in T . cruzi, causative agent of Chagas disease [i.e., there are three CAMP-independent protein kinases, inhibitible by polyamines (153)and there is, in the case of this species, apparently only one CAMP-binding protein which bears no relation to the regulatory subunits of the CAMP-dependent protein kinase regulatory subunits of other eukaryotes (154, 1-55)]. 2 . Plasmodium The occurrence of protein kinase activity was demonstrated also in Plasmodium, the causative agents of malaria; schizonts contained higher levels of protein kinase activity than immature forms of the parasite; preferred substrates were phosphovitin and casein; spermine and spermidine stimulated protein kinase activity; cyclic nucleotides had no effect and the flavone quercetin inhibited (156). 3 . Paramecium

It appears that both CAMP- and cGMP-dependent protein kinases occur in the ciliate Paramecium tetraurelia and that they form part of the system of amplification by which the calcium-potassium action potential, created in resonse to environmental stimuli, is translated into changes of direction of ciliary movement (157, 158). In experiments designed to identify endogenous substrates of CAMP-and cGMP-dependent protein kinases by in vitro phosphorylation, eight

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ciliary proteins were phosphorylated reproducibly in response to the cyclic nucleotides (159).Cyclic GMP, and cGMP-dependentprotein kinases, calmodulin, and the calmodulin-binding protein calcineurin (a phosphoprotein phosphatase) were localized immunocytochemically;they were associated primarily with the cilia, suggesting their participation in the regulation of ciliary motility (160). There is evidence for a role of protein phosphorylation-dephosphorylation in Ca2 -dependent exocytosis, mediated by the trichocysts, the secretory organelles, of P. tetraurelia. The trichocysts are docked at the surface of the cell at specific sites where exocytosis occurs, Experimentally, upon stimulation with picric acid, the trichocysts and the plasma membrane fuse and the secretory organelle is released by the cell. A correlation between the rapid dephosphorylation of protein of molecular weight 65,000 and secretion, stimulated by picric acid, was observed. When secretion was inhibited by Mg2+ or when a temperature-sensitive, secretion-deficient mutant was grown at the nonpermissive temperature, dephosphorylation of the 65,000-dalton protein did not occur (161). +

4. Tetrahymena Several protein kinases were detected in the cilia of Tetrahymena pyriformis. One of these enzymes was apparently cyclic nucleotide-independent and catalyzed the phosphorylation of ciliary tubulin; this protein kinase was associated with the axonemes of the cilia (162).Another four protein kinases were isolated from the soluble fraction of the cilia of T . pyriformis (16.3);two of them were cGMP-dependent, the third required CAMP for activity, and the fourth functioned with either cyclic nucleotide. The two cGMP-dependent kinases catalyzed the phosphorylation of histone and casein whereas the other two kinases acted primarily on histones and protamine. Tubulin was apparently not a substrate. The protein kinases associated with the cilia of Tetrahymena presumably catalyze the phosphorylation of proteins with a role in the control of cellular motility. Other protein kinases found in the cytosol of the organism appear to catalyze the phosphorylation of proteins with (unidentified) metabolic and developmental functions. A CAMP-dependent protein kinase was partially purified from the cytosol of T. pyriformis; the enzyme had relatively high specificity for histones and protamine. The fact that its specific activity varied significantly during growth suggests a regulatory role (164). A T. pyriformis ribosomal protein of molecular weight 38,000 is phosphorylated in response to the starvation of the organism; the protein appears to be equivalent to the S6 protein associated with the small ribosomal subunit of higher organisms. In the case of Tetrahymena it seems to be at the interface between the two subunits, at least when the ribosomes are dissociated by dialysis against a buffer low in magnesium (165). A correlation between the phosphorylation of the S6-like ribosomal protein of T. thermophila and the resistance of the organism to several inhibitors of protein synthesis has been demonstrated (166).

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IV. General Comments What conclusions may be drawn from this survey of the occurrence and functions of protein phosphorylation in single-celled organisms? Phosphoproteins have been found in all organisms and phosphorylation may be assumed to be a ubiquitous mode of the posttranslational modification of proteins and of their functions. To what extent the full spectrum of protein kinases found in multicellular organisms exists also in single-celled organisms is not known. There is no firm evidence for the occurrence of cyclic nucleotide-dependent protein kinases in prokaryotes and in trypanosomes; as pointed out earlier, a limited number of species have been examined and it would be premature to draw conclusions from the negative findings. The CAMP-dependent protein kinases of the single-celled eukaryotes are quite similar to those of the vertebrates and the domains required for the interaction of catalytic and regulatory subunits have been conserved. Tryosine kinases occur in E . coli and yeast and may be assumed to be widespread, if not ubiquitous, in their distribution. It will be of interest to see to what extent Ca2 -regulated kinases occur in single-celled organisms. Not only does the phosphorylation of proteins occur in both prokaryotes and single-celled eukaryotes, but its role is the same as that in the multicellular organism. A number of cases are described in the preceding sections where the phosphorylation of key enzymes permits the tuning of metabolism to the requirements of the individual cell; unlike in multicellular organisms, the regulation of kinase activity in single-celled organisms is controlled by cellular metabolism rather than by hormones. That the role of protein phosphorylation in singlecelled organisms is not confined, however, to the regulation of cellular metabolism is demonstrated by the case of Dictyosrelium where the evolutionary potential for a role of protein phosphorylation in intercellular, integrative signal transmission is realized. Single-celled eukaryotes may serve then as valid models for the study of the effects of protein phosphorylation. These effects may be direct or indirect; the reversible phosphorylation of certain enzymes and structural proteins, leading to changes in their activities, is an example of the direct effect, whereas effects on the synthesis of proteins, on mitosis and meiosis, on cellular differentiation and development, etc., are indirect in the sense that they involve a number of discrete steps of which the phosphorylation of the substrate of the kinase is the first. The effects of phosphorylation on the activities of enzymes and structural proteins and the metabolic consequences of the changes in their activities are generally understood. By contrast, little is known about the mechanisms by which the phosphorylation of proteins exerts the indirect “biological effects. The role of protein phosphorylation in the synthesis of proteins may constitute an intermediate situation; it seems likely that the phosphorylation of the RNA poly+

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merase affects synthesis at the level of transcription and the phosphorylation of the ribosomal protein S6 at the level of translation. The differential effect of protein phosphorylation on the synthesis of inducible proteins remains to be elucidated. It may be anticipated that the study of protein phosphorylation in single-celled organisms will contribute to an understanding of the long-range effects of the phosphorylation of proteins. Unlike cultured vertebrate cells, single-celled organisms can be grown and manipulated in a completely natural environment; since many single-celled eukaryotes are haploid, mutants, defective in either the relevant protein kinase or its substrate or abnormal with respect to the biological end effect of protein phosphorylation, can be selected with relative ease and submitted to analysis under conditions where survival of the organism does not depend on the functioning of the system. It may not be too sanguine then to predict that the study of the phosphorylation of proteins in Saccharomyces, Mucor, Blastocladiella, Dictyostelium, etc., particularly in relation to developmental transitions, will lead to an understanding of the roles of protein phosphorylation in the biology of the organism.

ACKNOWLEDGMENTS The authors’ research is supported by the National Institutes of Health and the American Cancer Society. H.V.R. holds an Ida and Cecil Green investigatorship in Developmental Biochemistry. We are most grateful to the many investigators who furnished us with the unpublished as well as published findings of their research; we thank Carol Breibart for the preparation of the manuscript.

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Bradbury, E. M., Inglis, R. J., Matthews, H. R., and Sarner, N. (1973). EJB 33, 131. Bradbury, E. M., Inglis, R. J., and Mathews, H. R. (1974). Nature (London) 247, 257. Fischer, S. G., and Laemmli, U. K. (1980). Biochemistry 19, 2240. Mueller, R. D., Yasuda, H., and Bradbury, E. M. (1985). JBC 260, 5081. Jeamanowski, A., and Maleszewski, M. (1985). Biochemistry 24, 2360. Chambers, T. C., Langan, T. A., Matthews, H. R., and Bradbury, E. M. (1983). Biochemistry 22, 30. 107. Keuttel, M. R., Schwoch, G., and Jungmann, R. A. (1984). Cell Biol. Int. Rep. 8, 949. 108. Langan, T. A. (1978). Methods Cell Biol. 19, 143. 109. Huebner, V. D., and Matthews, H. R. (1986). JBC 260, 16106. 110. Atmar, V. J., Daniels, G. R., and Kuehn, G. D. (1978). EJB 90, 29. 111. Kuehn, G. D., Affolter, H.-U., Atmar, V. J., Seebeck, T., Gubler, U., and Braun, R. (1979). PNAS 76, 2541. 112. Atmar, V. J., and Kuehn, G. D. (1981). PNAS 78, 5518. 113. Mitchell, J. L. A,, and Wilson, J. M. (1983). BJ 214, 345. 114. Konijn, T. M., van de Meene, J. G. C., Bonner, J. T., and Barkley, D. S . (1976). PNAS 58, 1152. 115. Malchow, D., Bohme, R., and Rahmsdorf, H. J. (1981). EJB 117, 213. 116. Rahmsdorf, H. J., Malchow, D., and Gerisch, G. (1978). FEES Lett. 88, 322. 117. Kuczmarski, E. R., and Spudich, J. A. (1980). PNAS 77, 7292. 118. Maruta, H., Bakes, W., Dieter, P., Marme, D., and Gerisch, G. (1983). EMBO J . 2, 535. 119. Pagh, K., Maruta, H., Claviez, M., and Gerisch, G. (1984). EMBO J . 3, 3171. 120. Claviez, M., Pagh, K., Maruta, H., Bakes, W., Fisher, P., and Gerisch, G. (1982). EMBO J . 1, 1017. 121. Maruta, H., and Korn, E. D, (1977). JBC 252, 6501. 122. Collins, J. H., and Korn, E. D. (1980). JBC 255, 801I . 123. Scholey, J. M., Taylor, K. A., and Kendrick-Jones, J. (1980). Nature (London) 287, 223. 124. Gerisch, G., Fromm, H., Huesgen, A., and Wick, U. (1975). Nature (London) 255, 547. 125. Yoshida, M., Stadler, J., Bertholdt, G., and Gerisch, G. (1984). EMBO J . 3, 2663. 126. Darmon, M., Brachet, P., and Pereira da Silva, L. H. (1975). PNAS 72, 3136. 127. Juliani, M. H., and Klein, C. (1978). Dev.Biol. 62, 162. 128. Majerfeld, I. H., Leichtling, B. H., Meligeni, J. A., Spitz, E., and Rickenberg, H. V. (1984). JBC 259, 654. 129. de Gunzburg, J., Part, D., Guiso, N., and Veron, M. (1984). Biochemistry 23, 3805. 130. Rutherford, C. L., Vaughan, R. L., Cloutier, M. J., Ferris, D. K., and Brickey, D. A. (1984). Biochemistry 23, 461 1. 131. Schoen, C., Arents, J. C., and van Driel, R. (1984). BBA 784, 1. 132. Rahmsdorf, H. J., and Pai, S.-H. (1979). BBA 567, 339. 133. Renart, M. F., Sastre, L., and Sebastian, J. (1984). EJB 140, 47. 134. Leichtling, B. H., Spitz, E., and Rickenberg, H. V. (1981). BBRC 100, 515. 135. de Gunzburg, J., and Veron, M. (1982). EMBO J . 1, 1063. 136. de Witt, R. J. W., Arents, J. C., and van Driel, R. (1982). FEES Lett. 145, 150. 137. Schaller, K. L., Leichtling, B. H.,Rickenberg, H. V., de Gunzburg, J., and Veron, M. unpublished observations. 138. McCarroll, R., Olsen, G. J., Stahl, Y. D., Woese, C. R., and Sogin, M. L. (1983). Biochemistry 22, 5858. 139. Leichtling, B. H., Majerfeld, I. H., Spitz, E., Schaller, K. L., Woffendin, C., Kakinuma, S., and Rickenberg, H. V. (1984). JBC 259, 662. 140. Schaller, K. L., Leichtling, B. H., Majerfeld, I. H., Woffendin, C., Spitz, E., Kakinuma, S., and Rickenberg, H. V. (1984). PNAS 81, 2127. 101. 102. 103. 104. 105. 106.

14. PROKARYOTES AND SINGLE-CELLED EUKARYOTES

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141. Woffendin, C., Chambers, T.C., Schaller, K. L., Leichtling, B. H., and Rickenberg, H. V. (1986). Dev. Biol. 115, 1. 142. Juliani, M. H., Maia, J. C. C., and Bonato, M. C. M. (1983). FEBS Lett. 154, 400. 143. Parish, R. W., Muller, U., and Schmidlin, S. (1977). FEBS Lett. 79, 393. 144. Springer, W. R., Cooper, D. N. W., and Barondes, S. H. (1984). Cell (Cambridge, Muss.)39, 557. 145. Juliani, M. H., and Klein, C. (1981). BBA 662, 256. 146. Lubs-Haukeness, J., and Klein, C. (1982). JBC 257, 12204. 147. Coffman, D. S., Leichtling, B. H., and Rickenberg, H. V. (1981). J. Suprumol. Struct. Cell. Biochem. 15, 369. 148. Coffman, D. S., Leichtling, B. H., and Rickenberg, H. V. (1982). Dev. Biol. 93, 422. 149. Frame, L. T., and Rutherford, C. L. (1984). ABB 232, 47. 149a. Klein, P., Theibert, A , , Fontana, D., and Devreotes, P. N. (1985). JBC 260, 1757. 149b. Klein, C., Lubs-Haukeness, J., and Simons, S. (1985). J . Cell Biol. 100, 715. 150. Sogin, M. L., Elwood, H. J., and Gunderson, J. H. (1986). PNAS 83, 1383. 151. Walter, R. D. (1978). Hoppe-Seyler’s Z. Physiol. Chem. 359, 601. 152. Walter, R. D. (1978). Hoppe-Seyler’s Z. Phsyiol. Chem. 359, 607. 153. Walter, R. D., and Ebert, F. (1979). Tropenmed. Pnrusitol. 30, 9. 154. Rangel-Aldao, R., Tovar, G., and de Ruiz, M. L. (1983). JBC 258, 6979. 155. Rangel-Aldao, R., Allende, O., and Cayama, E. (1985). Mol. Biochem. Parusitol. 14, 75. 156. Wiser, M. F., Eaton, J . W., and Sheppard, J. R. (1983). J. Cell. Biochem. 21, 305. 157. Lewis, R. M., and Nelson, D. L. (1980). BBA 615, 341. 158. Schultz, J. E., and Jantzen, H. M. (1980). FEES Lett. 116, 75. 159. Eistetter, H., Seckler, B., Bryniok, D., and Schultz, J . E. (1983). Eur. J . Cell Biol. 31, 220. 160. Klumpp, S., Steiner, A. L., and Schultz, J. E. (1983). Eur. J. Cell Biol. 32, 164. 161. Gilligan, D. M., and Satir, B. H. (1982). JBC 257, 13903. 162. Murofushi, H. (1973). BBA 327, 354. 163. Murofushi, H. (1974). BBA 370, 130. 164. Majumder, G. C., Shrago, E., and Elson, C. E. (1975). BBA 384, 399. 165. Kristiansen, K., Plesner, P., and Kriiger, A. (1978). EJB 83, 395. 166. Hallberg, R. L., Wilson, P. G., and Sutton, C. (1981). Cell (Cambridge, Muss.)26, 47.

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Author Index Numbers in parentheses are reference numbers and indicate that an author’s work is referred to although the name is not cited in the text. Numbers in italics refer to the page numbers on which the complete reference appears.

A Aamodt, E., 292(60), 312 Abbott-Brown, D., 375(114), 380 Abdel-Fattah Mostafa, M., 70(133), 75 Abeles, R. H.,124(14), 143 Abita, J.-P., 222(33), 223(33), 225(33), 227(33), 229(54), 236(33), 238(33), 239(33), 240(54), 241(54), 247(82), 272(33), 278, 279 Abood, M. E., 376(142), 380 Abrams, B., 32(154), 45 Abumrad, N. A,, 116(62), 121 Abumrad, N. N., 116(62), 121 Accorsi, A., 32(161), 45 Achazi, R. K., 406(139), 416 Acosta-Urguidi, J., 287(28), 289(28), 290(28), 303(28), 312, 342(49), 345(69), 346(49), 355, 356 Adachi, K., 433(54), 435(67), 453 Adams, W. B., 287(23), 288(23), 290(23), 303(23),311. 342(47), 345(47, 70), 355, 356 Adelstein, R. S., 291(54), 297(54, 131), 303(131), 312, 314, 382(2), 384(2), 385(2), 386(18, 20, 21, 24, 27), 387(27), 388(24), 389(42), 390(42), 391(56, 60),392(56), 393(56), 396(83), 399(103, 104, 105, 106),

400(27, 105, 106), 401(20, 21, 103, 104, 105, 106, 110). 402(105, 106, 113), 404(56), 407(160, 162). 408(2), 409(113), 410(185, 188), 411(190), 413, 414, 415, 416, 417, 418 Adibi, S., 108(43, 44),120 Affabris, E., 375(131), 380 Affolter, H.-U., 442(111), 454 Aftring, R. P., 109(47), 120 Agabian, N., 422(12), 452 Agius, L., 130(81), 131(81), 145 Agnew, W. S., 351(110, 112), 357 Agrawal, D., 307(222), 316 Agrawall, H. C . , 307(222), 316 Aguilar-Parada, F., 244(77), 279 Ahern, T., 374(105),379 Ahmad, F., 126(29), 127(29), 144 Ahmad, P. M., 126(29), 127(29), 144 Ahmad, Z., 264(152), 281, 296(121), 314 Aiba, T., 386(26), 414 Ailhaud, G., 170(173), 176 Aitken, A., 25(71), 43, 105(34, 3 3 , 108(35), 120, 154(53), 163(104), 174, 175 Aitken, G. A., 303(233), 307(233), 308(233), 309(233), 31 7 kerstrom, B., 150(29), 173

457

AUTHOR INDEX Akhtar, M., 199(137), 213 Akino, M., 219(14), 249(14), 273(14), 278 Aksoy, M. O., 386(25), 388(25), 396(79), 397(79, 90, 91), 409(91), 414, 415 Albanesi, J. P., 406(147, 148, 149, 150), 407(156), 417 Albert, K. A,, 265(155), 281, 287(30), 289(30), 290(30), 298(139), 299(30), 303(30, 198), 304(198), 308(139), 312, 314, 315, 329(48), 332, 341(31), 347(83), 348(83), 350(83), 355, 356 Alberts, A. W., 191(74), 192(93), 212 Alberts, W. A., 124(12), 125(12), 143 Albertsson-Wikland, K., 166(154), 176 Albrecht, P., 185(41), 211 Albuquerque, E. X.,350(146), 358 Alemany, S . , 62(81), 73 Alexander, M. C., 143(132), 146, 165(146),

Anderson, R. L., 29( 128), 44 Anderson W., Jr., 401(110), 416 Anderson, W. F., 360(4), 377 Andrade, R., 342(143), 346(143), 357 Andrews, D. W., 265(156), 267(156), 281 Anholt, R., 347(81), 349(81), 356 Anthony, D. T., 350(108), 357 Appel, S. H., 309(252), 317, 349(105), 357 Appella, E., 407(154), 410(186), 411(186), 417. 418

Apperson, A., 423(15), 426(15), 452 Appleman, M. M., 165(125), 175 Aquino, A. A,, 150(26), 173 Aracava, Y.,350(146), 358 Aragon, J. J., 29(124), 44 Ardron, D., 29(132), 40(132), 44 Arebalo, R. E., 187(46), 200(149, 150), 201(151), 203(158), 211, 214 176 Arents, J. C., 444(131, 136), 445(131), 454 Ariano, M. A., 293(81), 313 Alger, B. E., 290(31), 299(31), 312, 342(142), 346(142), 357 Armstrong, C. M., 337(132), 357 Alkon, D. L., 287(28), 289(28), 290(28), Armstrong, D. T., 170(171), 176 303(28), 312, 342(49, 51), 345(51, 68, 69), Armstrong, J. C., 273(178), 275(178), 282 346(49), 355,356 Arner, A., 398(99), 415 Alkondon, M., 350(146), 358 Amtzen, C. J., 427(33), 452 Allen, B. L., 31(147), 45, 32(148), 41(148), Arreola, J., 342(141), 357 45 Artaud, F., 276(192), 282 Allen, J. F., 427(33), 452 Ashavaid, T., 341(37), 355 Allende, O., 448(155), 455 Ashby, C. D., 293(64), 312, 436(73), 453 Allfrey, V. G., 25(77), 43 Ashe, J. H.,287(29), 289(29), 290(29), 312 Allhiser, C. L., 255(131), 260(131), 280 Ashour, A.-L. E., I1(37), 42 Allison, W. S., 19(55),42 Assimacopoulos-Jeannet, F. D., 130(89), Allmann, D. W., 195(100), 212 131(89), 132(89), 145, 165(123), 175 Astwood, E. B., 149(12), 152(12), 173 Allred, J. B., 126(31), 132(97, loo), 144, 145 Aswad, D. W., 302(182, 191), 303(191, 227, 228, 233), 307(227, 228, 233), 308(182, Almers, W., 353(126), 357 233), 309(233), 315, 316, 317 Almon, R. R., 349(105), 357 Atmar, V. J., 442(110, 111),442(112),454 Al-Nassar, I., 87(67), 94 Attwood, P. V., 124(13), 143 Alousi, A., 252(110), 280 Au, A., 300(164), 315 Aloyo, V. J., 298(143), 309(143), 314 Avigan, J., 202(205), 215 Amemiya, K.,422(8), 451 Aviv, H., 375(126), 380 Ames, M. M., 258(143), 260(143), 281 Avruch, J., 126(26), 132(26), 134(26), Amesz, H.,363(46), 364(57), 378 135(26), 137(26), 143(132), 144, 146, Anagnoste, B., 253(121), 280 163(106), 165(146), 175. 176 Anderson, J., 50(35, 36), 58(35, 36), 61(35, Awn, E., 170(173), 176 36), 72 Anderson, R. G. G., 257(140), 281, 294(97), Axelrod, D., 349(98), 356 Axelrod, J., 324(13), 331 298(97), 313 Ayad, S. R., 165(134), 175 Anderson, R. G. W., 181(20, 21), 182(20, Ayling, J. E., 219(19), 234(19), 236(19), 278 21), 183(20), 184(20), 190(63), 191(21, Azran, F., 191(73), 212 63), 192(63), 194(63), 207(20), 210, 211

459

AUTHOR INDEX

Baudry, M., 300(165), 315 Bauer, H., 70(136), 71(136), 75 Bacon, G.W., 128(48, 50), 129(68), 131(50), Bean, B. P., 337(131), 340(23), 355, 357 133(50), 137(50), 138(50), 139(50), 140(50, Beaty, N. B., 126(23), 127(23, 42, 43), 121), 141(128), 142(128),144, 145, 146 128(43), 144 Badwey, J. A . , 69(122), 74 Beaudet, A. L., 373(99), 379 Baer, J. E . , 249(88), 279 Beavo, J. A . , 71, 75, 162(102), 163(102), Baglioni, C., 368(70), 373(101), 375(119, 175, 343(57), 356 122, 130), 379, 380 Bechtel, P. J., 162(102), 175 Bair, C . E . , 163(115), 175 Becker-Ursic, D., 431(45), 452 Baker, F. C . , 187(52), 192(52), 202(52), 211 Beebe, S. J., 162(76), 174 Baker, P. F., 308(235), 317 Beg, Z . H., 180(6), 181(6), 195(100), Balasubramaniam, S., 169(162), 176 196(111, 112). 197(114, 117). 198(111, Baldessarini, R. J., 253(122), 254(122), 280 128, 129), 201(6, 153), 202(154, 205), Balkow, C., 169(170), 171(170),176 203(117), 209(129), 210, 212, 213, 214, Balkow, J., 362(12), 371(12), 375(12),377 215 Balkow, K., 365(64), 366(64), 378 Beggs, M., 119(64), 121 Ball, E. G.,130(94), 145 Behman, H. R., 170(171), 176 Ballard, F. J., 208(196), 215 Beins, D. M., 169(162),176 Balogh, A , , 6(18), 18(18),41 Beirne, E., 230(58), 244(58), 279 Bakes, W., 443(118, 120), 454 Belardetti, F., 375(127), 380 Baraban, J. M., 290(31), 299(31), 312, Belfrage, P., 148(1, 2, 4, 5, 9), 149(1, 2, 4, 342(142), 346(142), 357 15, 16, 17, 18, 20, 25), 150 (1, 2, 20, 25, Bzkriiny, K., 396(78), 415 29), 151 (2, 9, 20, 30, 34), 152(16, 38), B6rriiny, M., 396(78), 405(131), 415, 416 153(1, 2, 16, 47), 154(2, 16, 47, 49, 52, 54), 155(52), 156(5, 59), 157(2, 47, 49, 65, Barbaric, S . , 207(185), 214 Barchas, J. D., 250(98), 257(141), 258(98, 66, 69, 70, 71), 158(49, 66, 71, 190), 159(2, 16, 66), 160(2, 5, 49, 66, 70), 142), 260(98), 265(154), 270(163), 161(71), 162(78, IOO), 163(5), 165(132), 271(166), 280, 281 Barchi, R. L., 351(115, 116), 357 166(2, 4, 71), 167(78), 168(4), 169(34), Barden, R. E . , 124(11), 143 172, 173, 174, 175. I77 Barenholz, Y.,194(95), 212 Bell, G. D., 192(88), 212 Barhanin, J., 351(113), 357 Bell, G.I., 432(52), 452 Barkley, D. S . , 442(114), 454 Belman, S., 324(14), 331 Belsham, G.J., 135(107), 139(115), Barnes, E., 266(157), 267(157), 268(157, 140(115), 141(115, 123), 143(123), 146, 160). 269(157), 281 165(145, 149), 176 Barondes, S., 347(72), 356 Belville, J. S., 408(174), 417 Barondes, S. H., 446(144), 455 Barrera, C. R . , 80(15), 93 Bengstsson, G., 150(22), 151(22), 156(22), Barringer, J. A., 233(39), 278 173 Bengur, A. R., 410(186), 411(186), 418 Barmn, J. T., 396(78), 415 Barron, L. L., 116(56), 120 Benjamini, E., 25(67), 43, 61(75), 62(75), 73 Barsotti, R. J., 388(36), 395(72a), 398(36), Benkovic, P. A., 11(33), 34(175), 42, 45 Benkovic, S . J., 5(13), 11(33), 18(13), 404(121), 414, 415, 416 Bartfai, T., 289(33), 290(33), 293(33), 312 32(13), 33(13), 34(175, 176), 36(176), 41, Bartions, V. E. R . , 23(64), 43 42, 45 Bartnicki-Garcia, S., 436(75), 453 Benne, P., 363(46), 378 Bartrons, R . , 6(24), 29(24, 125), 42, 44 Benne, R., 360(5), 362(37), 364(57), 377, Basset, P., 302(187), 303(187), 315 378 Basu, S. K., 182(33), 184(33), 197(33), Bennet, V., 407(159), 417 208(33), 21 I Bennett, J., 427(33), 452

B

460 Bennett, M. K., 296(109), 297(109, 126), 306(109), 314 Bennett, W., 300(163), 315 Bennett, W.F., 255(132), 256(132, 136), 257(136), 264(136), 280, 408(174), 417 Benoit, V., 166(159), 176 Benovic, J. L., 325(18), 332 Bensch, W. R., 182(28), 211 Benson, J. A., 287(24), 288(24), 290(24), 303(24), 311, 342(46), 345(46, 71), 355, 356 Benzie, C. R . , 374(106), 379 Berg, P., 182(33), 184(33), 197(33), 208(33), 211 Berger, J. E., 149(10), 173 Berglund, L., 25(76), 43, 48(12), 50(33), 51(37), 52(43), 53(43), 54(43), 59(69), 60(37, 69), 61(74), 62(74), 68(12), 72, 73, 151(32), 173 Bergstrom, G., 50(33), 55(54, 5 9 , 56(54), 57, 72, 73, 207(176), 214 Berland, M., 90(81), 95 Bernadetto, A , , 375(127), 380 Bemdt, J., 195(102, 103), 204(161), 212, 214 Bernhardt, R . , 82(25), 93, 306(216), 316 Bernier, I., 299(152), 315 Bernier, L., 343(55), 355 Berridge, M. J., 89(70), 94, 287(14), 293(14), 298(14), 321 Bertholdt, G., 444(125), 454 Bertics, P. J., 330(55), 332 Bertrand, O., 56(58), 73 Besharse, J. C . , 100(20), 104(20), 120 Besmond, C., 48(8), 72 Bessman, S., 223(39), 278 Beth, A , , 16(50), 21(50), 42 Beuzard, Y . , 373(94), 379 Beynen, A . C . , 128(47), 130(90, 91), 131(47, 90, 91), 132(47, 90, 91). 144, 145 Bilham, D., 303(233), 307(233), 308(233), 309(233), 317 Bilham, T . , 25(71), 43, 163(104), 175 Birnbaum, R. S . , 161(74), 162(88), 163(88), 166(74), 174 Bimbaumer, L., 325(18), 332 Bjornsson, 0. G . , 200(204), 215 Biserte, G., 236(62), 279 Bjorck, L., 150(29), 173 Bjorgell, P., 157(66, 71), 158(66, 71), 159(66), 160(66), 161(71), 166(71), 174 Bjorntorp, P., 149(13), 173

AUTHOR INDEX Blackmore, P., 37(202), 46 Blackshear, P. J . , 163(106), 165(146), 175, 176 Blair, J., 62(79), 63(79), 73, 141(126), 146 Blair, J. B., 50(34), 53(34, 44),54(34, 44, 51), 57(34), 65(34, 44),66(34, 101). 67(101, 103, 105), 72, 73, 74, 241(68), 279 Blandin-Savoja, F., 247(82), 279 Blaschko, H. J., 248(85), 279 Blaskovics, M. E., 247(81), 279 Blass, J. P., 309(250), 317 Bleile, D. M., 80(8, 11). 84(35), 86(35), 93, 94 Blessing, R. H., 124(16), 144 Blith, D. L., 143(136), 146 Blithe, D. L., 330(54), 332 Blithe, D. R., 330(74), 333 Blobel, G., 182(32), 211 Bloch, K., 124(4), 126(4), 143 Bloch, K. E., 185(42), 211 Block, K. P.,98(1), 99(1), 119 Bloom, F. E., 287(17), 311 Bloxham, D. P., 27(44), 44, 50(22), 56(59), 57(22, 59, 65), 67(108), 72, 73, 74 Bloxham, E. P.,199(137), 213 Blum, W., 422(7), 451 Blumberg, P., 411 (190), 418 Blumenthal, D. K., 405(127, 130), 416 Blytt, H. J., 31(147), 45, 129(71), 145 Boadle-Biber, M., 252(1 IS), 263(1IS), 266(118), 277(201, 202), 280, 282 Boadle-Biber, M. C . , 276(190), 282 Bohme, R . , 443(115), 454 Bollbs, L., 396(87), 398(87), 415 Bollbs, L. L., 406(136), 416 Bonato, M. C . , 438(94), 439(96), 453 Bonato, M. C . M., 446(142), 455 Bond, J. S., 270(164), 281 Bond, M., 395(73), 415 Bonner, I. T., 442(114), 454 Bonner, M. J., 170(181), 177 Bordier, C., 150(24), 173 Borgia, P. T . , 436(76), 453 Borgstrom, B., 156(60), 174 Borin, G., 25(80, 81), 26(80), 43 Borland, M. K., 67(106), 74 Bornstein, P., 309(246), 317, 3 7 3 113, I14), 379, 380 Borthwick, A . C., 127(44a), 133(44a), 138(44a), 141(124), 143(124), 144, 146 Bortz, W. M., 171(188), 177

46 1

AUTHOR INDEX Bosca, L., 29(124), 33(172), 44, 45 Bourgoin, S., 276(192, 194), 277(194), 282 Bourne, H. R . , 376(141, 342), 377(146), 380 Bove, J., 195(103), 196(113), 197(116),212, 213 Bowden, G. T., 330(62), 332 Bowden, J. A , , lOO(8, 9), 119 Bowers, B., 407(153, 155), 417 Bownds, D., 328(25), 332 Bownds, M. D., 328(31), 332 Boyd, G. S . , 151(36), 168(36), 169(36, 161, 164, 167), 173, 176 Boyd, R. W., 107(38), 110(38), 120 Boyer, J., 166(159), 176 Brachet, P., 444(126), 454 Brackett, N., 294(97), 298(97), 313 Brackett, N. L., 257(140), 281 Bradbury, E. M., 60(73), 73, 440( 101), 441(101, 104, 106), 454 Bradbury, J. M., 291(50), 312 Bradford, A. P., 105(35), 108(35), I20 Brand, I. A , , 27(90), 28(118), 31(145), 43,

Bmcks, D. G., 129(64), 145 Bronaugh, R. L., 260(146), 268(146), 281 Brooker, G., 91(85), 95 Broschet, K. 0.. 407(166), 417 Brostrom, C. 0..292(62), 312 Brown, B. M . , 300(162), 303(162), 315 Brown, D. A,, 182(31), 211 Brown, G. G . , 421(3), 451 Brown, J. R . , 81(18), 93 Brown, K., 192(85),212 Brown, K. D., 330(61), 333 Brown, M. S . , 180(2), 181(18, 19, 20, 21, 22), 182(18, 19, 20, 21, 27, 30, 33), 183(20, 22, 36), 184(20, 22, 33, 36), 185(36), 186(45), 187(45, 53), 188(45, 53), 190(18, 22, 45, 63, 64,69, 201, 202), 191(21, 30, 45, 63, 81), 192(19, 63, 92), 194(63), 195(104), 197(33), 198(104), 204(159), 207(20, 22, 36), 208(33, 36), 210, 211, 212, 214, 215 Brown, R. E., 372(92), 375(92, 120, 129), 379, 380 44, 45 Brown, W. E., 197(120), 199(120), 213 Brand, K . , 115(53), 120 Browning, M. D., 300(163, 165), 303(211, Brand, L., 227(51), 279 212), 305(210, 211, 212). 306(212), 315, Brandt, B. L., 353(127), 357 316 Brandt, D. R., 84(36), 94 Brownsey, R. W., 89(71), 94, 125(21), Brandt, H., 63, 74, 195(109),213 126(21, 32, 37), 128(51, 59), 129(59), Brandt, K. G . , 198(125, 126), 213 130(37, 51, 59, 74, 77), 131(37, 51, 59, Braun, R . , 442(111), 454 74, 77). 132(51, 74, 77), 133(37), 134(37, Brautigan, D. L., 309(246), 317 51, 77, 104), 135(77, 104, 107, 108), Breckenridge, B. M . , 292(62), 312 136(37), 137(37, 77, 108, 112), 138(37, 108, 112), 139(77, 112, 115), 140(115), Breeman, W. A. P., 170(178), 176 Brenneman, A . R . , 219(13), 249(13), 278 141(21, 115, 122, 123, 124), 142(51, 77), Brewer, H. B., 180(6), 181(6), 196(112), 143(21, 123, 124, 135), 144, 145, 146, 197(114, 117), 198(129), 201(6), 203(117), 165(145, 148, 149), 176 209(129), 210, 213 Bruckwick, E. A , , 254(125), 257(141), 265(154), 280, 281 Brewer, H. B. Jr., 202(154), 213, 214 Brewer, H. G . , 196(110), 198(111), 213 Brugge, J. S., 300(158), 315 Bricker, L. A., 199(138),213 Brum, G . , 338(15, 16), 339(16), 341(15), 354, 355 Brickey, D. A., 444(130), 454 Bridenbaugh, R., 396(89), 398(89), 415 Brunati, A. M . , 25(80), 43 Bridges, B., 128(59), 129(59), 130(59), Brunelli, M., 343(54), 355 131(59), 145 Brunn, H., 70(132), 75 Bridges, B. J . , 78(5), 82(24), 93 Bruns, G. P., 360(1), 377 Bridges, B. R., 89(72), 94 Brunschede, G. Y . , 195(104), 198(104),212 Bridges, W. F., 219(11), 278 Bryniok, D., 449(159), 455 Brochetto, M. R . , 438(92), 453 Buc, H., 48(4), 49(4), 68(4), 69(4), 72 Brochetto-Braga, M. R., 438(91), 453 Buchanan, B., 6(18), 18(18),41 Brock, D. J. H . , 27(93), 43 Buchanan, B. B., 6(19), 41 Bmkes. .I. P.., 349(107). 357 ,~ Budde, T. H., 170(177), 176

.

AUTHOR INDEX Buechler, K. F., 128(47), 129(69), 131(47), 132(47), 144, 145 Buhrle, C., 302(181), 315 Bullar, B., 406(137), 416 Bullard, W. P., 276(191), 282 Bulter, R. G . , 408(178), 417 Burgess, B., 5(4), 15(4), 24(4), 28(4), 37(4), 41. 12(41), 37(41), 42 Burgess, D. R., 407(166), 417 Burgess, W. H . , 401(110a), 416 Burgett, M. W., 78(4), 83(4), 85(4), 93 Burgoyne, R. D., 302(192), 303(192), 306(213), 316 Burke, B. E., 301(173), 304(214, 215), 306(173, 214, 215), 315, 316 Burlini, N., 434(61), 453 Buse, M. G . , 109(47), 120 Buss, J. E., 405(132), 416 Bustamante, P., 428(36), 452 Bustos, G., 269(161), 281 Butcher, R. W., 152(37), 162(37, 79), 163(37, 113, 115), 173, 174, 175 Butler, T. M., 395(73), 396(80), 397(80, 94), 398(94), 404(121), 415, 416 Butley, M. S . , 291(46), 312 Butterworth, K. R.,252(107), 280 Buxton, D. B., 116(56), 120 Byerly, L., 340(28), 355 Byford, M. F., 56(59), 57(58), 73 Bygdeman, S., 252(108), 280 Bygrave, F. L., 88(51), 94 Bylund, D. B., 403(114), 416 C

Cabrer, B., 372(91), 375(91), 379 Cachelin, A. B., 287(25), 288(25), 289(25), 290(25), 303(25), 311, 340(21, 22). 342(48), 346(48), 355 Calkins, D., 293(64), 312, 436(73), 453 Calvo, J., 342(141), 357 Camardo, J., 344(60), 356 Camardo, J. S . , 287(20), 288(20), 290(20), 303(20), 311, 342(44), 343(59), 344(59, 61), 347(61), 355, 356 Cameron, R.,305(201), 316 Campanile, C. P., 241(67), 242(67), 279 Campbell, E. A . , 371(83, 84), 379 Cande, W. Z . , 395(70, 74), 415 Caplow, M., 292(57, 58). 312

Capony, J.-P., 341(33), 355 Capulong, Z . L., 63(88), 74, 195(109), 213 Caraboeuf, E., 337(10), 354 Caracosa, J. M., 143(138), 146 Carbone, E., 337(129), 357 Cardenas, J. M., 70(130), 75 Cardo, M.-T., 195(103),212 Carlin, R. K., 301(174), 315 Carlos, A . , 391(54), 414 Carlson, C. A., 134(102, 103), 145 Carlson, F. A , , 143(136), 146 Carminatti, H.,50(23), 72 Carnegie, R. R . , 25(75), 43 Caron, M. G . , 303(199), 304(199), 316, 320(1, 2), 321(5, 8), 323(9, lo), 324(10), 325(18), 327(21), 331, 332 Carpenter, G., 287(16), 311, 329(50), 330(51), 332 Carr, F. P. A , , 229(55), 230(55), 234(55), 243(55), 279 Carroll, D., 69(120), 70(120), 74 Carroll, R. C., 408(178), 417 Casadei, J. M., 351(116), 357 Casazza, J. P., 18(52), 42 Casnellie, J. E., 302(190), 303(190), 315, 351(120), 357. 409(184), 417 Caspani, G., 434(61), 453 Cassel, D., 376(139), 380 Cassidy, P. S . , 396(85, 87, 88), 398(85, 87), 415 Castagna, M., 242(71, 72), 279, 298(146), 314 Castaneda, E., 374(108), 379 Castano, J. G . , 27(111, 114), 28( 11 I , I14), 44 Castellanos, R. M. P., 431(47), 452 Castellucci, V., 287(18, 19), 288(18, 19), 290(18, 19), 303(18, 19), 311 Castellucci, V. F., 343(54, 55, 56, 58), 347(58, 73), 355. 356 Cate, R. L., 84(33), 94 Caterson, I. D., 88(55), 94 Catt, K. J., 162(89, 90, 91). 169(89, 90, 91, 165), 170(89, 90, 91), 174, 176 Catterall, W. A., 302(195), 303(195), 316. 342(39, 40, 138), 351(114, 118, 119, 120, 121), 352(121, 122), 353(121, 128), 355, 357 Cavalie, A., 340(29), 342(139), 355, 357 Cavanagh, H. D., 328(26), 332 Cavanee, W. K., 190(70), 212

AUTHOR INDEX Cayama, E., 448(155), 455 Cebra, J. J., 407(164), 408(164), 417 Cerione, R. A., 323(10), 324(10), 325(18), 331, 332 Cermolacce, C., 170(173), 176 Cem, C., 69(125), 74 Chacko, S . , 386(24), 388(24, 33, 35, 37), 390(33, 46), 414 Chad, J. E., 287(27), 289(27), 290(27), 303(27), 312 Chambers, T. C., 441(106), 446(141), 454, 455

C h a m s , H., 229(54), 240(54), 241(54), 279 Chan, K.-F.J., 310(256), 317 Chan, T. M., 66(100), 67(100, 104), 74 Chandra, T. S., 391(54), 414 Chang, C. C. Y., 191(72),212 Chmg, F.-H., 377(146), 380 Chang, N., 222(33), 223(33), 225(33), 227(33), 236(33), 238(33), 239(33), 272(33), 278 Chang, T. Y., 191(72), 212 Changeux, J.-P., 302(194), 303(194), 310(194), 316, 329(39, 43, 44),332, 347(77, 79). 348(86, 87, 90,91), 349(100), 350(90), 356, 357 Chantler, P. D., 384(7, 9). 388(7, 9, 31), 389(39), 413, 414 Chappe, B., 185(41), 211 Chasan, R., 399(101), 407(165),416, 417 Chatterjee, M., 396(84), 398(84), 415 Chatterjee, T., 33(167), 34(180), 35(188), 45, 46

Chaumet-Riffaud, P., 48(8), 72 Cheland, W. W . , 12(38),42 Chemouilli, P.,347(77), 356 Chen, H. W., 188(54), 189(60), 190(60, 70), 191(71), 192(54), 193(54), 205(54), 211, 212 Chen, J., 191(74), 212 Chen, J.-J., 376(132), 380 Chen, J. S., 192(93), 212 Chen, L., 170(184), 177 Cheng, K.,91(86, 92), 95 Cherbas, L., 362(31), 378 Cherhazi, B., 301(175), 305(175),315 Cherrington, A. D.,40(205), 46 Chessa, G., 25(80), 26(80), 43 Cheung, W. Y . , 295(107), 309(249), 313, 317 Chin, D. J . , 181(18, 19, 21, 22), 182(18, 19, 21, 30, 33), 183(22), 184(22, 33), 190(18,

463 22, 201, 202), 191(30), 192(19, 21), 197(33), 207(22), 208(33), 210, 211 Chinkers, M., 330(53), 332 Chiu, A. Y . , 344(62), 356 Choate, G., 31(144), 35(144), 44 Chock, P. B., 87(47), 94, 162(93, 96, 97), 163(107), 175, 310(256), 317, 389(42), 390(42), 392(61), 414 Chou, C.-H. J., 298(142), 307(142, 224), 314, 316 Chow, J. C., 195(105),212 Chrisman, T., 5(4, 5 ) , 12(41), 13(5), 15(4, 5), 17(5), 18(5), 22(5), 24(4, 51, 28(4, 5). 29(5), 37(4, 5 , 41), 41, 42, 200(141), 201(141), 213 Chrisman, T. D., 7(29), 22(29), 37(29), 42, 242(70), 279 Christiansen, R. Z . , 130(90), 131(90), 132(90), 145 Chuang, D. T., 99(5), 100(5), 119 Chung, T., 425(26), 452 Ciechanover, A., 207(184), 214 Cimadevilla, 362(25), 378 Cimbala, M. A., 50(34), 53(34, 44), 54(34, 44),57(34), 65(34, 44),66(34), 72, 73 Cioe, L., 375(127),380 Cladaras, C., 48(18), 50(21), 72 Clark, C. T., 249(88), 279 Clark, M. G . , 31(146), 44, 45, 67(102), 74 Clarke, C. F., 182(25), 192(25, 91). 210, 212 Clarke, S. D.,128(62), 129(62), 132(62), 145 Claus, T., 7(29), 12(41), 22(29), 37(29, 41), 42 Claus, T. H., 5(1, 4, 5, 6, 14), 6(14, 22, 23), 7(14), 9(32), 12(22, 23), 12(22), 13(5, 6), 14(4, 5, 6, 14, 22, 23), 15(5, 6, 14), 17(5, 6), 18(5), 20(32), 22(5, 6, 14, 60), 24(4, 5, 6), 27(91, 104, 110, 113), 28(1, 4, 5, 6, 23, 91, 110, 113, 115). 29(5, 6, 91), 30(6, 113), 32(150, 157, 160), 33(160), 34(157, 182), 35(157, 182, 186), 37(1, 4, 5, 6, 14, 22, 200), 38(200), 40(6, 205), 41, 42, 43, 44, 45, 46, 48(3), 49(3), 53(45, 48), 54(45, 50), 55(53), 59(48, 71), 60(48), 61(48), 62(50, 71), 65(45, 95). 66(48, 95, 100). 67(3, loo), 71(3), 72, 73, 74, 200(141), 201(141), 213 Claviez, M., 443(119, 120), 454 Clegg, R. A., 130(85), 131(85), 133(101), 137(101), 138(101), 139(101), 140(101), 142(129),145, 146

464 Clegg, R. J., 192(88),212 Cleland, W. W., 274(184), 282 Clemens, M. J., 362(38), 363(38, 50). 364(38), 372(92), 373(103), 374(103, 104), 375(92, 121), 378, 379, 380 Clifton, D., 6(20), 42 Cloutier, G., 252( 117), 280 Cloutier, M. J., 444(130), 454 Cobb, C., 7(27), 8(27), 42 Cobb, M. H., 287(15), 300(15), 311 Cochet, C., 330(66), 333 Codina, J., 325(18), 332 Coffman, D. S . , 447(147, 148), 455 Cognard, C., 337(133), 357 Cohen, D. C., 190(65), 192(65), 193(65),212 Cohen, P., 23(65), 25(70, 71, 73), 26(84), 42, 43, 51(38), 60(38), 62(77, 78, 80, 81, 82, 83), 63(77, 80, 82, 83), 64(83), 65(77), 71, 72, 73, 75, 126(27), 127(27), 137(27, ill), 138(1I I), 139(111), 140(11l), 141(125), 142(131),144, 146, 154(50, 51, 53), 156(64), 163(104, 105), 165(150), 174, 175, 176, 198(132), 200(140), 201(140), 208(190), 213, 215, 225(46, 47, 48), 243(46), 264(150, 153), 266(153), 278, 281, 296(123, 124), 297(124), 303(232, 233, 234), 307(232, 233), 308(233, 234), 309(232, 233, 243, 244, 245, 248), 310(255),314, 317, 336(4), 354 Cohen, S., 287(16), 311, 329(50), 330(51, 53, 69), 332, 333 Colbran, J. R., 149(19), 150(19), 151(19, 3 3 , 170(35), 173 Cole, H. A , , 386(16), 389(44), 405(129), 413, 414, 416 Coll, K. E., 88(50), 94 Collins, C. A., 208(198), 215 Collins, J. H.,406(146), 407(151, 152, 153), 417, 443(122), 454 Colosia, A , , 5(4), 7(29), 12(41), 15(4), 22(29), 24(4), 28(4), 37(4, 29, 41), 41, 42 Colvin, R. A , , 341(37), 355 Comerci, C., 329(47), 332 Comita, P . B., 185(40),211 Condit, J. R., Jr., 396(83), 398(83), 415 Conelman, K. A., 407(165), 417 Connelly, J. L., 100(8, 9, 10). 119 Connolly, T. M., 409(181), 417 Connors, J. M., 330(64), 333 Conrado, R., 342(42), 345(42), 355

AUTHOR INDEX

Conti, M. A . , 386(24), 388(24), 399(103), 401(103), 407(160), 414, 416, 417 Conti-Devirgilis, L., 205(168), 214 Cook, G. A., 109(50), 113(50), 115(50), 120, 125(20), 126(20), 129(20), 130(20), I44 Cook, J. H . , 328(32), 332 Cook, K. G . , 92(97), 95, 104(29, 32, 33), 105(32, 33, 34, 353, 107(38), 108(35), 109(29), 110(38), 120, 151(34, 35), 169(34), 170(35), 173 Cooke, R . , 404(126), 416 Cooley, L. B., 406(138), 416 Coon, M. J., 19(54), 42, 100(7), 119 Cooper, A. D., 200(148), 205(169), 214 Cooper, D. N. W., 446(144),455 Cooper, J. A., 330(52, 66),332, 333 Cooper, J. R . , 249(88), 271(170), 279, 281 Cooper, M. E., 165(143),176 Cooper, R. H., 78(5), 82(24), 83(29), 84(34), 93 Coppersmith, J. C., 351(118), 357 Corbin, J. D., 32(157), 34(157), 35(157), 45, 152(40), 153(40), 162(76, 84), 173, 174, 242(70), 279, 291(40), 312 Cordle, S. R . , 149(19), 150(19), 151(19), 173 Corkey, B. E., 88(50), 94 Cornell, N. W., 199(135),213 Cornish-Bowden, A . , 236(64), 279 Corredia, C., 33, 45 Correira, J. J., 7(27), 8(27), 42 Corstorphine, C. G . , 130(79), 131(79), 132(79), 139(79), 145 Costa, M. R. C., 302(195), 303(195),316, 351(120, 121). 352(121, 122), 353(121), 357 Costello, C. E., 185(40), 211 Cote, G. P., 406(147), 407(151, 154, 155, 156). 417 Cotman, C. W., 297(128), 306(128), 314 Cottam, G. L., 48(18, 19, 20, 21), 57(62, 64),65(93, 96), 66(96, 99), 67(99), 72. 73, 74, 207(179), 214 Cotton, P. C., 300(158), 315 Cotton, R . G . , 247(83), 279 Cottreau, D., 48(8), 72 Coughlin, B. A . , 310(254), 317 Cox, R. P., 99(5), 100(5), 119 Cozzone, A . J., 423(18), 425(18), 452 Craig, R., 393(67), 415 Craviso, G. L., 251(103), 280

465

AUTHOR INDEX Cree, T. C., I16(59, 60). 121 Crettaz, M., 130(74), 131(74), 132(74), 145 Creveling, C. R., 253(120), 280 Crisp, D. M., 48(9), 72 Croall, D. E.,207(174), 214 Crompton, M., 89(67), 94 Cross, D. G., 34(179), 45 Cross, R. A., 396(75a), 415 Crouch, D., 363(53), 378 Crow, M. T., 404(122), 416 Cseke, C., 6(18, 19), 18(18), 41 Cuatrecasas, P., 91(89, 93), 95, 165(136), 175, 330(63, 81), 331(85), 333 Cumming, D. A., 6(23), 12(23), 14(23), 28(23), 42 Cummings, R. D., 181(20), 182(20), 183(20), 184(20), 207(20), 210 Cunningham, L. W., 19(56), 42 Cume, S., 191(74), 212 Curtis, B. M., 342(39, 138), 355, 357 Curtis, R. M., 342(40), 355 Cushman, S . W., 143(133), 146 Czech, M. P., 91(91), 95, 330(65, 72). 333 D Dabney, B. J . , 373(99), 379 Dabrowska, R., 297(129), 303(129), 314, 386(25), 388(25), 413 Daggy, P., 91(86), 95 Dahlqvist, U., 50(36), 51(42), 53(42), 54(42), 55(42, 54), 56(54), 58(36), 61(36), 72, 73 Dahlqvist-Edberg, U., 34, 35(187), 45, 46, 57(67), 69(118), 73, 74, 200(142), 201(142), 207(178), 213, 214 Daile, P., 25(75), 43 Dalldorf, F., 292(58), 312 Daly, J. W., 253(120), 280, 350(146), 358 Damaille, J . G . , 341(33), 355 Damuni, Z . , 85(43), 86(46), 91(43), 92(46), 93(46), 94, 104(30), 107(30), 108(30), 109(30), 119(65, 66), 120, 121 Dancis, J., 99(6), 100(6), 119 D’Angelo, G . L., 250(95), 280 Daniel, J. L., 408(173), 409(176), 417 Daniels, G. R., 442(110), 454 Danner, D. J., lOO(11, 18, 20). 101(11), 102(18), 104(20), 109(11), 120 Danner, M. J., loo@), 119 Darmon, M., 444(126), 454

Dambrough, C., 362(14), 377 Das, A., 362(16, 21, 23), 363(21, 48, 49). 377, 378 Das, H. K., 362(21, 23), 363(21, 49), 377, 3 78 Dasgupta, A., 362(16), 363(48), 376(137), 377, 378, 380 Dasgupta, J. D., 349(96), 356 DaSilva, A, M., 439(96), 453 Dastidar, P. G . , 330(71), 333 Datta, A., 362(17), 364(55), 377, 378 Datta, A. G . , 32(154), 34(180), 45 Dautrevaux, M., 236(62), 279 Davidson, H., 192(87), 212 Davies, D. R., 15(49), 15(49), 22(49, 62), 23(49), 24(49), 37(49), 42, 43, 132(99), 14s Davies, J., 431(45), 452 Davies, J. I., 148(7), 172 Davis, C. G . , 302(193), 303(193), 316, 329(42, 4 3 , 332, 348(85, 88, 89, 93), 350(85), 356 Davis, E. R., 300(170), 315 Davis, J. Q . , 407(159), 417 Davis P. F., 85(39), 94 Davis, R. J., 330(65), 333 Davison, A. M., 189(59), 211 Davydova, E. K . , 377(148), 380 Dawes, J., 328(25), 332 Deal, W. C. Jr., 27(86, 87, 101), 43, 44 DeBenedetti, A., 368(70), 379 Debus, G., 277(200), 282 DeBuysere, M., 89(62), 94 DeBuysere, M. S . , 116(55), 120 DeCamilli, P., 291(49, 56), 293(75, 77), 305(201, 202, 203), 306(217), 312, 313, 316 Degerman, E.,165(132), 175 DeGunzburg, J., 444(129, 135, 137), 445(129), 454 DeHaas, C. G. M., 200(146), 201(146), 213 DeHaen, C., 166(155), 176 De Haro, C., 362(17, 18. 20). 363(81, 20), 377 Deibler, G. E., 307(221), 316 DeJonge, H. R., 293(71a), 303(71a), 313 DeLanerolle, P., 396(83), 398(83), 399(105), 400(105), 401(105), 402(105, 113), 409(113), 415, 416 DeLarco, J. E.,330(60), 333

466 DeLaunay, J., 374(111), 379 Delcour, A. H., 350(147), 358 Delidakis, C., 371(83), 379 Delluva, A. M., 249(87), 279 DeLorenzo, R. J., 294(90), 296(113), 297(127), 301(90, 172, 173), 304(214, 215), 306(113, 173, 214, 215), 307(113), 313, 314, 315, 316 DeMaine, M. M., 5(13), 18(13), 32(13), 33(13), 41 DeMartino, G. N., 181(20), 182(20), 183(20), 184(20), 207(20, 174), 210, 214 Dembure, P., 307(224), 316 Dempsey, M. E., 187(47), 211 Denton, R. M., 78(5), 82(24), 83(29), 84(34), 85(41), 88(48), 89(63, 65, 69, 71, 72, 73), 90(73, 74, 75), 91(73), 93, 94. 95, 108(42), 120, 125(17, 18, 21), 126(21, 32, 37), 127(44, 44a), 128(44, 51, 57, 59), 129(44, 57, 59), 130(37, 44,51, 57, 59, 74, 76, 77, 80, 82, 89), 131(37, 44,51, 59, 74, 76, 77, 80, 82, 89). 132(51, 74, 77, 80, 82, 89), 133(37, 44a), 134(37, 51, 77, 104), 135(77, 104, 107), 136(37), 137(37, 77, 113), 138(37, 44a), 139(77, 115). 140(115), 141(21, 115, 122, 123, 124). 142(51, 77), 143(21, 123, 124, 135), 143, 144, 145, 146, 165(145, 148, 149), 176 De-Paoli, A. A., 91(86), 95 DePaoli-Roach, A. A., 264(152), 281, 296(121), 314, 396(86), 398(86), 415 DePeyer, J. E., 287(25), 288(25), 289(25), 290(25), 303(25), 311, 340(21, 22), 342(48), 346(48), 355 DeRiemer, S. A., 287(30), 289(30), 290(30), 299(30), 303(30), 312, 341(31), 355 DeRuiz, M. L., 448(154), 455 DeSalu, 0.. 373(93), 379 Deschpande, S . S . , 350(146), 358 Desnuelle, P., 155(58), 174 DeTitta, G. T., 124(16), 144 Detre, J. A,, 302(182), 308(182), 315 Deutscher, J., 425(28), 426(28, 30). 452 Devi, S. U., 194(97), 212 Devillers-Thiery, A., 347(77), 356 Devreotes, P. N., 447(149a), 455 Dewerchin, M. A., 437(85), 453 DeWitt, R. J. W., 444(136), 454 Deykin, D., 170(183), 177 Dhillon, G. S . , 162(92, 101), 163(101), 164(116), 166(116), 174, 175

AUTHOR INDEX Dhondt, J.-L.,236(62), 279 Diamond, I., 300(164), 302(193), 303(193), 315, 316, 329(42, 45), 332, 348(85, 88, 89, 93), 350(85), 356 DiBartolomeis, M. J., 291(52), 312 Dicker, P., 330(61), 333, 374(106), 379 Dieter, P., 443(118), 454 Dietschy, J. M., 204(159), 214 Dijan, P., 170(173), 176 Diller, E. R., 182(28), 211 Dillon, P. F., 396(79), 397(79), 415 Dills, S. S., 423(15), 426(15), 452 Dils, R., 126(28), 128(49), 144 Dini, L.,205(168), 214 DiSegni, G., 361(6), 377 Dixon, G. H., 51(38), 60(38), 72, 81(18), 93 Dolei, A., 375(127), 380 Dolphin, A. C., 305(205), 316 Donella-Deana, A., 154(53), 174 Donlon, J., 224(36, 40), 225(42), 227(42), 228(42), 230(42, 58), 232(42), 238(42), 240(42), 241(42), 243(42), 244(58), 278 Donner, J., 158(190), 177 Doolittle, R. F., 184(37), 211 Doroshenko, P. A., 287(26), 289(26), 290(26), 303(26), 311, 340(26, 30). 355 Deskeland, A. L., 221(25), 278 Deskeland, A. P., 223(37), 235(37), 236(37), 237(37), 238(37), 242(37, 70), 243(37), 278, 279 D~keland,S . O., 163(103), 175, 223(37), 235(37), 236(37), 237(37), 238(37), 242(37, 70), 243(37), 278, 279 Doucet, J. P., 305(209), 316 Dower, W. J., 375(129), 380 Downes, C. P., 287(13), 292(13), 293(13), 298(13), 309(13), 311 Dratz, E. A., 184(38), 211 Dreyer, E., 266(157), 267(157), 268(157, 160), 269(157), 281 Dreyer, W. J., 328(24, 32). 332, 347(74), 356 Dreyfus, J.-C., 48(8), 69(116), 72, 74 Driska, S . P., 396(79), 397(79), 415 Drummond, A. H., 342(46), 345(46), 355 Drummond, G. I., 292(83), 293(83), 313 Duckworth, H. W., 80(12), 93 Dufau, M. L., 162(89, 90, 91), 169(89, 90, 91, 165), 170(89, 90, 91), 174, I76 Dugan, R. E., 180(4), 188(4), 193(4), 210 Dunaway, G. A. Jr., 27(88, 89), 43 Dunbar, B., 56(59), 57(59), 73

467

AUTHOR INDEX Duncan, R., 375(123), 380 Dunlap, K.,341(135, 136, 137), 357 Dunn, L. A., 294(94), 313 Dunwiddie, T. V., 293(79), 300(163), 313, 315 Durham, L. A., 170(172), I76 Dwyer, J., 348(92). 356 Dyson, R. D., 70(130), 75

E Earp, H. S., 330(70), 333 Easom, R. A,, 204(163), 205(162, 163), 214 Easter, D. J., 128(49), 144 Eaton, C. R., 399(104), 401(104), 416 Eaton, J. W., 448(156), 455 Ebashi, S., 397(95), 415 Ebersohl, R. D., 162(77), 166(77), 167(77), 174 Ebert, F., 448(153), 455 Ebstein, B., 260(146), 268(146), 281 Eckert, R., 287(27), 289(27), 290(27), 303(27), 312 Edelman, A. M., 257(141), 258(142), 265(154), 271(166), 281, 400(108), 409(184),416, 417 Edelstein, I., 33(167, 170), 34(178), 45 Edgell, N. J., 127(44a), 133(44a), 138(44a), 141(122), 144, 146, 165(148), 176 Edlund, B., 50(35, 36, 37), 51(37), 58(35, 36), 60(37), 61(35, 36), 72 Edwards, P. A,, 181(14, 17), 182(25), 187(51), 190(17, 66, 68), 192(25, 89, 90, 91), 202(17), 210, 211, 212 Egan, J. J., 329(47), 332, 409(179), 417 Eggleston, D., 275(187, 188), 282 Eggleston, L. V., 48(14), 72 Ehrenfeld, E., 362(10), 375(10), 377 Ehrlich, Y . H., 302(186), 303(186), 314 Eichner, R., 153(48), 173 Eigenbrodt, E., 70(131, 132, 133, 134, 135, 136), 71(136, 137). 75 Einig, I., 291(56), 312 Eisenach, J. C . , 329(41), 332, 348(94), 356 Eisenberg, E., 382(2), 384(2), 385(2, 13). 391(13, 56, 60),392(13, 56, 61), 393(56), 404(56), 408(2), 413, 414 Eisenberg, F., 219(11), 278 Eisenstein, A. B., 244(80), 279 Eistetter, H.,449(159), 455 Ekdahl, K. N., 34(184), 45

Ekman, P., 34(184), 35(187), 45, 46, 50(36), 51(42), 53(42), 54(42), 55(54, 55, 57), 56(54, 55, 57), 57(55), 58(36, 66), 59, 61(36), 65(94), 66(66), 69(118), 72, 73, 74, 200(142), 201(142), 207(176, 177, 178), 213, 214 Eldor, A., 411(191), 418 El-Doq, H. A., 35, 46 Eley, M. H.,80(15), 93 Elks, M . L., 165(129, 130, 131), 175 Ellingboe, J., 152(39), 173 El-Maghrabi, M. R., 5(1, 4, 5, 6, 7, 14, 15, 17), 6(7, 14, 15, 23, 25), 7(7, 14, 15, 17, 25, 26, 27, 29, 31), 8(15, 27, 31). 9(32), 12(23), 12(15, 25, 41, 42), 13(5, 6, 17, 25, 42, 4 3 , 14(4, 5, 6, 14, 17, 22, 23, 25, 26, 42, 45). 15(5, 6, 14, 17, 25, 26, 31, 50), 17(5, 6, 25), 18(5, 7, 26, 45, 53), 20(32, 53), 21(25, 42, 50, 53), 22(5, 6, 14, 17, 25, 26, 29, 31, 60),23(15), 24(4, 5, 6, 7, 26, 31, 66), 25(31), 26(84), 27(91, 113), 28(1, 4, 5, 6, 23, 91, 113, 115), 29(5, 6, 91), 30(6, 113), 32(160, 164), 33(160, 164, 166). 34(166, 176, 182, 185). 35(182, 186), 36(176), 37(1, 4, 5, 6, 14, 29, 41, 200), 38(200, 203), 40(6, 7).41, 42, 43, 44, 45, 46, 53(48), 54(50), 59(48, 71), 60,61, 62(50, 71, 80), 63(80), 66(48), 73, 200(141), 201(141), 213 El Mestikawy, S . , 269(162), 281 Elsas, L. J., 100(11, 20), 101(11), 104(20), 109(11), 119, 120 Elson, C. E., 449(164), 455 Elson, E. L., 349(98), 356 Elwood, H. J., 448(150), 455 Elzinga, M., 386(17a), 401(110), 413, 416 Enami, M., 424(20), 452 Endo, A., 181(24), 182(24, 26, 27, 29), 183(24), 191(24, 26, 29, 7 9 , 210, 211, 212 Endo, H.,389(38), 414 Engelmann, R., 426(30), 452 England, P. J., 89(64), 94 Engstrom, L., 25(76), 35(187), 43, 46, 50(32, 33, 35, 36), 51(37, 39, 40, 41, 42), 52(43), 53(42, 43, 47), 54(42, 43). 55(42, 54, 55), 56(54, 5 9 , 57(55), 58(35, 36), 59(69), 60(37, 69), 61(35, 36, 74). 62(40, 74, 76), 72, 73, 200(142), 201(142), 207(176), 213, 214, 223(38), 278 Epstein, A., 436(78), 453 Epstein, P., 438(90), 453

468

AUTHOR INDEX

Erickson, S . K., 189(59), 191(80), 200(148), 205(169), 211, 212, 214 Erikson, E., 309(247), 317 Erikson, I., 55(57), 56(57), 57 Erikson, R. L., 309(247), 317 Eriksson, H., 158(190),177 Eriksson, I., 207(177), 214 Erlanson-Albertsson, C., 156(60), 174 Erlichman, J., 291(42, 5 9 , 312 Ernst, V., 362(29, 24, 30), 365(65), 366(65), 367(30), 369(72), 370(30, 79, 80, 81), 371(30, 81, 82), 372(89), 374(111), 378, 3 79 Erondu, N. E.,296(109), 297(109, 126), 306(109), 314 Espinal, J., 102(21), 103(21), 109(49), 110(21), 111(21), 112(21, 49), 113(21, 49), 115(49), 118(49, 63). 119(64), 120, 121 Euler, U. S . , 252(108), 280 Eusebi, F., 351(148), 358 Evans, C., 48(19, 20), 72 Evenson, K. J., 199(139),200(139), 202(156, 157), 203(139, 156), 204(139), 205(139), 213, 214 Everson, W. V., 163(105), 175 Exton, J. H., 37(202), 46, 50(28, 29), 66(98, loo), 67(100, 104). 72, 74, 162(86), 163(86), 165(123), 174, 175 Eyer, P., 11(34), 42

F Fagard, R., 369(77), 370(77), 373(96), 374(112), 375(96), 379 Fain, J. N., 90(80), 95, 148(6, 8), 160(8), 162(83), 163(112), 172, 174, 175 Fairbanks, K. P., 191(78), 192(78), 193(78), 212 Faloona, G. R., 244(77, 78), 245(78), 279 Fambrough, D., 349(97, 102), 356, 357 Fan, C.-C., 162(81), 163(81), 174 Farese, R. V., 90(76), 95 Farrell, P., 262(12), 371(12), 375(12), 377 Farrell, P. J., 362(32), 378 Faniaux, J. P., 236(62), 279 Fatania, H. R., 78(7), 92(96, 98), 93, 95, 99(3), 100(3, 19). 101(19), 102(21), 103(21, 26), 104(3, 19, 28), 105(19), 106(19), 107(19, 26, 40), 108(40), 109(19, 26, 28, 4 3 , llO(21, 4 9 , 111(3, 21, 45)’ 112(21, 4 3 , 113(21),119, 120

Faupel, R. P., 58(68), 73 Faust, J. R., 181(18, 21, 22), 182(18, 21, 27, 30), 183(22), 184(22), 190(18, 22, 201, 202). 191(21, 30, 81), 207(22), 210, 211, 212, 215 Feamley, I. M., 105(35), 108(35), 120 Fecheimer, M., 407(164), 408(164), 417 Federman, P., 372(90), 379 Fedulova, S . A., 340(25), 355 Fehlmann, M., 143(137), 146, 165(144, 147). 176 Feingold, K., 205(169), 214 Feingold, K. R., 205(165), 214 Feinstein, M. B., 329(47), 332, 348(92), 356, 409(179), 417 Feliu, J. E., 27(11l), 28( 11l), 44, 54(49), 59(70), 63(86), 65(49, 70), 66(49), 71(49), 73, 74 Feman, E. R., 373(101), 379 Fenwick, E. M., 340(27), 355 Feramisco, J. R., 25(78), 43 Femstrom, J. D., 273(182), 274(182), 282 Ferrendelli, J. A., 293(82), 313 Ferrer, A., 197(118), 198(118),213 Ferris, D. K., 444(130), 454 Fessa, G., 25(81), 43 Fetzer, V., 249(94), 280 Fewtrell, C., 331(82), 333 Filsell, 0. H., 31(146), 45 Finegold, K. R., 198(133),213 Finer-Moore, J., 183(36), 184(36), 185(36), 207(36), 208(36), 211 Fischbach, G., 341(136, 137), 357 Fischbach, G. D., 349(104), 357 Fischer, E. H., 293(64), 312, 431(46), 436(73), 452, 453 Fischer, S . G . , 441(103), 454 Fish, S . , 124(14), 143 Fishbean, R., 11(42), 42 Fisher, D. B., 219(8, 12), 221(23), 222(12, 23, 28, 29), 223(23), 225(23), 236(29), 249(12), 250(29), 275(12), 278 Fisher, H. F., 34(179), 45 Fisher, J. M., 362(8), 377 Fisher, M. J., 26(84), 43, 62(80), 63(80), 73, 229(57), 241(66), 279 Fisher, P., 443(120), 454 Fishman, P. H., 303(200), 304(200). 316 Fister, P . , 70(135), 75 Flatmark, T., 221(25), 223(37), 235(37),

AUTHOR INDEX

236(37), 237(37), 238(37), 242(37, 70). 243(37), 278, 279 Flatt, J. P., 130(94), 145 Flawiii, M. M., 435(68, 69), 453 Fleischer, N., 291(41, 42), 312 Fletcher, W. H.,287(29), 289(29), 290(29), 312 Flicker, P. F., 390(50), 407(157), 414, 417 Flockeai, V., 293(72), 303(72), 313, 338(16), 339(16), 341(35), 342(139), 355, 357 Flockhart, D. A., 32(157), 34(157), 35(157), 45, 53(48), 59(48), 60(48), 61(48), 66(48), 73 Florio, V. A,, 67(106), 74 Flory, W., 50(24), 54(24), 72 Foe, L. G . , 25(138), 30(138), 40(138), 44 Fogelman, A. M., 181(14, 17), 182(25), 187(51), 190(17, 66, 68), 192(25, 89, 90, 91), 202(17), 210, 211, 212 Font, E., 197(124),213 Fontana, D., 447(149a), 455 Forest, C., 170(173), 176 Fom, J., 294(89, 89a, 98), 301(89, 89a), 305(204), 313, 316 Forray, C . , 294(87), 313 Forte, L. R., 403(114), 416 Foster, D. W., 128(55), 130(55), 144 Foster, J. L., 53(44), 54(44), 65(44), 66(101), 67(101, 105), 73, 74, 241(68), 279 Fothergill, L. A,, 56(59), 57(59), 73 Foukes, J. G . , 62(78), 73. 163(105), 175, 225(47), 278, 309(247), 317, 370(80), 372(89), 379 Fowler, S., 170(182), 177 Fowier, V. M., 407(159), 417 Fox, A., 337(134), 357 Fox, C. F., 330(67, 71), 333 Fox, E., 5(1, 14, 15). 6(14, 15), 7(14, 15, 29), 8(15), 12(15), 15(14), 16(14), 22(14, 29), 23(15), 28(1), 37(1, 14, 29), 41, 42 Fox, J. E. B., 408(177), 417 Fox, R. W., 29(133), 40(133), 44 Foyt, H. L., 401(111), 416 Frackelton, A. R., Jr., 330(64), 333 Frame, L. T., 447(149), 455 Francois, J., 14(47), 22(47), 26(85), 27(85), 33, 40(85), 42. 45, 433(55), 453 Franenkel, D. G., 6(20), 42 Frank, E., 349(101), 357 Frank, R. N., 328(26), 332

469 Fredholm, B., 153(48), 173 Fredriksen, D. W., 388(32), 414 Fredrikson, G., 148(2, 4), 149(2, 4, 15, 17, 21, 25), 150(2, 21, 25, 29), 151(2, 21, 30, 34), 153(2), 154(2, 49), 157(2, 49, 65), 158(49), 159(2), 160(2, 49), 162(78), 166(2, 4), 167(78), 168(4), 169(34), 172, 173, 174 Freedland, R. A,, 271(171), 282 Freedman, M. L., 362(8), 373(95, loo), 377, 3 79 Freedman, S . B., (7), 354 Freedman, S. D., 301(172), 315 Frey, T., 292(58), 312 Fridkin, M., 349(144), 358 Friedman, B. A., 330(64, 68), 333 Friedman, J. A., 129(68), 145 Friedman, M., 375(128), 380 Friedman, P. A., 219(8, 15), 272(15), 273(15), 274(15), 275(15), 278 Friedman, S., 219(11), 278 Friedman, S. A,, 128(48), 144 Friend, D. S., 185(43), 211 Friis, R. R., 70(136), 71(136), 75 Fritz, P. J., 48(13), 68(13), 72 Fritz, R. B., 298(141), 314 Fromm, H., 18(51, 52), 33(169, 173), 42, 45, 443(124), 454 Froscio, M., 330(58), 332 Fucci, L., 19(54),42 Fuchs, S., 349(144), 358 Fujii, S., 69(115, 117), 70(117, 128), 74 Fujii, Y . , 55(56), 56(56), 73 Fujiki, H., 330(63), 333, 422(7), 451 Fujisaki, H., 407(150), 417 Fujisawa, H . , 221(26), 255(128, 129), 256(128), 257(138, 139), 258(128), 263(138, 139, 147, 148), 264(128, 129, 147, 148, 149), 274(185), 275(185), 276(196, 197, 198), 277(147, 148, 196, 198, 199), 278, 280, 281, 282, 294(93), 296(115), 300(166, 168), 302(166), 303(166), 306(220), 307(220), 313. 314, 315, 316 Fujiwara, K., 395(75), 415 Fukunaga, K., 296(111), 300(169), 303(169), 306(169), 307(169), 314, 315 Fukushima, T., 233(59), 279 Fuller, S. J., 88(54), 94 Funayama, S., 36(193), 46

470

AUTHOR INDEX

Funcke, H. J., 88(53), 94 Fung, B. K.-K.,377(146), 380 Fung, C. H., 124(15), 144 Furman, R. H., 149(13), 173 Furst, M., 29(131), 40(131), 44 Furuya, E., 5(3), 22(3, 61, 63), 28(3, 117), 29(3, 117, 129), 36(198), 37(61, 199), 41, 43, 44, 46 Fuxe, K., 302(179), 315 G

Gadasi, H., 406(146), 417 Gagosian, R. B . , 185(40), 211 Gaillard, D., 170(173), 176 Gal, E. M., 273(178), 275(178), 282 Galasko, G., 91(86, 92), 95 Galick, H. A , , 191(80), 212 Gallager, D. W., 275(189), 282 Gallagher, J. J., 187(49), 211 Gallis, B., 309(246), 317, 409(184), 417 Gallo, L. L., 169(169), 176 Gallo-Torres, H., 244(80), 279 Galvagno, M. A,, 437(86), 453 Gammeltoft, 165(141, 142), 176 Gancedo, C., 36(193, 195). 46, 434(59, 60), 453 Gancedo, J. M., 36(195), 46, 434(59, 60), 453 Ganson, N. J., 33(169, 173), 45 Garbers, D.-L., 349(96), 356 Garcia, M. C . , 342(141), 357 Garcia-Sainz, J. A., 90(80), 95, 162(83), I73 Garcia-Segura, L. M., 190(63), 191(63), 192(63), 194(63), 211 Garfield, M. K . , 302(186), 303(186), 315 Garfinkel, D., 85(45), 94 Garfinkel, L., 85(45), 94 Gargouil, Y.M., 337(10), 354 Garnache, A., 228(53), 279 Gamak, M., 425(21), 452 Gamier, D., 337(10), 354 Garreau, H., 68(112), 74 Garren, L. D., 169(163), 176 Garrison, J., 7(30), 37(30), 42 Garrison, J. C., 67(106, 107), 74, 229(56), 241(56, 67), 242(67), 279 Carte, S. J., 324(14), 331 Gasior, E., 431(43, 44). 452 Gates, R. E., 34(179), 45 Gaumert, R., 195(102, 103), 212

Gautvik, K., 349(101), 357 Gaylor, J. L., 194(96), 212 Gazzano, H., 143(137), 146, 165(144, 147), I 76 Gebhard, R. J., 206(171), 214 Geelen, M. J. H., 130(90, 91), 131(90, 91), 132(90, 91), 145, 199(136, 139), 200(139, 146, 1 5 3 , 201(146), 202(155), 203(139), 204(139), 205(139), 213. 214 Geelen, M. T.H., 128(47), 131(47), 132(47), 144 Geiger, P., 223(39), 278 Geogiadis, A., 244(80), 279 George, J. N., 390(47), 407(47), 408(47), 414 George, R. J., 256(136), 257(136), 264(136), 280 Gergely, J., 384(4), 413 Gerighty, M., 373(95), 379 Gerisch, G., 443(116, 118, 119, 120, 124), 444(125), 454 Gennann, P., 70(127), 74 Gerrard, J. M., 408(178), 417 Gerthoffer, W. T., 396(81), 397(81), 398(81), 415 Ghosh-Dastidar, P., 362(21, 23). 363(21), 377, 378 Gibbons, G. F., 376(137), 380 Gibbins, J. M., 130(82), 131(82), 132(82), 145 Gibbons, G. F., 200(204), 215 Gibson, D. M., 124(9, lo), 129(69), 130(90), 131(90), 132(90), 143, 145, 180(3, lo), 183(35), 185(35), 195(100, 107, 108, IlO), 196(10, 107, 108, IlO), 198(106, 108, IIO), 199(136, 139), 200(110, 139, 155), 202(155, 156, 157), 203(139, 156), 204(139, 205(139), 206(35), 207(35, 192), 208(191, 192). 210, 211, 212, 213, 214, 215 Gikner, J., 69(120), 70(120), 74 Gil, G., 182(33), 184(33), 190(201, 202), 197(33, 116, 119, 121, 122), 199(122, 134), 208(33), 211, 213, 215 Gilbert, W., 56(60), 57(60), 73 Giles, W., 338(12), 354 Gill, G. N., 169(163), 176, 293(69, 74). 313, 330(55, 66), 332, 333, 362(43), 363(43, 43,378 Gilligan, D. M., 449(161), 455 Gillim, S. E., 109(50), 113(50), 115(50), 120

AUTHOR INDEX

Gilman, A . G., 162(99), 175, 292(61), 312, 328(23), 332, 376(138), 377(143), 380 Giner, A., 46 Ginsberg, B., 273(178), 275(178), 282 Ginsburg, A . , 422(10), 451 Gispen, W. H . , 298(143), 300(163), 309(143, 237, 239, 240, 241), 314, 315, 317 Gitomer, B., 225(45), 278 Glass, D. B., 25(68, 69, 74, 83), 43 Glassman, A., 299(153), 301(153), 315 Glazer, A . N., 19(57), 42 Gliemann, J., 165(141), 176 Glikin, G. C., 435(71), 453 Glinsmann, W. H . , 303(231), 307(231), 316 Glomset, J . A . , 191(77), 192(77, 86), 193(77), 212 Glossman, H., 70(136), 71(136, 137), 75 Glowinski, J., 269(162), 281 Gobel, C., 293(73), 303(73), 313 Goelz, S. E., 301(175), 305(175), 315 Goetze, A . , 331(82), 333 Goff,C. G . , 422(9), 451 Goh, E. H . , 188(57), 193(57), 211 Goh, S. H., 375(113), 379 Goldberg, A. L., 103(23), 109(23), 120 Goldberg, N. D., 287(12), 293(12), 311 Goldberg, R. I., 182(34), 211 Goldbeter, A . , 162(95), 175 Goldenring, J. R . , 296(113), 297(127), 306(113), 307(113), 314 Goldfarb, M., 377(144), 380 Goldstein, J. L., 180(2), 181(18, 19, 20, 21, 22), 182(18, 19, 20, 21, 27, 30, 33). 183(20, 22, 36), 184(20, 22, 33, 36), 185(36), 186(44), 187(44, 53), 188(44, 53), 190(18, 22, 44,63, 64,69, 201, 202), 191(21, 30, 44, 63, 81), 192(19, 63, 921, 194(63), 195(104), 197(33), 198(104), 204(159), 207(20, 22, 36), 208(33, 36), 210, 211, 212, 214, 215 Goldstein, M., 251(99), 253(121), 260(146), 265(155), 268(146), 280, 281, 302(179), 315 Gomez, S. L., 438(91, 93), 453 Gonzalez, B., 296(113), 306(113), 307(113), 314 Gonzalez, C., 293(64), 312, 436(73), 453 Goodall, McC., 249(86), 279 Goodman, D. B. P., 337(6), 354 Goodman, D. S . , 170(183), 177, 191(78), 192(78), 193(78), 212

47 1 Goodman, H. M., 161(73, 74), 162(88), 163(88, IlO), 166(73, 74, 151, 152, 153), 173, 174, 175, 176 Goodwin, C. D., 195(101), 212 Goodwin, G. W., 200(147), 201(147), 214 Gorban, A . M. S., 151(36), 168(36), 169(36, 164), 173, 176 Gorden, P., 330(77), 333 Gordis, C., 310(253), 317 Gordon, A . , 300(164), 315 Gordon, A. S . , 302(193), 303(193), 316, 329(42, 4 9 , 332, 348(85, 88, 89, 93), 350(85), 351(116), 356, 357 Gordon, J., 438(95), 453 Gordon, P., 330(78), 333 Gordon, P. B., 208(194), 215 Gordon, R., 252(111, 115), 280 Gordon-Weeks, P. R . , 306(213), 316 Gorecka, A . , 386(25), 388(25), 414 Gospodarowicz, D., 190(65), 192(65), 193(65), 212 Goss, D. J., 364(60), 366(60), 378 Goto, S., 300(169), 303(169), 306(169), 307(169), 315 Gottsschalk, M. E., 33(167), 45 Gould, R. G., 188(58), 189(59), 200(148), 211, 214 Goumans, H . , 363(46), 364(57), 378 Gourdin, M.-F., 330(79), 333 Grab, D. J., 301(174), 315 Graber, S . G., 302(186), 303(186), 315 Grace, M., 362(21), 363(49), 377, 378 Graefe, M., 30(141), 41(141), 44 Grahame-Smith, D. G., 273(183), 282 Grahame-Smith, S. G . , 271(167), 281 Grankowski, N., 431(43), 452 Graves, D. J., 25(67), 43, 61(75), 62(75), 73 Gray, D. W., 219(20), 221(20), 223(20, 35). 226(20), 234(20, 3 3 , 235(35), 236(63), 238(35), 239(35), 244(35), 245(35), 246(35), 278, 279 Grayzel, A. I. P., 360(2), 377 Grazi, E., 32(161), 45 Green, H., 376(133), 380 Greenbaum, A . L., 128(58), 129(58), 144 Greene, H. L., 27(108, 109), 34(108, log), 44, 50(31), 65(31), 68(109), 72, 74 Greene, L. A . , 265(155), 281 Greene, L. E., 391(60), 414 Greengard, P., 25(72), 43, 50(30), 72, 198(131), 213, 253(123), 264(150),

472 265(155), 280, 281, 286(4, 5, 6, 7, 8). 287(9, 10, 11, 18, 19, 21, 22, 30), 288(18, 19, 21, 22), 289(7, 30, 32, 33, 34), 290(8, 18, 19, 21, 22, 30, 32, 33, 34, 35, 36, 37, 38), 291(35, 36, 37, 38, 39, 47, 48, 51), 292(37, 38), 293(33, 37, 38, 67, 68, 75, 76, 77), 294(35, 36, 89, 89a, 91, 92, 96, 98), 295(37, 99, 100, 101, 102, 103, 104, 105, 106), 296(37, 100, 102, 110, 117, 118, 119, 124), 297(35, 36, 110, 117, 124, 125), 298(35, 36, 96, 139, 147), 299(30, 37, 148), 300(160), 301(7, 8, 35, 36, 89, 89a, 91, 96, 100, 175), 302(7, 176, 177, 178, 182, 183, 190, 191), 303(5, 6, 8, 18, 19, 21, 22, 30, 37, 99, 100, 190, 191, 197, 198, 211, 212, 227, 228, 233, 234), 304(7, 8, 160, 197, 198), 305(32, 100, 175, 201, 202, 203, 204, 205, 206, 207, 208, 209, 210, 211, 212). 306(7, 36, 37, 110, 212, 218), 307(177, 178, 227, 228, 229, 230, 233). 308(8, 35. 36, 37, 96, 139, 147, 182, 233, 234), 309(36, 147, 233). 310(8, 256, 257), 311,312,313,314, 315,316,317, 329(46, 48, 49), 332, 336(1, 3), 338(12), 341(31), 343(56, 58), 347(58, 82, 83, 84), 348(82, 83), 349(84), 350(83, 84, 147), 351(84), 354,355,356,358, 421(2), 451 Gregg, R. G., 205(167), 214 Gregolin, C., 127(40), 144 Gribkoff, V. K . , 287(29), 289(29), 290(29),

312 Griffiths, H. S . , 386(16), 413 Grimaldi, P., 170(173), 176 Gros, F., 349(100), 357 Groschel-Stewart, U., 411(191), 418 Grosfeld, H., 362(22), 377 Gross, M., 362(27), 368(71), 369(71, 76),

378, 379 Grossberg, S. E., 375(125),380 Gruen, E., 289(33), 290(33), 293(33), 312 Grunberger,G., 330(77, 78), 333 Goodridge, A. G.,128(54, 56), 129(54, 56),

144 Goodson, J., 126(31), 132(100), 144,145 Gubler, U . , 442(111), 454 Guchhait, R. B., 126(35), 144 Guerriero, V., Jr., 401(111, Illa),416 Guerritore, A,, 434(61), 453 Giittler, F., 244(79), 279 Guguen-Guillouzo, C., 374(112), 379 Guidotti, A., 293(66), 313

AUTHOR INDEX Guiso, N., 444(129), 445(129),454 Gunderson, J. H., 448(150), 455 Gupta, N. K . , 362(16, 21, 23), 363(21, 48, 49), 377, 378 Gupta, R. K., 124(15), 144 Gupta, S. L., 375(124), 380 Gurin, S., 249(87), 279 Guroff, G.,254(126, 127), 271(126), 280 Guy, P., 62(79), 63(79), 73 Guy, P. S., 126(33, 38), 128(33), 139(33), 141(33, 38). 144 Guynn, R. W., 85(44), 94, 125(19), 144

H Haagsrnan, H. P., 200(146), 201(146), 213 Habenicht, A. J. R., 191(77), 192(77), 193(77),212 Hack, S., 247(81), 279 Hacker, M. L., 80(11, 14), 93 Haddox, M. K., 287(12), 293(12), 311 Hadjian, R. A., 348(92), 356 Haeberle, J. R . , 386(19), 396(77, 86), 398(86), 413,415 Haen, C., 91(83), 95 Hagiwara, S . , 340(28), 355 Hajjar, D. P., 170(182), 177 Hale, G.,80(9), 93 Halestrap, A. P., 89(69), 94, 103(25), 107(25), 109(25), 120, 127(44), 128(44), 129(44), 130(44, 76), 131(44, 76), 144, 145 Hall, E. R . , 57(64), 73. 207(179), 214 Hall,S. W., 328(38), 332 Hall, Z., 349(106, 107), 357 Hallberg, R. L., 449(166), 455 Halpain, S., 302(180), 315 Halperin, M. L., 125(18), 128(57), 129(57), 130(57), 144 Hamill, 0. P., 340(24), 355 Hamilton, I. R . , 422(11), 428(11), 452 Hamilton, J. A., 208(191), 215 Hamilton, L., 80(15), 93 Hamm, H. E., 328(31), 332 Hammer, J. A., 111, 406(147), 407(148, 149, 150), 417 Hamon, M., 269(162), 276(192, 194), 277(194), 281, 282 Han, A. C . , 200(147), 201(147), 214 Hanbauer, I., 254(125), 280 Hansbury, E., 187(46), 211 Hansford, R. G., 88(49), 94

473

AUTHOR INDEX Harada, K.,68(110), 74 Harano, Y.,48(16), 72 Hardeman, E. C., 181(16, 24). 182(16, 24, 29), 183(24), 191(24, 29). 210, 211 Harden, T. K., 321(3), 327(20), 331. 332 Hardesty, B., 360(3), 362(15, 25), 370(78), 371(78, 85), 377, 378, 379 Hardgrave, J. E., 200(149, 150), 201(151), 203(158), 214 Hardgrave, P. A., 184(38),211 Hardie, D. G., 62(79), 63(79), 73, 126(27, 33, 38), 127(27), 128(33), 130(79a, 92). 131(79a, 92, 114), 132(79a, 114), 133(92, IOI), 137(27, 91, 101, 110, 11 1, 112), 138(92, 101, 11I , 112), 139(33, 92, 101, 111, 112, 120), 140(92, 101, 111, 114, 120), 141(33, 38, 126), 142(131), 144, 145, 146

Hardman, J. G., 291(40), 292(40), 312 Hardwicke, P. M. D., 390(48, 49), 414 Harford, J., 331(86), 333 Haxing, H. U., 143(136), 146, 330(74), 333 Harper, A. E., 98(1), 99(1), 115(52), 116(59, 60,61), 119, 120, 121. 227(51), 279 Harris, D. G., 32(148), 41(148), 45 Harris, E., 191(74), 212 Harris, J. E., 253(122), 254(122), 280 Harris, R. A., 70(129), 74, 92(99), 95, 100(16), 101(16), 102(16), 104(16), 105(36, 37), 106(16, 36, 37), 107(16, 36, 37, 41). 109(16, 50), 113(50), 114(41), 115(41, 50), 116(37, 41), 120, 128(60), 129(60), 145, 199(135), 200(147), 201(147), 214 Harris, S. M., 305(202), 316 Hartl, F. T., 291(44, 45), 312 Hartman, F. C . , 20(58), 43 Hartmann, U., 88(52), 94 Hartshorne, D. J., 297(129), 303(129), 314, 384(10), 386(17, 17a, 25), 388(25, 36), 389(40), 391(53), 395(53, 68, 72, 72a), 396(89), 398(89), 413, 414, 415 Hartshorne, R. P., 351(114, 118, 119), 357 Harvey, E. V., 391(52), 414 Harwood, H. J. Jr., 198(125, 127), 206(172), 213, 214

Hase, J., 55(56), 56(56), 73 Hasegawa, H., 222(31, 34), 223(34), 224(34), 225(44), 226(34, 44), 227(44), 228(44), 240(44), 248(44), 262(44), 272(31), 278 Hashimoto, E., 297(133), 314 Haslam, R. J., 409(180), 417

Haston, W. S., 53(48), 59(48), 60(48), 61(48), 66(48), 73 Hathaway, D. R., 291(54), 297(54), 312, 386(19), 396(77, 86), 398(86), 399(104), 401(104), 413, 415, 41’6 Hathaway, G . M., 299(149), 303(149), 315 Hatton, J., 193(94), 212 Haubrich-Monee, T., 363(46), 378 Hauschildt, S., 115(53), 120 Havel, C. M., 185(43), 187(52), 192(52, 85), 202(52), 211, 212 Hawkins, R. A., 125(20), 126(20), 129(20), 130(20), 144 Hayaishi, O., 271(169), 273(179), 275(179), 281, 282

Hayashi, K., 169(165), 176 Haycock, J. W., 255(132), 256(132, 136). 257(136), 264(136), 265(155), 280, 281, 299(148), 315 Haystead, T. A. J., 130(79a), 131(79a), 132(79a), 145 Heald, S. L., 321(5), 331 Heaslip, R. J., 388(35), 414 Heffelfinger, S. C., 100(18), 102(18), 120 Hegardt, F. G . , 195(103), 196(113), 197(116, 118, 119, 121, 122, 124). 198(118), 199(122, 134), 212, 213 Heger, H. W., 129(70), 145 Hegstrand, L. R., 253(123), 260(145), 280, 28 I Heidmann, T., 347(79), 356 Heil, P. J., 7(27), 8(27), 42 Heimberg, M., 188(57), 193(57), 211 Heinemann, S., 353(127), 357 Heiniger, H. J., 189(60), 190(60),211 Heinrich, P. C . , 208(188), 215 Heinrikson, R., 35(186, 188), 46 Heinrikson, R. I., 25(138), 30(138), 40(138), 44

Heldan, E., 345(68), 356 Helenius, A,, 150(23), 173 Helfand, G. D., 219(19), 234(19), 236(19), 2 78

Heller, R., 150(27), 173 Heller, R. A., 152(42), 155(42), 173 Helmer-Matyjek, E., 265( 155), 281 Hemmings, B. A., 154(53), 165(150), 174, 176, 207(182), 214, 435(64, 65, 66), 453 Hemmings, H. C . , 25(72), 43 Hemmings, H. C . , Jr., 289(34), 290(34, 37), 291(37), 292(37), 293(37), 295(37),

474 296(37), 299(37), 302(178), 303(37, 228, 234), 396(37, 178), 307(228, 229, 230), 308(37, 234), 310(256), 312, 316, 317 Hempstead, B. L., 331(83, 84), 333 Hems, D. A,, 132(98), 145 Henderson, A. B., 370(78), 371(78), 378 Hendricks, S . A,, 300(159), 315 Hengstenberg, W., 425(29), 452 Henneberg, R., 200(203), 215 Hennebury, R. C . , 324(13), 331 Hennig, G., 89(68), 94 Henry, J.-P., 251(105), 280 Hensens, O., 191(74), 212 Henshaw, E. C . , 362(38, 41), 363(38, 41), 364(38, 41). 367(69), 373(102, 103), 374(103, 104), 378, 379 Hepp, K. D., 165(137), 175 Herbert, P., 371(83, 84). 379 Herman, R. H., 27(108, 109), 34(108, 109), 44, 50(31), 65(31), 68(109, I l l ) , 72, 74 Hermolin, J., 328(31), 332 Herr, B. E., 275(189), 282 Herrlich, P., 421(4, 5), 422(6), 428(37), 451, 452 Hers, H. G., 5(2), 6(24), 14(46, 47), 15(2, 49). 22(2, 46, 47, 49, 62), 23(49, 64), 24(49), 26(85), 27(85), 28(2, 116), 29(2, 24, 116, 119, 120, 121, 125), 32(165), 33(165), 37(46, 49), 40(85), 41, 42, 43, 44, 45, 48(2), 50(2), 54(49), 59(70), 63(85), 65(49, 70), 66(49), 71(2, 49), 72, 73, 74, 132(99), 145. 433(55), 434(56), 453 Hershey, J. W. B., 362(37), 375(123), 378, 380 Hershko, A., 207(184), 214 Hervy, C., 330(79), 333 Hery, F., 276(192, 194), 277(194), 282 Heschler, J., 338(15), 341(15), 354 Hesketh, J. E., 408(169), 417 Hess, G. P., 350(147), 358 Hess, P., 337(130), 357 Hewick, R. M.,347(74), 356 Hewlett, E., 165(130), 175 Heylen, A,, 29(119), 44 Heyworth, C., 165(143), 176 Hidaka, H., 399(107), 401(107), 402(107), 410(185, 187, 188), 411(187, 189), 415, 417, 418 Higashi, K., 296(1I l), 314 Higgins, M., 181(13), 193(13), 210 Higgins, M. J. P., 195(105),212

AUTHOR INDEX High, C. W., 405(130), 416 Hill, T. L., 385(13), 391(13), 392(13), 413 Hille, B., 342(41), 345(41), 355 Hinkins, S . , 388(36), 395(68, 72a), 398(36), 414, 415 Hinkle, P., 143(133), 146 Hiraoka, T., 89(62), 94 Hirata, F., 324(13), 331 Hirose, R., 299(151), 315 Hirsch, A. H., 150(28), 151(28), 156(28), 157(28), 170(28), 173 Hirschfield, J., 191(74), 212 Hirsch-Kauffmann, M., 422(6, 7), 428(37), 451. 452 Hissin, J. H., 143(133), 146 Hixson, C. S . , 430(40), 431(40), 444(40), 452 Hjelmquist, G., 50(32, 33, 3 3 , 58(35), 61(35), 72 Ho, R.-J., 152(37), 162(37, 81), 163(37, 81), 173, 174 Ho, Y. K., 181(19), 182(19), 192(19), 210 Hoar, C. G . , 56(59), 57(59), 73 Hoar, P . E., 396(85, 87, 88), 398(85, 87, 98), 402(112), 415, 416 Hochberg, A. A., 208(197), 215 Hodgson, D., 422(8), 451 Hokfelt, T., 302(179), 315 Hoeldtke, R., 250(96), 254(96), 275(96), 280 Horlein, D., 309(246), 317 Hofer, H. W., 11(34), 29(131, 135, 136), 30, 31(147), 32(148), 40(131), 41(148), 42, 44, 45 Hoffer, B. J., 293(79), 313 Hoffman, C., 191(74), 212 Hofmann, E., 29(126, 127), 31(143), 44 Hofmann, F., 293(68a, 72, 73), 303(72, 73), 313, 338(16), 339(16), 341(35), 342(139), 355. 357, 400(109), 405(109), 416 Hoffschulte, H., 36(190), 46 Holland, R., 130(92), 131(92, 114). 132(114), 133(92, IOl), 137(92, 101), 138(92, 101). 139(92, 101), 140(92, 101, 114), 145, 146 Holland, W. C., 252(106), 280 Hollenberg, C. H., 149(12), 152(12), 165(120), 173, 175 Holloway, D. E., 206(171), 214 Holm, C., 149(20, 25), 150(20, 2 3 , 151(20), 173 Holmes, S. L., 375(124), 380 Holroyde, M. J., 405(134), 416 Holuigue, L., 427(34, 35), 452

475

AUTHOR INDEX Holzer, H., 36(190, 192, 194), 41(196), 46, 207(180), 208(188), 214, 215, 434(57, 5 8 , 62, 63), 453 Honeyman, T. W., 90(77), 95. 163(110, I l l ) , 175 Honnor, R. C . , 162(92, IOl), 163(101), 164(116), 166(116), 174, I75 Hoogsteen, K., 191(74),212 Hopkirk, T. J., 50(22), 57(22), 72. 137(113), 141(122), 146, 165(148, 149), 176 Horchner, P., 360(2), 377 Horecker, B. L., 5(11), 14(48), 32(11, 149, 154, 156, 158), 33(168), 41, 42, 45, 209(199, 200), 215 Horlein, D., 375(113, 114), 379, 380 Home, P., 341(38), 355 Homer, K. A., 162(90), 169(90), 170(90), 174 Hosey, M. M., 29(137), 32(159), 35(159), 40(137), 41(159), 44, 45 Hott, J. W., 396(77), 415 House, P., 165(140),176 Houslay, M. D., 165(133, 134, 135, 143), 175, 176, 302(189), 303(189),315 Houston, M. E., 404(123), 416 Hovanessian, A., 372(92), 375(92), 379 Howard, C. F., 170(180),177 Huang, C.-K., 303(211, 212), 305(212), 306(212), 316, 329(47),332 Huang, C. Y., 310(256), 317 Huang, F. L., 303(231), 307(231), 316 Huang, J. J., 155(44), 156(44), 173 Huang, L., 91(92), 95 Huang, L. C., 91(85, 86), 95 Huang, Y . C . , 292(62), 312 Hucho, F., 78(4), 82(26), 83(4), 85(4), 93 Hudlicka, O., 404(125), 416 Hudson, T. H., 163(112), 175 Hue, L., 28(116), 29(116), 37(202), 44, 46, 48(2), 50(2), 54(49), 59(70), 63(85), 65(49, 70). 66(49), 71(2, 49), 72, 73, 74 Hiibner, G., 82(25), 93 Huebner, V. D., 442(109), 454 Hiilsmann, W. C., 48(10), 50(25), 55(25), 65(92), 72, 74, 170(178, 179), 176 Huesgen, A,, 443(124), 454 Huff, J., 191(74), 212 Huff, J . W., 192(93), 212 Huganir, R., 286(5), 303(5), 311 Huganir, R. L., 300(160), 303(197, 198), 304(160, 197, 198), 315, 316, 329(46, 48,

49), 332, 347(80, 82, 83, 84), 348(82, 83, 9 3 , 349(80, 84), 350(83, 84, 95, 147), 351(84), 356, 358 Hugh-Jones, T., 306(216), 316 Hughes-Fulford, M., 205(169), 214 Hughes, W. A,, 89(71), 90(74), 94, 95, 103(25), 107(25), 109(25), 120, 126(32), 130(77), 131(77), 132(77), 134(77, 104), 135(77, 104), 137(77), 139(77), 142(77), 144, 145 Huiatt, T. W., 387(29), 393(29), 414 Humble, E., 25(76), 35(187), 43, 45, 51(42), 53(42), 54(42), 55(42, 54, 5 3 , 56(54, 5 3 , 57(55, 61), 58(61), 60,61(61, 74), 62(74), 72, 73, 207(177), 214, 223(38), 278 Humphreys, J., 83(32), 94 Humphreys, J. S., 85(43), 86(46), 91(43), 92(46), 93(46), 94, 104(30), 107(30), 108(30), 109(30), 119(66), 120, 121 Huneeus, V. Q . , 191(76, 79, 80),212 Hunt, T., 362(9, 10, 12, 13, 14, 32), 365(64), 366(64), 369(74), 371(12, 74, 83, 84), 375(10, 12, 115). 377, 378, 379. 380 Hunt, V., 191(74),212 Hunter, T., 299(157), 303(157), 315, 330(52, 66), 332, 333, 350(109), 357. 375(115), 380 Hurley, J. B., 376(142), 377(143), 380 Hussey, C. R., 29(132), 40(132), 44 Hutcheson, E. T., 81(18), 93 Hutson, N. J . , 82(20), 88(56, 59), 89(20, 59), 93, 94 Hutson, S. M., 116(59, 60), 12'1 Huttner, W. B., 295(100), 296(100), 301(100), 303(100), 305(100, 202, 203). 313, 316 Huttunen, J. K., 150(26, 27), 152(39, 41, 43), 156(43), 173 Huvos, P., 373(103), 374(103), 379 Hylemon, P. B., 201(153), 214

I Ichihara, A., 188(55), 211 Ichikawa, Y., 408(171), 417 Ichiyama, A,, 271(169), 273(179), 275(179), 281, 282 Ihara, M., 191(82),212 Ikebe, M., 386(17, 17a). 386(26), 388(36), 389(41), 390(41, 4 9 , 391(53), 392(62),

476

AUTHOR INDEX

395(53, 68, 72, 72a), 396(45), 398(36), 399(107), 40 1(107), 402( 107), 406( 144), 413, 414, 415, 416, 417

Illiano, G., 165(136),175 Imamura, K., 48(11), 50(27), 68(11), 72 Imura, H., 308(236), 317 Inagaki, M., 399(107), 401(107), 402(107), 410(187), 411(187, 189). 415, 417, 418 Ingebritsen, T. S., 62(77, 78, 79, 82, 83), 63(77, 79, 82, 83), 64(83), 65(77), 73, 141(125, 126), 146, 154(50, 51), 174, 180(3), 182(28), 195(107, 108, 110), 196(107, 108, IIO), 198(107, 108, 110, 132), 199(139), 200(110, 132, 139, 155), 202(159, 203( 139), 204( 139), 205( 139), 210, 211, 213, 214, 225(47, 48), 278, 309(244, 245, 248), 310(255), 317, 336(4), 354

Inglis, R. J . , 440(101), 441(102), 454 Inohara, S . , 298(138), 303(138), 314 Inoue, H., 48(5), 49(5), 50(5), 72, 126(36), 134(36), 144 Inoue, M., 265(153a, 153b), 281, 298(135, 136), 314 Inui, M . , 397(96),415 Irisawa, H., 339(19), 355 Iritani, N., 128(53), 144 Irvine, R. F., 89(70), 94 Isaksson, O., 157(71), 158(71), 161(71), 166(71), 174 Ishibashi, H., 65(96), 66(96, 99). 67(99), 74 Ishihama, A., 424(20), 452 Ishikawa, T., 431(48, 49), 432(48, 49, 50, 51), 433(54), 435(67), 439(97, 98), 452, 453

Iwaki, M., 221(24), 222(24), 237(65), 278 Iwamoto, G. A., 404(123), 416 Iwasa, Y., 257(137), 265(137), 281 Iwashita, S., 330(67), 333

J Jabalquinto, A. M., 204(160), 214 Jackson, R. J., 362(12, 13, 14, 32), 365(64), 366(64), 371(12, 83, 84), 375(12, 115), 377, 378, 379, 380

Jacobs, S . , 91(89, 93), 95, 330(81), 331(85), 333

Jacquemin, C . , 165(138), 176 Jacubowitz, S . , 408(175), 417 Jagendorf, A. T., 427(32), 452

Jagus, R., 360(4), 363(52, 53), 377, 378 Jahn, R., 305(209), 316 Jakes, R., 386(15), 390(15), 413 James, M. E., 241(68), 279, 50(34), 53(34), 54(34), 57(34), 65(34), 66(34, 101). 67(101, 105), 72, 74 Jameson, L., 292(57, 58). 312 Janekovic, D., 422(7), 451 Jantzen, H. M., 448(158), 455 Jaramillo, F., 350(145), 358 Jarett, L., 91(87, 88, 90, 94), 95, 162(75), 165(126),174, 175 Jeanrenaud, B., 130(89), 131(89), 132(89), 145

Jedlicki, E., 225(43), 278 Jefferson, L. S . , 163(105), 175 Jenik, R. A , , 134(106), 146 Jenke, H., 181(16), 182(16), 210 Jenke, H. S . , 204(161), 214 JeMingS, K. R., 287(21), 288(21), 290(21), 303(21), 311, 345(63, 65), 346(65), 347(74), 356 Jensen, D., 151(32), 171(186), 173, 177 Jensen, D. F., 151(31), 170(31), 173 Jequier, E., 271(168), 272(173, 174, 176), 273(176), 275(173, 174, 176), 281, 282 Jeril, B., 148(1), 149(1), 150(1), 153(1), 172 Jerzmanowski, A., 441(105), 454 Jett, M.-F., 28(116), 29(116), 44, 63, 74 JimCnez DeAsba, L., 50(23), 72 Joh, T. H., 250(97), 258(133), 261(133), 262(133), 265(133), 280, 302(184), 303(184), 315 Johansson, O., 302(179), 315 John, M., 386(15), 390(15), 413 Johnsen, D. E., 241(67), 242(67), 279 Johnson, E. M., 25(77), 43, 291(51), 312 Johnson, G. L., 323(12), 331 Johnson, J. D . , 405(134), 416 Johnson, M. A., 408(172), 417 Johnson, R. A., 7(28), 42 Johnson, W. A., 100(10), 103(10), 119 Johnson, W. H., 406(138), 416 Jolli?s, P., 299(152), 309(241),315, 317 Jones, A. W . , 403(114), 416 Jones, R., 407(150), 417 Jones, R. G . , 130(86), 131(86), 145 Jones, T., 221(25), 278 Joseph, S. K . , 88(50), 94 Joshua, H., 191(74), 212 Joy, L. L., 151(31), 170(31), 173

477

AUTHOR INDEX Judewicz, N. D., 435(70, 71), 453 Juliani, M. H., 438(92, 93, 94), 439(96), 444(127), 446(142, 145), 453. 455 Julien, J.-P., 299(155), 303(155), 315 Jungas, R. L., 130(95), 145, 166(157), 167(157), 176 Jungerman, K., 48(17), 72 Jungmann, R. A., 442(107), 446(107), 454 Jurin, R. R . , 188(56), 211

K Kacini, M. R., 31(147), 32(148), 41(148), 45 Kaczmarek, L. K., 287(21, 22), 288(21, 22), 290(21, 22), 303(21, 22), 311, 341(31), 342(45), 345(63, 64,65, 66). 346(65), 347(74), 355, 356 Kaempfer, R., 361(6), 377 Kagimoto, T., 27(112), 28(112), 44 Kahane, I., 411(191), 418 Kahn, A., 48(4, 8), 49(4), 50(26), 56(58), 68(4, 112), 69(114, 116), 71(4), 72, 73, 74 Kahn, C. R., 143(136), 146, 330(54, 73, 74, 75, 80), 332, 333 Kaibuchi, K., 242(71, 72). 279, 298(146), 314 Kakinuma, S . , 445(139, 140), 454 Kakiuchi, S., 253(119), 280, 296(112), 309(251), 314, 317, 397(96), 415 Kaku, K., 70(128), 74 Kamata, T., 393(64), 415 Kaminsky, E. A., 388(37), 414 Kamm, K. E., 396(76), 397(90, 91, 92), 398(92), 403(76), 409(90), 415 Kanamura, K., 399(107), 416 Kanda, K., 296(112), 314, 397(96), 415 Kandel, E. R., 287(18, 19, 20), 288(18, 19, 20), 290(18, 19, 20), 303(18, 19, 20), 310(258), 311, 317, 342(44), 343(52, 53, 54, 55, 56, 58, 59), 344(52, 59, 60, 61), 347(58, 61, 73), 355, 356 Kandutsch, A. A., 189(60, 61), 190(61, 70), 191(71), 211, 212, Kane, J., 181(14), 210 Kaneko, I., 182(27), 211 Kaneko, T., 69(115, 117), 70(117, 128), 74 Kang, E. S . , 219(8), 278 Kanof, P., 291(47), 312 Kapatos, G., 259(144), 260(144), 261(144), 265(144), 267(144), 281 Kaplanski, D. A,, 368(71), 369(71), 379

Kappelman, A. H., 219(15), 272(15), 273(15), 274(15), 275(15), 278 Karadsheh, N. S . , 11(35), 42 Karell, M. A,, 328(31), 332 Karlin, A , , 329(40), 332 Karlsson, F. A,, 330(73, 74), 333 Karnieli, E., 143(133), 146 Karpatkin, S., 373(100), 379 Kashiba, A., 299(151), 315 Kass, R. A , , 339(18), 340(18), 355 Kassis, S., 303(200), 304(200), 316 Kasten, T. P., 27(89), 43 Kasuga, M., 143(136), 146, 330(54, 73, 74, 75), 332, 333 Katada, T., 376( 140), 380 Kato, T., 272(175), 282 Katoh N., 294(95, 97), 298(97, 145), 313, 314 Katz, A. M., 341(36, 37), 355 Katz, I., 251(102), 280 Katz, J., 40(204), 46, 71, 75 Katz, N., 48(17), 72 Kaufman, S., 218(1, 2, 3, 4, 6), 219(3, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 21, 22), 221(1, 23, 24), 222(12, 23, 24, 28, 29, 30, 31, 32, 33, 34), 223(33, 34). 224(34, 36, 40, 41), 225(23, 33, 42, 43, 44, 45), 226(34, 44), 227(33, 42, 44,50), 228(42, 44, 52). 230(42, 27), 231(50), 232(42, 50), 233(22, 23), 234(7, 21), 235(60), 236(7, 29, 30, 33, 60), 237(60, 65), 238(21, 33, 42), 239(33, 50, 60), 240(42, 44), 241(42, 50), 242(60), 243(22, 42, 60, 74, 7 3 , 245(50), 248(2, 3, 4, 44, 84), 249(12, 13, 14, 16, 90), 250(29, 96), 251(101, 102), 252(101), 253(101, 124), 254(96, 124), 255(124), 258(124), 259(124, 144), 260(144), 261(144), 262(44, 124), 265(144), 267(144), 269(124), 272(6, 15, 16, 31, 33, 177), 273(10, 14, 15, 180, 181), 274(7, 15, 181), 275(12, 15, 96), 277, 278, 279, 280, 281, 282 Kavaler, J., 408(175), 417 Kawahara, Y . , 257(137), 265(137), 280 Kawamoto, S., 410(187), 411(187, 189). 418

Kawamura, M., 151(31), 170(31), 173 Kawka, D. W . , 192(93), 212 Kay, J. E., 374(105, 106), 379 Kazazis, D. M., 192(93), 212 Keay, S., 375(125), 380

478 Keech, D. B., 124(13), 143 Kehinde, O., 376(133), 380 Kehr, W., 277(200), 282 Keim, P., 35(186), 46 Keith, M. L., 197(115, 123), 213 Kelleher, D. J., 323(12), 331 Keller, T. C. S . , 111, 407( 163, 41 7 Kellet, G. L., 29(132), 40(132), 44 Kellogg, J., 91(86, 92). 95 Kelly, P. T., 296(117), 297(117, 128), 306(128), 314 Kelly, R. B., 300(171), 315 Kemp, B. E., 25(67, 78, 79), 43, 61, 62(75), 67(102), 73, 74, 247(83), 279, 386(15), 390(15), 413 Kemp, R., 27(105), 44 Kemp, R. G., 25(138), 27(97, 98), 29(133, 137), 30(138), 40(133, 137, 138), 43, 44 Kendrick-Jones, J., 384(6, 8), 386(15), 387(29), 388(8), 390(15), 393(29, 67). 395(70, 74), 406(8), 407(163), 413, 415, 417, 443(123), 454 Kennedy, M. B., 198(131), 213, 295(101, 102), 296(102, 109), 297(109, 126), 306(109), 313, 314 Kennelly, P. J., 180(9), 196(9), 197(123), 198(126), 200(9), 210, 213 Kerbey, A. L., 78(5), 81(19), 82(20, 21), 84(34), 88(57, 58, 59), 89(20), 93, 94 Kerlavage, A. R., 321(6), 331 Kemer, N., 436(80), 437(87, 88), 439(99), 453

Kerr, I. M., 372(92), 375(92, 120, 129), 379, 380

Kenick, W. G . L., 396(85, 87, 88, 89), 398(85, 87, 89, 98), 402(112), 406(136), 415, 416

Kessar, P., 89(67), 94 Kessler, R., 29(127), 44 Kester, M. V., 236(61), 279 Keuttel, M. R., 442(107), 446(107), 454 Khailova, L. S . , 82(25), 93 Khandelwal, R. L., 422(11), 428(1I), 452 Khoo, J. C., 148(3), 149(3), 151(31, 32), 152(44, 45), 153(48), 155(44, 56, 57), 156(3, 44), 160(3), 162(80), 170(31, 173, 185), 171(186),172, 173, 174, 176, 177 Khorana, H. G., 377(145), 380 Kibler, R. F., 298(142), 314 Kidokoro, Y.,353(127), 357 Kiechle, F. L., 91(89), 95, 165(126), 175

AUTHOR INDEX

Kiehart, D. P., 407(158), 417 Kiener, P. A., 69(I 19, 120), 70( 120), 74 Kiesch, J., 191(74), 212 Kii, R., 297(133), 314 Kikkawa, U., 242(71), 279, 298(137, 138, 146), 314 Kikuchi, T., 208(189), 215 Killilea, S. D., 195(109),213 Kilpatrick, B. F., 323(10), 324(10), 331 Kilpatrick, D. L., 91(95), 95 Kim, K.-H., 124(5), 126(5, 24, 25), 127(25, 41), 129(66, 67, 71), 130(75, 78. 93), 131(25, 78, 93), 132(25, 93), 134(102, 1031, 139(117, 118, 119), 140(41, 117, 118, 119), 141(127), 142(130), 143, 144, 145, 146, 200(144), 201(144), 213 Kim, M., 134(106), 146 Kimchi, A., 375(116, 128), 380 King, A. C., 330(63), 333 King, L., 287(16), 311, 330(51), 332 King, L. E. Jr., 70(126), 74 King, L. J., 298(134), 314 King, M. M., 310(256), 317 Kinnunen, P. K., 151(33), 173 Kirchberger, M. A., 341(36), 355 Kirshner, N., 249(86), 279 Kishimoto, A., 257(137), 265(137, 153a, 153b), 281, 298(135, 136, 137), 314 Kita, T., 187(53), 188(53), 192(92), 211, 212 Kitajima, S . , 5(16), 6(16), 7(16), 12(16, 43), 13(43), 14(43), 15(16, 43), 20(58), 21(16, 59), 22(16), 23(43), 30(140), 41, 42, 43, 44 Kityuma, S., 29, 44 Klausner, R. D., 331(86), 333 Klee, C. B., 291(54), 295(108), 297(54), 310(255), 312, 314, 317. 336(4), 354 Klein, C., 444(127), 446(145), 447(146, 149b), 454, 455 Klein, G. J., 130(87), 131(87), 145 Klein, H. P., 124(8), 143 Klein, M., 343(52, 53), 344(52, 60), 355, 356 Klein, P., 447(149a), 455 Kleine, L. P., 309(238), 317 Kleinschmidt, A. K., 127(40), 144 Kleinsek, D. A,, 201(152), 204(160), 214 Klier, F. G., 353(127), 357 Klug, A., 440(100), 453 Klumpp, S . , 449(160), 455 Knapp, S., 276(191), 282 Kneer, N., 34(177), 45 Kneer, N. M., 12(39, 40), 42

AUTHOR INDEX Knight, D. E., 308(235), 317 Knitson, G. S . , 375(123), 380 Knoppel, E. M., 192(89),212 Knowles, A., 328(22), 332 Knowles, J., 13(44), 42 Kobashi, K., 55(56), 56(56), 73 Koepfer-Hobelsberger, B., 90(78, 79), 95 Koeppe, R. E.,50(24), 54(24), 72 Koerner, T. A. W., Jr., 11(37),42 . Kohama, K., 397(95), 415 Kohl, E. A., 57(62), 65(93), 73, 74 Kojima, K., 249(93), 279 Kokubun, S., 340(21, 22), 355 Koller, T., 440(100), 453 Kondo, S., 251(99), 280 Konieczny, A., 362(44), 363(44, 53), 364(44), 366(44), 378 Konigsberg, P . W. H., 307(230), 316 Konigsberg, W. H., 25(72), 43 Konijn, T. M., 442(114), 454 Kono, N., 27(94, 95), 43 Kono, T., 165(122, 124, 127), 175 Kopin, I. G., 252(113), 280 Koppel, D. E., 349(98), 356 Korn, E. D., 382(1), 406(140, 145, 146, 147). 407(148, 149, 150, 151, 152, 153, 154, 155, 156), 408(1), 413, 416, 417, 443(121, 122). 454 Kornberg, H. L., 425(22), 452 Koshland, D. E., 423(16, 17), 425(24, 25, 27), 452 Koshland, Jr. D. E., 162(95), 175 Kosower, E. M., 362(10), 377 Kosower, N. S . , 362(11), 377 Koster, J. F . , 48(10), 50(25), 53(46), 55(25), 57(63), 65(92), 72, 73 Kostyuk, P. G., 287(26), 289(26), 290(26), 303(26), 311, 340(25, 26, 30), 355 Kotagal, N., 91(90), 95 Kotake, H., 339(19), 355 Kothari, H. V., 170(181), 177 Kountz, P., 3 5 , 6, 17), 7(17, 31), 8(31), 13(5, 6, 17), 14(5, 6, 17), 15(5, 6, 17, 31), 17(5, 6), 18(5), 22(5, 6, 17, 31), 24(5, 6, 31). 25(31), 28(5, 6). 29(5, 6), 30(6), 37(5, 6), 40(6), 41. 42 Kountz, P. D., 34(176), 36(176), 45 Kovanen, P. T., 186(45), 187(45), 188(45), 190(45), 191(45),211 Kowaloff, E. M., 126(26), 132(26), 134(26), 135(26), 137(26), 144

479 Kowalski, A., 143(137),146, 165(144, 147). 176 Krakower, G. R., 141(127), 142(130), 146 Kramer, G., 360(3), 362(15, 25). 371(85), 377, 378, 379 Kranias, E. G., 405(134),416 Krause, S., 406(138), 416 Krebs, E. G., 25(67, 68, 69, 74, 78), 43, 61(75), 62(75), 71(139), 73, 75, 152(40), 153(40), 156(63), 162(102), 163(102), 173, 174, 175, 293(64), 312, 343(57), 356, 400(108), 409(184), 416, 417, 420(1), 430(40), 431(40), 436(73), 444(40), 451. 453 Krebs, H. A., 48(14), 72 Kresze, G.-B., 80(10), 93 Krettaz, M., 330(54), 332 Krishnan, I., 375(130),380 Kristiansen, K., 449(165), 455 Kristjansson, G. I., 309(239), 317 Kriiger, A., 449(165), 455 Krueger, B. K., 294(89), 301(89, 89a), 313 Kruep, D., 27(89), 43 Kruijt, J. K., 57(63), 65(92), 73, 74 Krustik, E., 11(34),42 Kruyt, J. K., 50(25), 55(25), 72 Krystek, E.,29(136), 44 Kuczenski, R. T., 251(100), 280 Kuczmarski, E. R., 406(142), 407(142), 417, 443(117), 454 Kudlichki, W., 431(43, 44),452 Kuduz, J., 28(118), 44 Kuehn, G. D., 207(183), 208(183), 214, 442(110, 111, 112). 454 Kuffler, S. W., 285(2), 286(2), 311 Kuhl, C., 244(79), 279 Kuhn, D. M., 276(195), 277(195), 282, 302(185), 315 Kuhn, H., 328(24, 28, 30, 32, 38), 332 Kulczycki, A., Jr., 331(83, 84). 333 Kumon, A , , 408(170), 417 Kunau, W. H., 169(170), 171(170), 176 Kuntz, M. J., 200(147), 201(147), 214 Kuo, J. F., 50(30), 72, 257(140), 281, 291(39), 293(67, 78), 94(95, 97), 298(97, 140, 141, 142, 145), 307(142, 224), 310(257), 312, 313, 314, 316, 317, 421(2), 451 Kuri-Harcuch, W., 376(134), 380 Kuroda, M., 182(26), 191(26),211 Kuron, G., 191(74),212

480

AUTHOR INDEX

Kursky, M. D., 287(26), 289(26), 290(26), 303(26), 311, 340(30), 355 Kurtz, J. W., 208(198), 215 Kushmerick, M. J., 404(122), 416 Kuzma, G., 287(28), 289(28), 290(28), 303(28), 312, 342(49), 346(49), 355 Kuznicki, J., 407(153, 155, 156), 417 Kuzuya, H., 308(236), 317 Kyte, J., 184(37), 211

L Lacy, W. W., 116(63), 121 Laemmli, U. K., 103(27), 120, 441(103), 454 Lafontan, M., 90(81), 95 Lai, E., 162(98), 175 Lai, J. C . K., 309(250), 317 Lai, Y., 264(150), 281, 295(104, 105), 296(110, 119, 124), 297(110, 124), 306(1lo), 313, 314 Lakshmanan, M. R., 125(20), 126(20), 129(20), 130(20), 144 Laloux, M., 14(47), 22(47), 42 Lambert, B., 165(138), 176 Lampson, L. A., 351(116), 357 Lan, S.-F., 181(17), 190(17, 66, 68), 192(89, 90, 91), 202(17), 210, 212 Lane, M. D., 124(1), 126(1, 22, 23, 3 3 , 127(1, 23, 39, 40, 42, 43, 4 3 , 128(1, 43, 45, 61, 62). 129(61, 62, 63), 132(61, 62, 63), 134(105), 143, 144, 145 Langan, T. A., 60(72), 73, 255(130), 262(130), 265(156), 267(156), 280, 281, 441(106), 442(108), 454 Lange, Y., 185(44),211 Lansman, J. B., 337(130), 357 Lapidot, Y., 375(128), 380 LaPorte, D. C . , 425(25, 26, 27), 452 Lardy, H. A., 12(39, 40), 27(100, 102), 34(177), 42, 43, 44, 45, 208(197), 215 Lamer, J., 91(85, 86, 92, 9 9 , 95 Larsen, A. D . , 436(77), 453 Larson, R. E., 90(76), 95 Lasek, R. J., 299(154), 303(154), 315 Latorre, R., 342(42), 345(42), 355 Latshaw, S. P., 25(138), 30(138), 40(138), 44 Lattke, H. K., 129(64), 145 Lau, K. S . , 78(7), 92(96, 98), 93, 95. 99(3), lOO(3, 19), 101(19), 102(21), 103(21, 26), 104(3, 19, 28, 31), 105(19, 31), 106(19), 107(19, 26), 109(19, 26, 28, 4 3 , llO(21,

4 3 , 11 l(3, 21, 45), 112(21, 4 3 , 113(21), 119, 120 Lauris, V., 330(80), 333 Lavin, T. N., 321(5), 331 Lavis, V. R., 165(121), 175 Law, M. Y., 307(221), 316 Lawing, W. J., Jr., 409(181), 417 Lawson, R., 92(97), 95, 104(29, 32, 33), 105(32, 33, 34), 107(33, 38), 109(29), 110(38), I20 Lawson, T. W. R., 6(21), 31(21), 40(21), 41(21), 42 Lazar, M. A,, 250(98), 258(98, 142), 260(98), 270(163), 271(166), 280, 281 Lazdunski, M., 337(133), 342(140), 351(113, 117), 357 Leach, S. J., 25(79), 26(79), 43 Lear, S. R., 198(133),213 Leavis, P. C . , 384(4), 413 Lebleu, B., 372(91), 375(91, 131), 379, 380 Lederer, B., 6(24), 29(24), 42 Lee, E. Y. C., 63(88), 74, 155(56),174, 195(109),213 Lee, F., 268(160), 281 Lee, F.-L., 266(157), 267(157), 268(157), 269(157), 281 Lee, H. S., 195(108), 196(108), 198(108), 213 Lee, F. T., 151(30),173 Lee, K.-H.,126(25), 127(25, 41), 129(67), 130(75, 78), 131(25, 75, 78), 132(25, 7 3 , 140(41), 144, 145 Lee, L.-S., 330(56, 57), 332 Lee, R. H., 300(162), 303(162), 315 Lefkowitz, R. J., 303(199), 304(199),316, 320(1, 2), 321(4, 5 , 8), 323(9, 10, ll), 324(10, 16, 17), 325(18), 327(19, 21), 331, 332 Legon, S., 362(13, 14). 377 Lehman, W., 384(5, 12), 391(12), 406(12), 413 Lehninger, A,, 243(73), 279 Leibach, F. H., 29(122), 44 Leichtling, B. H., 444(128, 134, 137), 445(128, 139, 140), 446(141), 447(147, 148), 454, 455 Leijten, L., 192(84), 193(84), 212 Lekawa, M. E., 287(29), 289(29), 290(29), 312 Lemmon, S. K., lOO(11, 20), 101(11), 104(20), 109(11), 119, 120

AUTHOR INDEX Lemongello, D., 181(14),210 Lemos, J. R., 347(75, 76), 356 Lengyel, P., 372(91), 375(91, 118), 379, 380 Lenke, R . R . , 243(76), 279 Lenox, R. H . , 302(186), 303(186), 315 Lent, B. A . , 127(41), 139(117, 118, 119), 140(41, 117, 118, 119), 144, 146 Leonard, C. S., 289(32), 290(32), 305(32), 312 Leoni, S., 205(168, 169), 214 LePeuch, C. J., 341(32), 355 Lepreau-Jose, M. J., 130(90), 131(90), 132(90), 145 Lerch, K., 431(46), 452 Lerner, P., 258(143), 260(143), 281 Leroux, A,, 362(33), 364(33), 378 Leroux, A . L., 369(72), 379 Letendre, C. H.,254(126, 127), 271(126), 280

Leuthje, J., 115(53), 120 Levenberg, B., 218(3), 219(3), 248(3), 277 Levey, G. S . , 199(138),213 Levin, D., 373(96), 375(96), 379 Levin, D. H., 362(24, 28, 29, 30, 40), 363(40), 364(40, 59, 62), 365(65), 366(62, 65), 367(30, 59), 368(59, 62), 369(59, 72, 751, 370(30, 79, 80, 81), 371(28, 29, 81, 82), 372(28, 86, 87, 88, 89). 374(11I), 375(28), 378, 379 Levinson, S. R . , 351(110, 112), 357 Levitan, 1. B., 287(23, 24, 25), 288(23, 24, 2 3 , 289(25), 290(23, 24, 25), 303(23, 24, 2 3 , 311, 342(46, 47, 48), 345(46, 47, 70, 711, 346(48), 347(72, 75, 76), 355, 356 Levitt, M., 249(91), 252(91), 279 Levin, P., 293(77), 313 Levitz, M., 99(6), 100(6), 119 Levy, H., 243(76), 279 Levy, L. K., 163(110, 111), 175 Lewis, J. A . , 373(103), 374(103), 379 Lewis, P. N., 60(73), 73 Lewis, R . M., 448(157), 455 Li, H.-C., 309(242), 317, 421(3), 451 Li, S. Y.,163(112), 175 Liang, T., 374(107, 108), 379 Liao, S., 374(107, 108). 379 Liddle, P. E., 29(132), 40(132), 44 Liebeman, D., 375(126), 380 Liebman, P. A . , 328(33, 34, 35, 36), 332 Limanek, J. S . , 191(72), 212 Lim Tung, H. Y.,119(65), 121

Lin, R. C., 192(83), 193(83), 212 Lincoln, T. M., 399(105), 400(105), 401(105), 402(105), 416 Lindahl-Kiessling, K., 374(106), 379 Linden, D. C . , 349(102), 357 Lindstrom, J., 347(81), 349(81), 356 Linn, T. C . , 78(3), 80(15), 82(26), 93 Liscum, L., 181(19, 20), 182(19, 20, 33). 183(20, 36), 184(20, 33, 36), 185(36), 192(19), 197(33), 207(20, 36), 208(33, 36), 210, 211 Litosch, I., 163(112), 175 Little, S. A , , 166(155), 176 Livesy, G., 116(58), 117(58), 121 Ljugstrom, O., 50(32, 33). 51(37), 52, 53, 54(43), 59(69), 60(37, 69), 65(94), 66, 72, 73, 74 Llinas, R.,289(32), 290(32), 305(32), 312 Lloyd, T., 251(100, 104), 252(100), 253(100, 124), 254(124), 255( 124), 258( 124), 259(124), 262(124), 269(124), 280 Loach, P. A., 428(36), 452 Lobos, D. V., 187(52), 192(52), 202(52), 211 Lockfeld, A. J., 250(98), 258(98), 260(98), 280 Lodish, H . F., 373(93), 379 Loeb, J. N., 162(94), 175 Loffler, G., 88(52, 54), 89(68), 94 Logel, J., 190(67),212 Lohmann, S. M., 291(48, 56), 293(75, 76), 312, 313 Lolley, R. N., 300(162), 303(162), 315 Lombet, A., 351(113, 117), 357 Lonberg, N., 56(60), 57(60), 73 London, C., 162(92, lo]), 163(101), 164(116), 166(116), 174. 175 London, I. M., 360(1, 2), 362(9, 16, 19, 24, 26, 28, 29, 30, 31, 33, 40), 363(19, 40), 364(33, 40, 58, 59, 62), 365(65), 366(62, 65, 68), 367(30, 58, 59). 368(58, 59, 62, 68), 369(59, 72, 73, 75, 77), 370(30, 77, 79, 80, 81). 371(28, 29, 81, 82), 372(28, 86, 87, 88), 373(94, 96, 97), 374(68, 104, 1 I l ) , 375(28, 96), 377, 378, 379 Lornitzo, F: A . , 134(106), 146 Loten, E. G . , 162(85, 86, 87), 163(86, 87), 165(117, 123), 174, 175 Louw, W., 401(110a), 116 Lovell-Smith, C. J., 165(119), 175 Lovenberg, E., 271(168), 281 Lovenberg, W., 254(125), 257(141),

482

AUTHOR INDEX

258(143), 260(143), 265(154), 272(173, 174, 176), 273(176), 275(173, 174, 176, 186), 276(195), 277(195), 280, 281, 282, 294(94), 302(185), 303(185), 313, 315 Lovett, J. S . , 438(89), 453 Lowel, M., 204(161), 213 Lowenstein, J. M., 126(36), 134(36), 144 Lowry, 0. H., 27(106), 44 Lowey, S . , 387(30), 390(45b, 51), 393(30, 65), 394(65), 395(65), 414, 415 Lowry, 0. H., 298(134), 314 Lubs-Haukeness, J., 447(146, 149b), 455 Luby, L., 28(117), 29(117), 44 Luby, L. J., 29(134), 40(134), 44 Lucas, T. J., 401(110a), 416 Lucero, H. A,, 427(34, 3 9 , 4 5 2 Lukas, T., 7(31), 8(31), 15(31), 22(31), 24(31), 25(31), 42 Lund, P., 116(58), 117(58), 121 Luppis, B., 14(48), 42 Luskey, K. L., 181(18, 19, 21, 22), 182(18, 19, 21, 30, 33), 183(22, 36). 184(22, 33, 36), 185(36), 190(18, 22, 201, 202), 191(21, 30), 192(19), 197(33), 207(22, 36), 197(33), 207(22, 36), 208(33, 36), 210. 211, 215 Lux, H. D., 337(129), 357 Ly, S., 130(93), 131(93), 132(93), 145 Lynch, G., 300(163, 165), 315, 408(174), 417 Lynen, F., 171(188), 177 Lynham, J. A , , 409(180), 417 Lynn, W. S . , 91(82), 95 Lysz, T. W., 276(193), 277(193), 282

M Ma, G. Y . , 132(98), I45 McAdam, W. J., 247(83), 279 Macaulay, S . L., 165(126), 175 McCaffrey, P. G . , 330(68), 333 McCarroll, R., 445(138), 454 McCarthy, B. J., 185(43), 211 McCormack, J. G . , 88(48), 89(63, 64,65, 73), 90(73, 75). 91(73), 94, 95, 130(80, 82), 131(80, 82), 132(80, 82). 145 McCreery, M. J., 194(99), 212 McCully, V., 57(64), 66(99), 67(99), 73, 74, 207(179), 214 McCune, R. W., 293(74), 313 McCune, S. A , , 188(56), 211

MacDonald, R. J., 181(18), 182(18), 190(18), 210 McDonald, J. M., 91(87), 95, 162(77), 166(77), 167(77), 174 MacDonnell, P. C., 254(126, 127), 271(126), 280 McDowell, J. H., 328(28), 332 McGany, J . D., 40(204), 46, 128(55), 130(55), 144 McGrane, M., 5(1, 4, 5, 6), 12(41), 13(5, 6), 15(4, 5, 6 ) , 176, 61, 18(5), 22(5, 61, 24(4, 5, 6), 28(1, 4, 5, 6), 29(5, 6), 30(6), 32(164), 33(164), 34(182), 35(182), 37(1, 4, 5, 6, 41), 40(6), 41, 42, 45, 200(141), 201(141), 213 McGrane, M. M., 33(166), 34(166, 176), 45, 54(50), 62(50), 72 MacGregor, J. S . , 35, 46 McGuinness, T. L., 198(131), 213, 264(150), 281, 289(32), 290(32), 295(102, 104, 105), 296(110, 117, 124), 297(110, 117, 124, 125), 305(32), 306(110), 312, 313, 314 McGuire, D. M., 187(47), 211 McGuire, J. S . , Jr., 296(113), 297(127), 306(113), 307(113), 314 Machicao, F., 143(138), 146 McIlwain, H., 253(119), 280 Mackall, J. C., 126(22), 144 Mackerlova, L., 257(140), 281, 294(97), 298(97), 313 McKinner, M., 294(87), 313 McLean Grogan, W., 170(172), 176 McNamara, J. 0..309(252), 317 McNeillie, E. M., 130(83), 131(83), 132(83), 142(129), 145, 146 McPherson, J., 375(113), 379 McPherson, J. M., 375(114), 380 McQueen, C. A., 251(103), 280 Madison, D. V., 342(50, 143), 346(50, 143), 347(50), 355, 357 Maeno, H., 291(51), 312 Magilen, G., 300(164), 315 Magun, B. E., 330(62), 333 Maher, J. F., 84(38), 94 Mahler, H. R., 309(238), 317 Mahoney, E. M., 170(185), 171(186), 177 Maia, J. C. C., 438(91, 92, 93, 94), 439(96), 446(142), 453, 455 Majerfeld, I. H., 444(128), 445(128, 139, 140), 454

AUTHOR INDEX Majerus, P. W., 409(181), 417 Majumdar, A . , 362(16), 377 Majumder, G. C . , 449(164), 455 Mak, A . , 405(131), 416 Mak, A . S . , 405(128), 416 Makino, H., 165(124), 175 Makk, G., 257(141), 265(154), 281 Malaise-Lagae, F., 29( 119), 44 Malaisse, W. J., 29(119), 44 Malano, J., 36(193), 46 Malchow, D., 443(115, 116), 454 Malencik, D. A . , 396(87), 398(87), 415 Malenka, R. C., 342(143), 346(143), 357 Maleszewski, M., 441(105), 454 Makinson, A. M., 291(46), 312 Mallorga, P., 324(13), 331 Malloy, P. J., 425(23), 452 Malviya, A. N., 302(188), 303(188), 315 Manai, M., 423(18), 424(18), 452 Manalan, A . , 310(255), 317, 336(4), 354 Manchester, K. L., 364(61), 366(61), 378 Manclark, C. R., 376(141), 380 Mandel, P., 408(169), 417 Mandell, A. J., 251(100), 276(191), 280, 282 Manganiello, V. C . , 162(82), 163(82), 165(118, 119, 129, 130, 131, 132), 171(189), 174, 175, 177 Mangiantini, M. T., 205(168, 169), 214 Mann, M., 252(107), 280 Manne, V., 364(54, 56). 378 Manning, D. R.,328(23), 332, 376( 138), 380, 404(120), 405(130), 416 Manning, R., 126(28), 144 Manos, P. N., 109(47), 120 Mansour, T., 5(8), 27(8), 41 Mansour, T. E., 11(36), 27(99), 31(144), 35(144), 42, 43, 44 Marchiori, F., 25(80, 81), 26(81), 43 Marchmont, R. J., 165(133, 134, 135), 175, 302(189), 303(189), 315 Marco, R., 55(52), 73 Marcus, F., 29(137), 32(159), 33(167, 170), 34(178), 35(159, 188), 40(137), 41(159), 44. 45, 46 Margolis, S., 195(101), 212 Mariano, T. M., 376(136), 380 Marie, J., 48(4, 8). 49(4), 50(26), 56(58), 68(4, 112), 69(114, 116), 72. 73, 74 Markey, K. A . , 251(99), 280 Marme, D., 443(118), 454

483 Maroney, P. A , , 375(122), 380 Marsch-Moreno, M., 376(134), 380 Marshall, S. E., 89(73), 90(73, 7 9 , 91(73), 94,95 Marston, S. A., 391(57), 392(57), 393(57), 414 Marston, S. B., 397(97a), 415 Martensen, T. M., 11(36), 42 Martenson, R. E., 307(221), 316 Martin, A. R., 285(2), 286(2), 311 Martin, B. R., 85(41), 94 Martin, D. B., 130(88), 131(88), 132(88), 145 Martin, L., 192(84), 193(84), 212 Martin, M. W., 294(88), 313 Marty, A . , 340(24, 27), 355 Martynyuk, A. E., 287(26), 289(26), 290(26), 303(26), 311, 340(26, 30), 355 Maruta, H., 406(140, 146), 416, 417. 443(118, 119, 120, 121). 454 Maryanoff, B. E., 34(175, 176). 36(176), 45 Mason, K., 219(21), 234(21), 238(21), 278 Massaras, C. V., 69(119), 74 Massoglia, S. L., 190(65), 192(65), 193(65), 212 Massey, T. H., 27(86, 87), 43 Mastropaolo, W., 367(69), 379 Masui, H., 169(163), 176 Matarese, G. P., 375(127), 380 Mathews, M. B., 373(98), 376(135, 136), 379, 380 Matrisian, L. M., 330(62), 333 Matsui, K., 296(111), 314 Matsumoto, K., 431(48, 49), 432(48, 49, 50, 51), 433(54), 435(67), 452, 453 Matsuura, S., 272(175), 282, 408(170), 417 Mattenson, D. R., 337(132), 357 Matthews, H. R . , 440(101), 441(102, 106), 442(109), 454 Matts, R. L., 362(40), 363(40), 364(40, 58, 59, 62), 366(62, 68), 367(58, 59), 368(58, 59, 62, 68), 369(59), 374(68), 378, 379 Matus, A . , 306(216), 316 Mauser, L., 362(39), 363(39), 364(39, 55, 56). 378 Maxon, M. J., 431(47), 452 Maxwell, C. R., 362(8), 377 May, J. M., 25(83), 43, 91(83), 95 May, M. E., 109(47), 120 May, W. S . , 331(85), 333

484 Mayer, R. J., 126(28, 32), 134(104), 135(104), 144, 145 Mayer, S. E., 152(41), 153(48), 162(80), 173, 174 Muon, M. J., 36(195), 46, 434(59, 60), 453 Mazzei, G. J., 307(224), 316 Means, A. R., 401(111, Illa), 416 Mechi, N., 375(131), 380 Medynski, D. C . , 377(146), 380 Meek, D. W., 33, 45 Meggio, F . , 25(80, 81), 26(80), 43 Mehlman, M. A,, 125(20), 126(20), 129(20), 130(20), I44 Mehra, L., 375(124), 380 Mehta, H. B., 362(42), 363(42), 364(60), 366(60), 378 Meisenhelder, J., 330(66), 333 Meisheri, K. D., 389(43), 390(43), 414 Meister, E., 99(4), 119 Melcer, I., 271(170), 281 Meligeni, J. A,, 255(132), 256(132), 280, 444(128), 445(128), 454 Melloni, E., 32(149), 33(168), 45, 209(199), 215 Mendelewski, J., 362(27), 369(76), 378, 379 Mendicino, J., 29(122), 44 Meng, H. C . , 152(37), 162(37), 163(37), 173 Menon, A. J., 194(97),212 Mercer, J. E. B., 247(83), 279 Meredith, M. J., 127(45), 128(45), 129(63), 130(63), 132(63), 134(105), 144, 145 Merkel, L., 389(43), 390(43), 414 Merlie, J.-P., 349(100), 357 Merlin, G., 375(128), 380 Memck, W. C . , 362(34, 3 3 , 376(135), 378, 380 Merrifield, R. B., 25(77), 43 Merryfield, M. L., 86(46), 92(46), 93(46), 94, 104(30), 107(30), 108(30), 109(30), I20 Messner, D. J., 351(118), 357 Metzger, H., 331(82), 333 Mewes, R., 341(35), 355 Michaelis, W., 185(41), 211 Michalek, A., 397(94), 398(94), 415 Michel, T., 324(16), 332 Michell, R. H., 293(85, 86). 313 Michelson, A. M., 371(82), 379 Michetti, F., 33(168), 45 Michetti, M., 209(199, 200), 215 Michler, A., 349(103), 357 Middleton, B., 192(88), 193(94), 212

AUTHOR INDEX

Middleton, P.,350(145), 358 Milani, A,, 169(166), 176 Mildner, P., 207(185), 214 Mildvan, A. S . , 124(15), 144 Miles, K., 300(160), 304(160), 315, 329(49), 332, 347(84), 349(84), 350(84), 351(84), 356 Milfay, D., 302(193), 303(193), 316. 329(42, 45), 332, 348(88, 89), 356 Miller, A. H., 370(78), 371(78), 378 Miller, B. F., 170(181), 177 Miller, E. A., 153(48), I73 Miller, J., 328(25), 332 Miller, J. A., 351(112), 357 Miller, J. R., 403(115), 416 Miller, 0. N., 244(80), 279 Miller, P., 306(217), 316 Miller, P. E., 293(75, 76, 77), 302(178), 307(178), 313, 315 Miller, R. E., 153(48), I73 Miller, R. H., 98(1), 99(1), 116(61), 119, 121 Miller, R. J., (7), 354 Miller, S . J., 183(35), 185(35), 206(35), 207(35, 192), 208(192), 211, 215 Miller, T. B., 91(85), 95 Miller, T. B., Jr., 228(53), 279 Mills, I . , 163(112), I75 Milstien, S . , 222(33), 223(33), 225(33, 43). 227(33), 236(33), 238(33), 239(33), 243(74), 272(33), 278, 279 Minakuchi, R., 298(138, 144), 303(138), 314 Minasian, E., 25(79), 26(79), 43 Minick, C. R., 170(182), 177 Mitchell, J. L. A., 442(113), 454 Mitropoulos, K. A., 187(49, 50), 188(50), 189(50), 194(50), 211 Mitschelen, J. J., 180(1), 188(1), 192(1), 195(106), 198(106), 199(106), 203(1), 210, 213 Mitsui, K., 363(47), 378 Mitsunaga, M., 168(160), 169(160), 176 Miyamoto, E., 296(111), 299(150, 151), 300(169), 303(169), 306(169), 307(169, 223), 309(251), 314, 315, 316, 317 Miyanaga, O., 48(19, 20), 72 Miyatake, A,, 29(134), 40(134), 44 Miyzaki, K., 299(151), 315 Mobley, P., 302(176), 315 Moir, A. J. G., 405(129, 133), 416 Mojena, M., 63, 74 Molinaro, M., 351(148), 358

485

AUTHOR INDEX Molish, I. R., 408(176), 409(176), 417 Monaghan, R., 191(74), 212 Moncada, V., 330(77), 333 Montal, M., 347(81), 349(81), 356 Montgomery, K., 405(128), 416 Mooers, S. U., 396(80), 397(80, 94), 398(94), 415 Mooney, R. A., 162(77), 166(77), 167(77), 174 Moore, A. C., 351(110), 357 Moore, R. L., 404(123, 124), 405(127), 416 Mooseker, M. S., 407(165), 417 Morad, M., 338(13), 354 Moreno, S., 436(79, 80, 81), 437(79, 82, 86, 87, 88),453 Morgan, M., 404(117), 416 Morgan, R. A , , 53(44), 54(44), 65(44), 72 Morgenroth, V. H., 252(118), 263(118), 266(1 18), 280 Morgenroth, V. H., III, 253(123), 267(158), 268(158), 269(158), 280, 281 Mori, T., 257(137), 265(137), 281, 298(137), 314 Moriarity, D., 130(88), 131(88), 132(88),145 Morikofer-Zwez, S., 34(181), 45 Morimoto, K., 397(96), 415 Morimura, H., 48(16), 72 Morin, R., 170(184), 177 Morris, P . A., 408(178), 417 Morrison, M., 70(126), 74 Morrow, C. D., 376(137), 380 Morrow, C. J., 187(52), 192(52), 202(52), 211 Mortimore, G. E., 208(193, 195), 215, 223(35), 234(35), 235(35), 238(35), 239(35), 244(35), 245(35), 246(35), 278 Moser, A. H.,198(133),213 Moser, S. H., 205(165), 214 Moskowitz, N., 299(153), 301(153), 315 Moss, J., 124(1), 126(1), 127(1, 39), 128(1), 143, 144, 165(130), 175, 377(147), 380 M r a s , S., 397(90), 409(90), 415 Mrsa, V., 207(185), 214 Muchmore, D. B., 166(155), 176 Miiller, D., 36(194), 46, 207(180), 214 Mueller, R. D., 441(104), 454 Miiller, T. H., 265(155), 281 Miiller, W. A., 244(77), 279 Muhhad, A,, 408(168), 411(191), 417, 418 Muir, L. W., 431(46), 452 Mukherjee, S. P . , 91(82), 95

Mukouyama, E. B., 366(67), 378 Muller, U., 446(143), 445 Munday, M. R., 130(84, 86, 96), 131(84, 86), 132(84), 139(120), 140(120), 145, 146 Muniyappa, K., 29(122), 44 Munk, P., 80(8, 15), 93 Murachi, T., 207(173, 175), 208(189), 209(175), 214, 215 Murad, F., 92(85), 95, 162(82), 163(82), 165(139), 174, 176 Murakami, N., 408(170), 417 Murofushi, H., 449(162, 163), 455 Murphy, R. A., 396(79, 81, 84), 397(79, 81, 90, 91), 398(81, 84), 409(90), 415 Murray, A. W., 330(58), 332 Murray, K., 7(31), 8(31), 15(31, 50). 18(53), 20(53), 21(50, 53), 22(31), 24(31), 25(31), 42 Murray, N., 307(225, 226), 316 Murray, T., 165(128), 175 Musacchio, J. M., 250(95), 251(103), 280 Mushynski, W. E., 299(155), 303(155), 315 Myant, N. B., 180(5), 210

N Nachmias, V. T., 406(143), 408(175), 416, 417

Nada, T., 36(190), 46 Nag, S., 388(34), 414 Nagano, M., 66(99), 67(99), 74 Nagata, K., 408(171), 417 Nagatsu, J., 252(114), 280 Nagatsu, T., 249(91), 252(91), 272(175), 279, 282 Naghshineh, S., 169(169), 176 Naim, A. C., 265(155), 281, 287(18, 19, 21, 22), 288(18, 19, 21, 22), 289(33), 290(18, 19, 21, 22, 33, 35, 36, 37), 291(35, 36, 37), 292(37), 293(33, 37, 68), 294(35, 36, 96), 295(37, 103, 106), 296(37, 116), 297(35, 36, 116), 298(35, 36, 98, 139), 299(37), 301(96), 302(182, 183), 303(18, 19, 21, 22, 35, 36, 37, 228), 306(36, 37), 307(228, 229), 308(35, 36, 37, 96, 139). 309(36), 310(256), 311, 312, 313, 314. 315, 316, 317, 343(56, 58). 347(58), 356 Nakagawa, M., 69(125), 74 Nakai, N., 55(56), 56(56), 73 Nakamura, H., 397(95), 415 Nakamura, S., 271(169), 273(179), 275(179), 281, 282, 297(133), 314

486 Nakamura, T., 188(55),211 Nakao, T., 168(160), 169(160), 176 Nakashima, K.,69(115, 117), 70(117, 128). 74 Nakasone, H., 397(95), 415 Nakata, H., 221(26), 249(92, 93), 263(147, 148), 264(147, 148), 274(185), 275(185), 277(147, 148), 278, 279, 281, 282, 300(166), 302(166), 303(166), 315 Nakayama, H.,351(111),357 Nambi, P., 303(199), 304(199), 316, 321(5, 8), 323(9, 11), 324(16, 17), 331. 332 Namihira, G . , 80(15), 93 Naqui, D., 27(89), 43 Narabayashi, H., 6(21), 31(21), 40(21), 41(21), 42 Narahashi, T., 353(125), 357 Nath, N., 391(54), 414 Neary, J. T., 287(28), 289(28), 290(28), 303(28), 312, 342(49), 345(69), 346(49), 355, 356 Neef, H., 82(25), 93 Negrel, R., 170(173), 176 Neher, E., 340(24, 27), 355 Neher, R., 169(166), 176 Nelson, D. L., 448(157), 455 Nelson, J. A., 191(80),212 Nelson, N., 347(81), 349(81), 356 Nemenoff, R. A., 143(132), 146, 163(106), 165(146),175, 176 Ness, G . C . , 181(15, 23), 192(93), 194(99), 210, 212 Nestler, E. J., 286(6, 7, 8), 289(7, 34), 290(8, 34), 301(6, 7, 8, 175), 302(7), 303(8), 304(7, 8), 305(175, 206, 207), 306(7), 308(8), 310(8), 311, 312, 315, 316, 336(2), 354 Netzel, R., 35(187), 46 Neumann, D., 349(144),358 Newsholme, E. A., 27(107), 32(153), 44. 45 Ngai, P. K., 397(97), 415 Nicholls, D. G., 170(174), 176 Nicholls, J. G . , 285(2), 286(2), 311 Nichols, R. A,, 299(148), 315 Nicoll, G. W., 56(59), 57(59), 73 Nicoll, R. A , , 342(50, 143), 346(50, 143), 347(50), 355, 357 Nielsen, K. H., 219(17), 234(17), 236(17), 278 Nielsen, R. C., 125(20), 126(20), 129(20), 130(20), 144

AUTHOR INDEX

Nieto, A., 27(111, 114), 28(111, 114), 44 Nikawa, J., 128(52), 129(52), 144 Nilius, B., 337(130), 357 Nilsen, T. W., 375(122), 380 Nilsson Ekdahl, K.,57, 58(66), 59, 66(66), 73 Nilsson, N. O., 148(2), 149(2, 15, 18), 150(2), 151(2), 153(2), 154(2, 49), 157(2, 49, 65, 69, 70), 158(49), 159(2), 160(2, 49, 70, 72), 162(72, 78), 163(72), 166(2), 167(78), 172, 173, 174 Nilsson, S., 149(20), 150(20, 29), 151(20), 173 Nimmo, G. A., 25(70), 43, 303(232), 307(232), 309(232), 317 Nimmo, H. G . , 32(155), 33, 45, 71, 75, 156(64), 174 Nishikawa, M., 399(105, 106), 400(105, 106), 401(105, 106), 402(105, 106, 113), 409(113), 410(185, 188). 415. 416, 417, 418 Nishikori, K.,128(53), 129(53), 144 Nishimura, S., 191(82),212 Nishiura, I., 207(173), 214 Nishiyama, U., 293(71), 313 Nishizuka, A. Y . , 265(153a),281 Nishizuka, Y., 198(130),213, 242(71, 72), 257(137), 265(137, 153b), 271(169), 273(179), 275(179), 279, 281, 282, 292(84), 293(71, 84), 294(84), 297(133), 298(135, 136, 137, 138, 144, 146), 303(138), 308(84, 236), 313, 314, 317, 324(15), 331, 409(182), 417, 430(39), 431(30), 444(39), 452 Nissler, K., 29(126, 127), 44 Nixon, C. S . , 408(172), 417 Nixon, J. C . , 233(59), 279 Noda, C., 188(55), 211 Noda, T., 434(62), 453 Noguchi, T., 48(5), 49(5), 50(5), 72 Noiman, E. S . , 409(183), 417 Noland, B. J., 187(46), 200(149), 201(151), 211, 214 Noma, A., 338(14), 339(19), 354, 355 Nonomura, Y., 397(95), 415 Nordstrom, J. L., 180(1), 188(1), 192(1), 196(106), 198(106), 199(106), 203(1), 210, 213 Norman, R. I., 351(113), 357 Northup, J. K., 327(20), 332 Norton, S . J., 236(61), 279

487

AUTHOR INDEX

Novak-Hofer, I., 347(75, 76). 356 Novotny, M. J . , 425(29), 452 Nowycky, M. C., 337(134), 340(23), 355, 357 Nudel, U., 375(126), 380 Nui, W. L., 99(5), 100(5), 119 Numa, S., 124(2), 126(2), 128(2, 52, 53). 129(52, 53), 143, 144 Nunnally, M. H., 405(127), 416 Nuzino, A., 223(39), 278 Nyfeler, F., 30(6), 37(6), 40(6), 41 0

O’Brien, P. J., 328(27), 332 O’Callaghan, J. P., 294(94), 313 Ochi, R., 340(29), 355 Ochoa, S., 362(17, 18, 20, 22, 39). 363(18, 20, 39, 47), 364(39, 54, 5 5 , 56, 63), 377, 378 Ochs, R. S., 70(129), 74 O’Connell, K., 307(222), 316 Odessey, R., 100(17), 102(17), 103(23, 24), 104(17, 24), 109(17, 24, 48), 120 Oeken, H.-J., 342(139), 357 Oestreicher, A. B., 309(239, 240), 317 Ogihara, S., 389(41), 390(41, 4 3 , 396(45), 406( 144). 414. 41 7 Ogiwara, H., 128(52), 129(52), 144 Bgreid, D., 163(103),175, 223(37), 235(37), 236(37), 237(37), 238(37), 242(37), 243(37), 278 Oka, K., 249(93), 279 Okajima, F., 66(97), 74 Okayama, H.,182(33), 184(33), 197(33), 208(33), 21 1 Okuno, S., 249(92), 279, 263(147), 264(147), 277( 147). 281 Olds, J . , 287(28), 289(28), 290(28), 303(28), 312, 342(49), 346(49), 355 Olivecrona, T., 150(22), 151(22), 156(22), 173 Oliver, C. N., 19(54), 42 Oliver, R. M., 78(2), 80(2, 8, 14). 93 Olsen, G. J., 445(138), 454 Olson, C. D., 187(47),211 Olson, M. S., 89(62), 94, 109(46), 116(55, 56), 120 Olson, N. J., 401(11 la), 416 Olsson, H., 150(29), 153(47), 154(47, 54), 157(47),173, 174

Onishi, H., 386(26), 387(28), 392(62), 393(28, 64,66), 395(66, 69), 414, 415 Opas, E. E., 409(179), 417 Oplatka, A., 408(168), 417 Orci, L., 190(63), 191(63), 192(63), 194(63), 211 Ores,C., 299(153), 301(153), 315 Oschry, Y.,166(158),176 Oshima, Y.,431(48), 432(48), 452 Oskarsson, M., 85(44), 94 Osterman, J., 48(13), 68(13), 72 Osterrieder, W., 338(15, 16), 339(16), 341(15), 354, 355 Ottaway, J., 404(117), 416 Otto, A., 29(126), 44 Ouimet, C. C., 289(34), 290(34), 295(105), 297(125), 302(178), 307(178), 312, 313, 314, 315 Ovchinnikov, L. P., 377(148), 380 Ozawa, E., 297(132), 314

P Paby, P., 244(79), 279 Packman, L. C., 80(13), 93 Paetkau, V., 27(100), 43 Paetzke-Brunner, I., 91(84), 95 Pagh, K., 443(119, 120), 454 Pai, S. H., 421(4, 5), 422(6), 428(37), 444(132), 446(132), 451, 452, 454 Pain, V. M., 362(38), 363(38, 50), 364(38), 373(102, 103), 374(103, 104), 378, 379 Palen, E., 431(44), 452 Palfrey, H. C., 296(118, 119, 120), 314 Pall, M. L., 436(72, 74), 453 Palmer, J. L., 143(132),146, 165(146), 176 Palmieri, S., 362(21), 363(21, 48), 377, 378 Pang, H., 185(40),211 Panini, S. R., 191(73), 205(164, 170), 212, 214 Panniers, R., 362(41), 363(41), 364(41), 3 78 Panter, S. S., 291(46), 312 Pappone, M.-C., 376(142), 380 Parham, P., 407(157), 417 Paris, C. G . , 347(73), 356 Parish, R. W., 446(143), 455 Park, C. R., 32(150), 45. 50(28, 29), 66(98), 72. 74, 162(84, 8 5 , 86), 163(113), 165(123),174, 175 Park, D., 302(179), 315

488 Park, D. H., 258(133), 261(133), 262(133), 265(133), 280, 302(184), 303(184), 315 Parker, C.W., 331(83,84),333 Parker, D.J . , 19(55), 42 Parker, P., 154(53), 174 Parker, P. J . , lOO(12, 13, 14). 101(12, 13,

14), 102(22), 103(22), 105(14), 107(14,39, 41), 109(12, 13, 14,22), 114(41),115(41), 116(14, 41), 120 Parker, R. A . , 183(35), 185(35), 195(108, IIO), 196(108,110), 198(108,110), 199(139), 200(110, 139,155), 202(155, 156,157),203(139, 156), 204(139), 205(139), 206(35), 207(35, 192), 208(191, 192),211, 213, 214, 215 Parkes, P. S . , 428(36), 452 Parkhurst, L.J . , 364(60), 366(60), 378 Parkin, S. M., 156(62), 175 Pamiak, M., 221(24),222(24,34),223(34), 224(34,41), 225(44), 226(34, 44).227(44), 228(44), 240(44), 248(44), 262(44), 278 Parniak, M. A , , 222(30), 236(30), 278 Part, D., 444(129), 445(129),459 Parthasarathy. R.,124(16), 144 Pask, H.T., 78(5), 84(34), 93, 94, 108(42), 120 Passeron, S . , 436(78, 79,80,8I ) , 437(79, 82, 83,84,86,87,88), 439(99), 453 Passonneau, J. V., 27(106), 44, 298(134), 314 Pastori, R.,436(81), 437(82), 453 Pastori, R. L., 436(80), 453 Patchell, V. B., 386(16), 389(44), 413, 414 Pate, T., 7(27), 8(27), 13(45), 14(45), 18(45, 53), 20(53), 21(53), 42, 200(141), 201(141), 213 Pate, T. M., 6(25), 7(25), 12(25,42), 13(25,

421,15(25, 42), 17(25), 21(25, 42), 22(25), 42 Patel, H., 119(64), 121 Patel, J . , 303(200), 304(2OO), 316 Patel, T.B . , 109(46), 120 Pater, A. 219(20), 221(20), 223(20), 226(20), 234(20), 278 Paterson, E. L., 218(5), 248(5), 278 Pathett, A . , 191(74), 212 Pato, M.D., 386(18,20,21,22,27),

387(27), 398(98), 400(27), 401(20,21), 413, 414. 415 Patston, P. A . , 102(21), 103(21), 107(40,41). 108(40), 109(49), 110(21), 111(21), 112(21,

AUTHOR INDEX

49), 113(21, 49), 114(41), 115(41, 49), 116(41), 118(49,63), 119(63), 120 Patten, G. S., 31(142, 146),44, 45 Patzelt, C . , 88(52, 54), 94 Paul, H., 108(43,44),120 Paul, R. J . , 397(93),415 Pauloin, A,, 299(152), 315 Paveto, C.,436(78), 453 Pawlson, L.G . , 165(119), 175 Paxton, R.,92(99), 95. 100(16), 101(16), 102(16), 104(16), 105(36, 37), 106(16,36, 37), 107(16,36,37), 109(16,50), 113(50), 115(50), 116(36), 120, 200(147), 201(147), 214 Payne, D. M . , 31(147),45 Payne, M. E., 296(122),314, 401(110), 416 Payne, N. J . , 165(143),176 Paznokas, J. L., 436(76), 453 Pearson, R. B., 386(15), 390(15), 413 Peczon, B. D., 50(24), 54(24), 72 Pedersen, L., 244(79), 279 Pedrosa, F.0.. 32(156), 45 Pekala, P. H . , 134(105), 145 Pelech, S.,62(80), 63(80), 73 Pelech, S. L., 171(187), 177, 200(145), 201(145), 213 Pelleh, S., 26(84), 43 Peleg, I., 411(191), 418 Pelley, J. W . , 78(4), 82(26), 83(4,28), 85(4), 86(28), 88(28), 93 Peltz, G.,407(157), 417 Pelzer, D., 340(29), 342(139),355, 357 Pemrick, S . M., 404(118), 416 Pereira Da Silva, L. H . , 444(126), 454 Perharn, R.N., 80(9, 12,13). 93 Perkins, J. P., 327(20), 332 Perrie, W. T., 404(116),416 Perry, S . V . , 386(16), 389(44),404(116,117, 125). 405(129, 133), 413, 414, 416 Persechini, A., 389(40), 404(119, 126),414, 416 Pessin, J. E., 323(12), 331 Peter, H.W., 129(70), 145 Peterkofsky, A., 428(38), 452 Peters, J. R., 303(199), 304(199), 316, 323(9, I]), 324(16, 17), 331, 332 Peterson, F. J., 206(171), 214 Peterson, P. T., 362(35), 378 Petle, D., 11(34), 42 Petrack, B., 249(280), 280

AUTHOR INDEX Petrali, E. H., 300(170), 315 Petryshyn, R., 364(58), 367(58), 368(58), 369(75), 372(86, 87, 88), 373(96, 97). 375(96), 376(132), 378, 379, 380 Pettit, F. H., 78(3), 80(11), 81(17, 18), 82(22, 26, 27), 83(27, 28, 32), 84(17, 35, 37), 85(17, 37, 39), 86(28, 35, 37), 88(28, 37), 93, 94, 200(143), 201(143), 213 Pettit, R. H., 100(15), 101(15), 102(15), 104(15), 105(15), 109(15), 120 Pfeuffer, T., 376(139), 380 Pfitzer, G., 389(43), 390(43), 414 Philipson, K. D., 341(34), 355 Phillips, C. E., 104(31), 105(31), 120, 181(23), 194(99), 210, 212 Phillips, D. R., 408(177), 417 Phillips, R., 228(52), 279 Phillips, R. S . , 222(32), 235(60), 236(60), 237(60, 65), 239(60), 242(60), 243(60), 278, 279 Pierce, M. W., 143(132), 146, 165(146), 176 Pietetti, L., 126(29), 127(29), 144 Pike, L. J., 325(18), 332 Pilkis, J., J(1, 4, 5, 6, 14, 15), 6(14, 15, 22, 23, 25), 7(14, 15, 25, 29), 8(15), 9(32), 12(22,23), 12(15, 22,25,41), 13(5,6, 25), 14(4,5,6, 14,22,23,25), 15(5,6, 14, 25, SO), 17(5,6,25), 18(5), 20(32), 21(25, SO), 22(5,6, 14,22,25,29), 23(15), 24(4,5,6), 27(110, 113), 28(1,4,5,6, 23, 110, 113, 1 IS), 29(5,6), 30(6, 113), 32(160, 164), 33(160, 164), 34(176), 36(176), 37(1,4,5,6, 14.22, 29,41), 40(6), 41,42,44,45, 200(140),201(140)213 Pilkis, S . J., 5(1, 4, 5, 6, 7, 14, 15, 17), 6(7, 14, 15, 22, 23, 25), 7(7, 14, 15, 17, 25, 26, 27, 29, 31), 8(15, 27, 31), 9(32), 12(22, 23), 12(15, 22, 25, 41, 42), 13(5, 6, 17, 25, 42, 4 3 , 14(4, 5, 6, 14, 17, 22, 23, 25, 26, 42, 4 3 , 15(5, 6, 14, 17, 25, 26, 31, SO), 17(5, 6, 25), 18(5, 7, 26, 45, 53), 20(32, 53), 21(25, 42, 50, 53), 22(5, 6, 14, 17, 22, 25, 26, 29, 31, 60), 23(15), 24(4, 5, 6, 7, 26, 31, 66), 25(31), 26(84), 27(91, 104, 110, 113). 28(1, 4, 5, 6, 23), 48(3), 49(3), 53(45, 48), 54(45, SO), 55(53), 59(48, 71), 60(48), 61(48), 62(50, 71, 80, 81). 63(80), 65(45, 9 9 , 66(48, 95, loo), 67(3, loo), 71(3), 72, 73, 74, 91, 110, 113, 1151, 29(5, 6, 91), 30(6, 113), 32(150, 157,

489 160, 164), 33(160, 164, 166), 34(157, 166, 175, 176, 182, 185), 35(157, 182, 186), 36(176), 37(1, 4, 5, 6, 14, 22, 29, 41, 200). 38(200, 203), 40(6, 7, 205), 41, 42, 43, 44, 45, 46, 200(141), 201(141), 213 Pinna, L. A., 25(80, 81), 26(80), 43 Pinphanichakam, P., 362(15), 377 Pittman, R. C . , 152(39), 173 Pizzighella, S . , 349(144), 358 Plesner, P., 449(165), 455 Podesta, E., 162(90, 91), 169(90, 91), 170(90, 91), 174 Podesta, E. J., 169(166), 176 Podleski, T. R.,349(98), 356 Podskalny, J. M., 330(77), 333 Pogell, B. M., 32(151, 152, 162), 45 Pogson, C., 26(84), 43 Pogson, C. J., 48(9), 62(80), 63(80), 72, 73 Pogson, C . I., 229(55, 57), 230(55), 234(55), 241(66), 243(55), 279 Pohlig, G., 434(62, 63), 453 Polakis, S . E., 124(1), 126(1), 127(1), 128(1), 43 Pollard, T. D., 292(59), 312, 382(3), 395(75), 406(141, 145), 407(158), 408(3), 413, 415, 416, 417 Pollock, R. J., 259(144), 260(144), 261(144), 265(144), 267(144), 281 Pomerantz, A. H., 25(77), 43 Ponta, H., 421(5), 422(6, 7). 428(37), 451, 452 Pontremoli, S . , 5(11), 14(48), 32(11, 149, 154, 156, 158, 161), 33(168), 41, 42, 45, 209(199, 200), 215 Ponzio, G., 165(144), I76 Poole, G. P., 57(65), 73 Poorman, R. A,, 25(138), 30(138), 40(138), 44 Pope, T. S . , 126(31), 144 Popt, J.-L., 347(79), 356 Popp, D. A,, 91(90), 95 Portenhauser, R., 88(52), 94 Porter, J. W., 127(39a), 134(106), 144, 146, 180(11), 210, 201(152), 214 Porter, M. E., 406(141), 416 Portman, 0. W., 170(180), 177 Postle, A. D., 67(108), 74 Postma, P. W., 423(14), 452 Potter, J. D., 405(134), 416 Poulis, P., 165(140), 176

490

AUTHOR INDEX

Powell, S. M., 200(147), 201(147), 214 Powers, D. M., 422(10), 451 Powers, P. A,, 436(72), 453 Pratt, M. L., 83(30), 84(38), 94 Prendergast, F. G., 401(110a), 416 Presek, P., 70(135, 136), 71(136, 137), 75 Press, J. C . , 390(51), 414 Prigge, W. F.,206(171), 214 Pritchwd, K.,397(97a), 415 Pugh, E. N., 328(33, 34), 332 Pullinger, C. R., 200(204), 215 Punch, D. L., 310(254), 317 Puszkin, S., 299(153), 301(153), 315

Q Quarum, M., 300(159), 315 Qureshi, A. A., 134(106), 146

R Rabadjiija, M., 252( 116), 280 Raben, M. S . , 149(12), 152(12), 173 Rabinovitz, M., 362(8), 377 Racker, E., 347(80), 349(80), 356 Rackoff, W. R., 330(70), 333 Rae, J. D., 25(79), 43 Raese, J. D., 265(154), 270(163), 271(166), 281 Raese, J. E., 257(141), 258(142), 281 Raftery, M. A., 329(41), 332, 347(78), 348(94), 351(110, I l l ) , 356, 357 Ragnarsson, U., 25(76, 82), 35(187), 43, 46, 51(41), 61(74), 62(74), 64(90, 91), 72, 73, 74, 200(142), 201(142), 213, 223(38), 278 Rahmsdorf, H. J., 421(4, 5 ) , 422(6, 7), 428(37), 443(115, 116), 444(132), 446(132), 451, 452, 454 Rainbow, T. C., 302(180), 315 Rall, T., 253(119), 280 Rall, T. W., 226(49), 278 Ralston, R., 362(16), 377 Ralston, R. 0..362(21, 23), 363(21, 48, 49), 377, 378 Ramaiah, A,, 5(9), 11(35), 27(9, 96), 41, 42, 43 Ramasanna, T., 194(97),212 Ramos, B. V . , 185(44),211 Ramsey, A. J., 130(86), 131(86), 145 Randall, D. D., 78(4), 82(26), 83(4), 85(4), 93

Randle, C. L., 307(222), 316 Randle, P. J., 78(5, 7), 81(19), 82(20, 21, 24), 83(29), 84(34), 85(41), 88(55, 56, 57, 58, 59), 89(20, 59, 60,61, 72). 92(96, 98), 93, 94, 95, 99(3), 100(3, 12, 13, 14, 19), lOl(12, 13, 14, 19), 102(21, 22), 103(21, 22, 261, 104(3, 19, 28, 31), 105(14, 19, 31), 106(19), 107(14, 19, 26, 40). 108(40, 421, 109(12, 13, 14, 19, 22, 26, 28, 45, 491, 110(21,45), 111(3, 21,45), 112(21, 45, 491, 113(21, 49), 115(49), 116(14, 57), 118(49, 63), 119(63, 64),119, 120, 121, 125(17), 144 Rane, S. G., 341(135), 357 Ranganathan, S., 201(152), 214 Rangel-Aldao, R., 291(42), 312, 438(93), 448(154, 1 5 3 , 453, 455 Ranu, R. S., 362(16, 19, 24, 26). 363(19), 365(65), 366(65, 66) 367(66), 369(73), 374(109, 110, 11l), 377, 378, 379 Rao, D. N., 248(84), 279 Rapport, S., 375(128), 380 Rasmussen, H . , 337(6), 354 Ravdin, P., 349(98), 356 Ray, P. D., 67(103), 74 Raynor, R. L., 298(140, 141), 314 Reardon, I., 35(188), 46 Redman, R., 368(71), 369(71), 379 Reed, B. Y . , 228(52), 279 Reed, L. J., 78(1, 2, 3, 4), 80(2, 8, 11, 14, 15, 16), 81(17, 18), 82(22, 23, 26, 27), 83(4, 23, 27, 28, 31, 32), 84(17, 35, 37), 834, 17, 37, 39, 43), 86(28, 31, 35, 37, 461, 88(28, 37), 91(43), 92(46), 93(46), 93, 94, 100(15), 101(15), 102(15), 104(15, 30), 105(15), 107(30), 108(30), 109(15, 30), 119(65, 66), 120, 121, 200(143), 201(143), 213 Rees, D. D., 388(32), 414 Rees-Jones, R. W., 300(159), 315 Reeves, H. C . , 425(21, 23), 452 Regen, D. M., 3 5 , 6), 6(25), 7(25), 12(25), 13(5, 6, 25), 14(5, 6, 25), 15(5, 6, 25 ), 17(5, 6, 25), 18(5), 21(25), 22(5, 6, 25), 24(5, 6), 28(5, 61, 29(5, 6), 30(6), 370, 6), 40(6, 205), 41, 42, 46, 200(141), 201(141), 213 Reichwdt, L. F., 300(171), 315 Reichel, P. A., 376(135, 136). 380 Reichlin, M., 291(41), 312 Reid, J. V. D., 252(111), 280

AUTHOR INDEX Reid, K . B. M.,81(19), 93 Reimann, E. M.,152(40), 153(40), 173 Reinacher, M.,70(135), 75 Reinauer, H., 170(177), 176 Reinhart, G. D., 27(103), 43, 44 Reinhart, P. H., 88(51), 94 Reis, D. J . , 250(97), 258(133), 261(133), 262(133), 265(133), 280, 302(184), 303(184), 314 Reitz, A. B., 34(176), 36(176), 45 Reitz, D. B . , 34(175), 45 Reizer, J., 425(29), 452 Renart, M. F., 444(133), 446(133), 447(133), 454

Renaud, J.-F.,342(140), 357 Renner, R., 165(137), 175 Renson, J . , 271(172), 282 Renyon, K. R . , 328(26), 332 Resink, T. J . , 165(150), 176 Reuter, H., 287(25), 288(25), 289(25), 290(25), 303(25), 311, 337(8, 9, 1 I), 338(9), 339(17), 340(17, 20, 21, 22). 342(48), 346(48), 354, 355 Revel, M., 372(90), 375(116, 126, 128), 379, 380 Revel, M.-O., 302(187, 188), 303(187, 188), 315 Rey, F., 229(55), 240(55), 241(55), 247(82), 2 79 Reydel, P., 69(125), 74 Reynolds, J. A . , 329(40), 332 Reznikov, D. C . , 202(205), 215 Rial, E., 170(174), 176 Ribeiro, J. A . , 353(124), 357 Richards, B. A., 191(71),212 Richards, C. S., 5(3), 22(3), 28(3), 29(3), 36(198), 37(201), 41, 46 Richards, E. G . , 29(134), 40(134), 44 Richelson, E., 294(87), 313 Rickenberg, H. V . , 444(128, 134, 137), 445(128, 139, 140). 446(141), 447(147, 148), 454, 455 Ries, B., 207(185), 214 Rigmaiden, M.,408(176), 409(176), 417 Rinaldi, M. L., 341(32, 33), 355 Riou, J. P., 32(157), 34, 135(157),45, 53(45), 54(45), 55(53), 65(45, 9 9 , 66(95), 72, 73, 74 Riquelme, P. T., 12(39, 40), 29(133, 137), 34(177), 40(133, 137), 42, 44, 45 Ritchie, I. M., 275(185), 282

Ritkey, J. A., 401(111a), 416 Rittenhouse, J., 33(170), 34(178), 35, 45, 46 Rivett, A . J . , 207(186), 215 Rizack, M.A , , 149(11, 14), 152(11, 14), 162(14), 173 Roach, P. J . , 264(152), 281, 296(121), 314 Roberts, A. F. C., 130(86), 131(86), 145 Roberts, G. C . K . , 80(12, 13), 93 Roberts, M. H. T., 275(187), 282 Roberts, W. K., 372(92), 375(92), 379 Robertson, H. D., 375(115), 380 Robertson, S. P., 405(134), 416 Robinson, D., 275(186), 282 Robinson, D. R., 156(62), 174 Robinson, D. S . , 272(174), 275(174), 272, 282 Robinson, E. A . , 407(154), 410(186), 411(186), 417, 418 Robinson, F. W., 165(122, 127), 175 Robinson, G. A . , 162(79), 174 Robishaw, J. D., 377(143), 380 Robson, N. A . , 130(85), 131(85), 145 Rocha, D. M . , 244(78), 245(78), 279 Roche, T. E., 78(4), 81(17, 18). 82(23), 83(4, 23, 30, 31). 84(17, 33, 36, 38). 85(4, 17), 86(31), 93, 94 Rodbell, M.,157(67), 174 Rodvien, R . , 273(94), 379 Rodwell, V. W., 180(1, 9), 188(1), 192(1), 195(106), 196(9), 197(115, 120, 123), 198(106, 125, 126, 127), 199(106, 120), 200(9, 203), 203(1), 210, 213, 215 Roehrig, K. L., 132(97), 145 Rosen, P., 170(177),176 Rogerge, C., 260(146), 268(146), 281 Rogers, D. H., 197(115), 208(187), 213, 215 Rognstad, R., 71, 75 Rogol, A. D., 166(156), 176 Roitelman, J., 194(98), 212 Romea, G., 375(131), 380 Romey, G., 337(133), 357 Ronft, H., 80(10), 93 Rosa, F., 373(96), 375(96), 379 Rosberg, S., 157(71), 158(71), 161(71), 166(71, 154), 174. 176 Rose, I. A , , 69(121), 74 Roselle, M.,431(41, 42), 452 Roseman, S., 423(14), 452 Rosen, H., 361(6), 377 Rosen, 0. M., 150(28), 151(28), 156(28), 157(28), 162(98), 170(28), 173, 175,

492 287(15), 291(41), 292(63), 293(71a), 300(15), 303(63, 71a), 311, 312, 313, 420(1), 422(12), 451, 452 Rosenbaum, J. L., 306(218), 316 Rosenfeld, A., 388(33), 390(33), 414 Rosenfeld, S. S., 391(58, 59), 392(58, 59), 414 Roskoski, R., Jr., 255(131), 260(131), 270(165), 271(165), 280, 281, 291(44, 4 3 , 312. 421(5), 451 Rosman, J., 373(95), 379 Rosner, M. R., 330(64, 68). 333 Ross, A. H.,330(64), 333 Ross, E. M., 292(61), 312 Ross, R., 191(77), 192(77), 193(77), 212 Rosselin, G., 229(54), 240(54), 241(54), 279 Rossi, B., 165(144), 176 Rossi, G. B., 375(127, 131), 380 Rossie, S., 353(128), 357 Roth, J., 300(159), 315, 330(75, 76, 77). 333 Roth, R. H., 252(109, 112, 118), 253(123), 260( 145). 263( 1 18), 266( 118), 267( 158), 268(158, 159), 269(158, 159, 161), 275(189), 280, 281, 282 Rothlein, J., 305(209), 316 Rothlein, J. E., 296(118), 314 Rothman, J. E., 187(48), 211 Rothrock, J., 191(74), 212 Rougier, O., 337(10), 354 Roy, R., 362(21, 23), 363(21, 48), 377, 378 Rozengurt, E., 50(23), 72, 330(61), 333 Rubin, C. S., 162(98), 175, 291(42, 43, 53, 292(63), 303(63), 312 Rubin, L. L., 350(108), 357 Rubin, R. A., 330(70), 333 Rudney, H., 181(13), 191(73), 193(13), 195(105), 197(115), 205(164, 170), 208(187), 210, 212, 213, 214, 215 Rudolph, S. A., 306(218), 316 Rubsamen, H., 70(136), 71(136), 75 Ruoho, A. E., 323(12), 321 Russel, T. R., 165(128), 175 Russell, D. W., 182(33), 184(33), 190(64), 192(87), 197(33), 208(33), 211, 212 Russo, M. A,, 401(111a), 416 Ruth, P., 341(35), 355 Rutherford, C. L., 444(130), 447(149), 454, 455 Rutter, W. J., 432(52), 452 Ryan, J., 182(29), 191(29), 211 Ryder, E., 127(40), 144 Rylatt, D. B., 25(70), 43

AUTHOR INDEX S

Sabine, J. R., 180(8), 189(62), 194(62), 210, 211 Sabir, M. A., 90(76), 95 Sabularse, D. C., 29(128), 44 Safer, B., 360(4), 362(35, 44), 363(44, 5 1, 52, 53), 364(44, 51), 366(44), 377, 378 Sagara, J., 408(171), 417 Sager, G., 69(124), 74 Saggerson, E. D., 128(58), 129(58), 144 Saheki, K., 48(6, 7), 72 Saheki, S., 48(6, 7), 68(110), 72, 74 Sahyoun, N. E., 330(81), 333 Saier, M. H., Jr., 423(13, 15). 425(28, 29), 426(15, 28), 452 Saitoh, T., 329(44), 332, 348(90, 91), 350(91), 356 Sakai, K., 297(133), 314 Sakai, T., 165:121), 175 Sakakibara, R., 5(16), 6(16), 7(16), 12(16, 43). 13(43), 14(43), 15(16, 43), 20(58), 21(16, 59), 22(16), 23(43), 29(130), 30(140), 41, 42, 43, 44 Sakmann, B., 340(24), 349(103), 355, 357 Sala, G. B., 162(89), 169(89, 163, 170(89), 174, 176 Salamino, F., 33(168), 45, 209(199, 200), 215 Salans, L. B., 143(133), 146 Salas, M. L., 46 Sala Tkpat, J., 48(8), 72 Sale, G. J., 89(60, 61), 94 Salers, P., 166(159), 176 Salimans, M., 364(57), 378 Salomon, D. S., 330(59), 333 Salomosson, I., 257( 140), 281, 294(97), 298(97), 313 Saltiel, A., 91(89), 95 Saltiel, A. R., 91(93), 95, 330(81), 333 Salzman, P. M., 267(158), 268(158), 269(158), 281 Sampson, J., 374(105), 379 Samuel, C. E., 375(123), 380 Sanchez, J. A., 342(141), 357 Sanders, C., 338(13), 354 Sanghvi, A., 180(12), 201(12), 209(12), 210 Sanno, Y.,68(110), 74 Sano, K., 242(71, 72), 279, 298(146), 314 Santana, M. A., 241(66), 279 Sarkar, D., 291(42, 43,55), 312 Sarner, N., 440(101), 454 Sarngadharan, M. G., 32(162), 45

493

AUTHOR INDEX Sarver, J. A,, 165(122), 175 Sasaki, T., 32(154), 45, 208(189), 215 Sasaki, Y . , 411(189), 418 Sastre, L., 444(133), 446(133), 447(133), 454 Satir, B. H., 449(161), 455 Sauviat, M. P., 337(10),354 Sawyer, D. F., 321(8), 331 Sawyer, S. T . , 330(69), 333 Scallen, T. J., 180(12), 187(46), 200(149, 150), 201(151), 203(158), 209(12), 210, 211, 214 Schaeffler, G. E., 247(81), 279 Schaller, K. L., 444(137), 445(139, 140), 446(141),454, 455 Schatzman, R. C., 298(141), 314 Schecket, G., 299(154), 303(154), 315 Schecter, I., 181(14), 210 Scheid, C. R., 90(77), 95 Schell, M. A., 347(80), 349(80), 356 Schellenberger, A., 82(25), 93 Schellenberger, W., 29(126, 127), 44 Schepens, J. T. G., 109(51), 115(51, 54). 120 Schiebler, W., 305(203, 209), 316 Schiltz, E., 56(59), 57(59), 73 Schimizu, H., 253(120), 280 Schimke, R. T., 375(129), 380 Schimmel, R. J., 90(77), 95, 163(108), 175 Schirmann, A., 88(52), 94 Schlatter, S., 30(141), 41(141), 44 Schlessinger, J., 349(98), 356 Schlichter, D. J., 302(190), 303(190),315 Schlumpf, J. R., 22(60), 27(110, 113), 28(110, 113, 115), 30(113), 34(182), 43, 44, 45 Schmid, A., 342(140), 351(113),357 Schmidlin, S., 446(143), 445 Schmidt, A., 375(116, 128), 380 Schmidt, J., 353(128), 357 Schmidt, M. R., 423(15), 426(15), 452 Schmidt, R. A., 192(86), 212 Schmitt, W., 56(59), 57(59), 73 Schneider, C. J., 192(86),212 Schneider, M., 206(172), 214 Schoen, C., 444(131), 445(131), 454 Scholey, J. M., 384(6), 387(29), 393(29), 395(70), 407( 163), 413, 414, 415, 443(123), 454 Scholz, H.,339(17), 340(17), 355 Schonberg, G., 191(74), 212 Schoner, W . , 48(15), 50(15), 70(131, 132, 133, 134, 135, 136), 71(136), 72, 75 Schook, W., 299(153), 301(153), 315

Schotland, D. L., 351(116), 357 Schotman, P.,309(237, 240, 241), 317 Schrag, K. J., 11(33), 42 Schroepfer, G. J., Jr., 180(7), 187(7), 190(7), 210 Schubert, D., 353(127), 357 Schuetze, S. M., 349(104), 350(108, 149, 357, 358 Schulman, H., 291(47), 294(91, 92), 296(114), 301(91), 312, 313, 314 Schulman, H. M., 362(7), 377 Schulman, L., 372(90), 375(128), 379, 380 Schultz, J. E., 448(158), 449(159, 160), 455 Schulze-Wethmar, F. H., 88(53), 94 Schuman, H.J., 252(106), 280 Schwartz, J., 347(73), 356 Schwartz, J. H., 287(18, 19), 288(18, 19), 290(18, 19), 303(18, 19). 310(258), 311, 317, 343(55, 56, 58). 347(58), 355, 356 Schwartz, R. P.,Jr., 392(61), 414 Schweiger, M., 421(4, 5 ) , 422(6, 7), 428(37), 451, 452 Schwoch, G., 442(107), 446(107), 454 Schworer, C., 38(203), 46 Schworer, C. M., 208(195), 215, 223(35), 234(35), 235(35), 238(35), 239(35), 242(69, 70), 244(35), 245(35), 246(35), 264(151), 278, 279. 281, 296(122), 314 Scolnick, E. M., 321(7), 331 Scow, R. O . , 155(58), 174 Scrutton, M. C . , 48(1), 5(1), 72 Scubert, W., 48(15), 50(15), 72 Seals, J. R., 91(87, 88, 91, 94), 95 Sebastian, J., 444(133), 446(133), 447(133), 454

Sebastiao, A. M., 353(124), 357 Seckler, B., 449(159), 455 Sedvall, G. C., 252(113), 280 Seebeck, T., 442(11l), 454 Seeburg, P. H., 377(146), 380 Sefton, B. M., 299(157), 303(157), 315. 350(109), 357 Seglen, P. O., 208(194), 215 Seidel, J. C . , 388(34), 391(54), 395(71), 414, 415

Seigelbaum, S. A., 343(59), 344(59), 356 Seigelchifer, M. A., 437(83, 84), 453 Seitz, H. J., 58(68), 73 Sekura, R. D., 376(141), 380 Selden, S. C . , 292(59), 312 Sellers, J. R., 384(9, Il), 386(22, 27), 387(27), 388(9), 389(39, 42), 390(42),

AUTHOR INDEX 391(52, 56, MI), 392(56), 393(56, 63), 399(102), 400(27, 102), 404(56), 406(11, 135). 410(185, 186), 411(186), 413, 414, 416, 417, 418 Sen, G. C., 372(91), 375(91), 379 Sena, G. R., 203(158), 214 Sener, A., 29(119), 44 Sensheimer, H.-P., 293(73), 303(73), 313 Serrero-Dave, G., 170(173), 176 Severson, D. L., 78(5), 89(72), 93, 94, 152(45, 46), 170(175, 176), 173, 176 Severson, D. M . , 108(42), I20 Sewell, E. T . , 100(18), 102(18), 120 Sexton, R. C., 191(73), 205(164), 212, 214 Shachter, E., 162(96, 97). 175 Shaila, S., 372(91), 375(91), 379 Shapiro, B., 166(158), 176 Shapiro, L., 422(8, 12), 451, 452 Shaw, J. M., 118(63), 119(63), 121 Shechter, I., 194(98), 212 Sheetz, M. P., 399(100, 101, 102), 400(102), 407(150, 157), 415, 416, 417 Shenkman, L., 251(99), 280 Shenolikar, S., 25(73), 43, 62(81), 73 Shepher, G. M., 285(1, 3), 286(3), 311 Sheppard, J. R.,448(156), 455 Sheppy, F.,249(94), 280 Sherr, H. P., 68(111), 74 Sherry, J. M . F.,386(25), 388(25), 414 Sheu, K.-F.R.,309(250), 317 Shichi, H . , 300(161), 303(161), 315, 328(27, 29, 37), 332 Shields, P. J., 275(188), 282 Shih, T. Y.,321(7), 331 Shima, S., 168(160), 169(160), 176 Shiman, R.,219(14, 20). 221(20, 27), 223(20, 35), 226(20), 234(20, 35), 236(27, 35, 63), 238(35), 239(35), 243(27), 244(35), 245(35), 246(35), 249(14), 273( 14), 278 Shimizu, K., 377(144), 380 Shinitsky, M., 194(95), 212 Shirakawa, S., 399(106), 400(106), 401(106), 402(106), 415 Shirron, C., 253(121), 280 Shneider, W. J . , 190(64), 211 Shoji, M., 257(140), 281, 294(97), 298(97), 313 Shorr, R. G. L., 321(8), 331 Shoukimas, J. J., 345(68), 356 Shoyab, M., 330(60), 333

Shrago, E., 449(164), 455 Shrewsbury, M. A . , 200(148), 214 Shuman, H., 389(38), 414 Shuster, M. J . , 342(44), 344(61), 347(61), 355, 356 Sibley, D. R . , 303(199), 304(199), 316, 321(4), 323(9, l l ) , 324(16, 17). 327(21), 331, 332 Siegel, M., 91(89), 95 Siegel, M. I., 91(93), 95 Siegelbaum, S. A., 287(20), 288(20), 290(20), 303(20), 311, 342(43, 44). 344(43, 61), 347(61), 355, 356 Sieghart, W . , 291(48), 294(98), 312, 313 Siegman, M . J.;396(80), 397(80, 94). 398(94), 415 Siegmann, M., 438(95), 453 Siekevitz, P., 301(174), 315 Siekierka, J., 362(39), 363(39, 47), 364(39, 54, 55, 56), 376(135, 136), 378, 380 Siess, E. A , , 85(40), 88(52, 53), 94, 129(64), 145 Sewers, D. J., 11(33), 42 Sigworth, F. J . , 340(24), 355 Silberkang, M., 185(43), 211 Silberman, S. R., 84(37), 85(37), 86(37), 88(37), 94 Silver, P. J., 396(82), 397(82), 403(115), 415, 416 Silverman, P. M . , 438(90), 453 Simon, J. R., 260(145), 268(159), 269(159), 281 Simon, M. I., 377(143), 380 Simon, M.-P., 48(4, 8), 49(4), 56(58), 68(4), 69(4), 72, 73 Simoni, R. D., 181(16, 24), 182(16, 24, 29, 31), 183(24), 191(24, 29), 210, 211 Simonnet, G., 276(194), 277(194), 282 Simons, K., 150(23), 173 Simons, S., 447(149b), 455 Simpson, I., 143(134), 146 Simpson, I. A , , 143(133), 146 Sinensky, M., 190(67), 212 Singer, I. I., 192(93), 212 Sipat, A. B . , 189(62), 194(62), 211 Siperstein, M., 205( 169), 214 Siperstein, M. D., 191(76, 79, 80), 198(133), 205(165, 166), 212, 213, 214 Sitaramayya, A., 328(35, 36), 332 Sitges, M., 197(116, 118, 122, 124), 199(122, 134), 213

AUTHOR INDEX Sitikov, A. S . , 377(148), 380 Sjoerdsma, A., 252(111, 115), 271(168), 272(173, 174, 176), 273(176), 275(173, 174, 176, 186). 280, 281, 282 Skotland, T., 221(25), 278 Slaughter, C. J., 190(64),211 Slayter, H. F., 395(71), 415 Sloan, S. K., 152(46), 173 Sloboda, R. D., 306(218), 316 Small, J. V., 386(23), 414 Smigel, M. D., 292(61),312 Smillie, L. B., 404(116), 405(131), 416 Smilowitz, H., 329(47), 332, 348(92), 356 Smith, C. W . J., 397(97a), 415 Smith, D., 377(146), 380 Smith, E. L., 19(57), 42 Smith, M., 194(99), 212 Smith, M. M., 165(127), 175 Smith, R., 393(67), 415 Smith, R. C., 395(70), 415 Smith, R. M., 162(75), 174 Smith, S., 128(49), 144 Smith, S. C., 247(83), 279 Smoluk, G. D., 299(155), 303(155), 315 Smyth, J. E., 141(124), 143(124),146 Snee, J., 151(35), 170(35), 173 Sneyd, J. G . T., 163(113), 165(117), 175 Snider, R. M., 294(87), 313 Snodgrass, P. J., 192(83), 193(83),212 Snyder, S. H., 290(31), 299(31), 312, 342(142), 346(142), 357 Sobel, A., 302(194), 303(194), 310(194), 316, 329(43), 332, 347(79), 348(86), 356 Sobieszek, A., 386(23), 390(45a), 396(75a), 414, 415 Sobue, K., 296(112), 314, 397(96), 415 Soderling, T. R., 7(31), 8(31), 15(31), 22(31), 24(31), 25(31), 38(203),42, 46, 162(84), 174, 242(69, 70), 264(151), 279, 281, 296(122), 314 Sogin, M. L., 445(138), 448(150), 454, 455 Solano, A. R . , 162(91), 169(91), 170(91), 174 Solaro, R. J., 405(133, 134). 416 Sold, G., 293(69), 313 Soling, H.-D., 27(90), 28(118), 31(145), 43, 44, 45

Sols, A., 29(124),44, 46, 55(52), 73 Sols, D. L., 33(172), 45 Somers, R. L., 300(161), 303(161), 315, 328(27, 29, 37), 332 Somlyo, A. P., 389(38), 395(73), 414, 415

495 Somlyo, A. V., 389(38), 395(73), 414, 415 Sommerschild, H., 349(101), 357 Sonenberg, M., 69(123), 70(127), 74 Song, C. S . , 126(24), 144 Sonne, 0..143(134),146 Sonnenborn, U., 169(170), 171(170), 176 Sonnhof, V., 302(181), 315 Sorensen-Ziganke, B., 29(135), 44 Sorenson, R. G., 309(238), 317 Souroujon, M., 349(144), 358 Spaeth, A. E., 130(72, 73), 145 Spagnuolo, S . , 205(168, 169). 214 Sparatore, B., 33(168), 145, 209(199, 200), 215 Speake, B. K., 156(62), 174 Spector, S., 252(115), 280 Spencer, T. A., 191(80),212 Sperman, T. N., 422(1I), 428(1 l), 452 Speny, P. J., 293(69), 313 Spiess, J., 425(23), 452 Spitz, E., 444(128, 134), 445(128, 139, 140), 454 Spivey, H. O., 50(24), 54(24), 72 Sprengers, E. D., 68(112, 113), 74 Springer, J., 191(74),212 Springer, W. R., 446(144), 455 Spudich, E. N., 426(31), 452 Spudich, J. A., 399(100, 101, 102). 400(102), 406(142), 407(142, 157), 415, 416, 417, 443(117), 454 Spudich, J. L., 426(30a, 31), 452 Spurgeon, S . L., 180(11), 210 Staal, G. E. J., 68(113), 74 Stacpoole, P. W., 206(172), 214 Stadel, J. M., 320(1), 321(5, 8), 324(16), 327(19), 331, 332 Stadler, J., 444(125), 454 Stadtman, E. R., 19(54), 42, 87(47), 94, 162(93, 96, 97), 163(107), 175 Staehlin, T., 362(36), 378 Stafford, W. F., 391(55), 395(71), 406(141), 414, 416 Stahl, Y. D., 445(138), 454 Stahlman, M., 328(25), 332 Stallings, W., 124(16), 144 Stam, H., 170(178, 179), 176 Staniszewski, C., 325(18), 332 Stansbie, D., 78(5), 93, 130(74), 131(74), 132(74), 145 Stapley, E., 191(74),212 Stark, G. R., 375(129), 380

496 Starling, J. A., 31(147), 45 Steck, A. J., 307(225, 226), 316 Steffen, H., 169(166), 176 Stein, J. C . , 291(43), 312 Stein, L. A., 392(61), 414 Steinback, K. E., 427(33), 452 Steinberg, D., 148(3), 149(3, lo), 150(26, 27), 151(31, 32), 152(39, 41, 42, 43, 45), 153(48), 155(42, 55, 56), 156(3, 43), 157(68), 170(31, 185), 171(186), 172. 173, 174, 177 Steinberg, F., 376(141), 380 Steiner, A. L., 449(160), 455 Steiner, D. E., 35(186), 46 Steiner, K. E., 66(100), 67(100), 74 Stephenson, F. A., 166(156), 176 Stepp, L. R., 82(27), 83(27), 93 Sternberger, L. A., 299(156), 315 Sternberger, N. H.,299(156), 315 Stevens, C. F., 340(20), 355 Stewart, A. A,, 62(82), 63(82), 73, 154(51), 174, 309(248), 310(255), 317, 336(4), 354 Stewart, B. H.,5(5), 13(5), 14(5), 15(5), 17(5), W ) , 22(5), W 9 , 28(5), 29(5), 37(5), 41, 200(141), 201(141), 213 Stewart, C. S . , 202(156), 203(156), 214 Stewart, G., 408(176), 409(176), 417 Stewart, H. B., 5(6), 6(25), 7(25, 26), 12(25), 13(6, 25), 14(6, 25, 26), 15(6, 25, 26), 17(6, 25), 18(26), 21(25), 22(6, 25, 26). 24(6, 26, 66), 28(6), 29(6), 30(6), 37(6), 40(6), 42, 43 Stewart, W. E . , 11, 375(117), 380 Stidwell, R. T., 407(166), 417 Stifel, F. B., 27(108, 109), 34(108, 109), 44, 50(31), 65(31), 68(109, 111). 72, 74 Stiffens, J. J., 11(33), 42 Stiles, G. L., 320(2), 331 Still, J., 195(103),212 Stitt, M., 6(18, 19), 18(18), 41 Stjame, L., 252(109, 112), 280 Stock, J. B., 162(95), 175 Stoeckenius, W., 426(30a), 452 Stoecklin, F. B., 34(181), 45 Stokstand, E. L., 218(5), 236(5), 278 Stone, S . R., 18(51, 52), 42 Stone, T. W., 293(80), 313 Stonik, I. A., 196(111, 112). 197(114, 117). 198(111, 128, 129), 202(154), 203(117), 209(213), 213, 214 Stoops, J. K., 124(6), 143

AUTHOR INDEX Storti, R. V., 396(78), 415 Stoscheck, C., 330(53), 332 Strack, I., 244(80), 279 Strada, S . J., 163(105), 175, 294(88), 313 Strfilfors, P., 148(1, 2, 4, 5), 149(1, 2, 4, 15, 16), 150(1, 2), 151(2, 34), 152(16), 153(1, 2, 16, 47), 154(2, 16, 47, 49, 52, 54), 155(52), 156(5), 157(2, 16, 47, 49, 65, 66), 158(49, 66, 90), 159(2, 16, 66), 160(2, 5, 49, 66),162(78, IOO), 163(5), 166(2, 4), 167(78), 168(4), 169(34), 172, 173, 175, 177 Strandholm, J. J., 70(130), 75 Strasser, R. H.,327(21), 332 Stratmen, F. W., 208(197), 215 Strickland, S . , 162(94), 175 Strittmatter, W. T., 324(13), 331 Strohsnitter, W., 90(77), 95 Strong, J. A,, 287(30), 289(30), 290(30), 299(30), 303(30), 312, 341(31), 345(67), 355, 356 Stroud, R. M., 183(36), 184(36), 185(36), 207(36), 208(36), 211 Strulovici, B., 323(10), 324(10), 327(19), 331, 332 Stmmwasser, F., 287(21), 288(21), 290(21), 303(21), 311, 342(45), 344(62), 345(63, 64, 65, 66), 346(65), 347(74), 355, 356 Stryer, L., 376(141, 142), 380 Stuart, D. K., 344(62), 356 Stubbe, J., 124(14), 143 Studier, F. W., 421(5), 451 Stuhmer, W., 353(126),357 Stull, J. T., 396(76, 82), 397(82, 92), 398(92), 403(76, 115), 404(119, 120, 123, 124, 126), 405(127, 130, 132),415. 416 Subers, E., 329(47), 332 Sue, F., 48(16), 72 Sugden, P. H.,81(19), 82(20), 88(59), 89(20, 59), 93, 94 Sugimori, M., 289(32), 290(32), 305(32), 312 Sugimoto, T., 272(175), 282 Sugimura, T., 330(64), 333 Sulakhe, P. V., 300(170), 315 Sullivan, K., 377(146), 380 Sullivan, P. T., 236(61), 279 Sutherland, E. W., 152(37), 162(37, 79), 163(37, 113, 115), 173, 174, 175, 226(49), 2 78 Sutton, C., 449(166), 455 Suyama, K., 70(127), 74

497

AUTHOR INDEX Suzuki, H., 366(67), 378, 387(28), 391(54), 393(28, 64),395(71), 414, 415 Swarup, G . , 349(96), 356 Swenson, T. L., 127(39a), 137(39a). 144 Sy, J., 36(197), 41(197), 46, 431(41, 42). 452 Sypherd, P. S . , 436(76, 77), 453 Sze,P. Y.,276(193), 277(193), 282 Szent-Gyorgyi, A. G., 384(5, 8, 9). 388(8, 9, 31), 389(39), 390(48, 49), 391(55), 406(8), 413, 414 Szentkiralyi, E. M., 384(8), 388(8), 391(55), 406(8), 413, 414 Szmigielski, A., 293(65, 66). 312, 313 Szyszka, R., 431(44), 452

T Tabuchi, H.,297(133), 314 Tada, M., 341(36), 355 Tager, H. S., 35(186), 46 Taira, T., 297(133), 314 Takabayashi, Y.,128(55), 130(55), 144 Takahashi, K.,387(28), 393(28), 414, 406(144), 417 Takahasi, J., 297(133), 314 Takai, Y.,242(71), 257(137), 265(137, 153a, 153b), 279, 281, 293(71), 298(135, 136, 137, 138, 144, 146), 303(138), 308(236), 313, 314, 317, 430(39), 431(39), 444(39), 452 Takayama, S., 330(80), 333 Taketa, K.,32(151, 152), 45 Takeuchi, K.,407(161), 417 Takimoto, S., 297(133), 314 Tallant, E. A., 309(249), 317 Tallman, J. F., 324(13), 331 Tana, J. B., 12(38), 42 Tanabe, T., 124(2), 126(2), 128(2, 52), 129(52), 143, 144 Tanaka, E., 300(169), 303(169), 306(169), 307(169), 315 Tanaka, K., 207(173), 214 Tanaka, R. D., 181(17), 187(51), 190(17, 68), 192(89, 91), 202(17), 210, 211, 212 Tanaka, T., 48(5, 6, 7, 11, 16), 49(5), 50(5, 27), 68(11, IIO), 72, 74 Tanaka, Y., 191(82), 212 Tanenbaum, M., 396(83), 398(83), 415 Tanigawa, K.,308(236), 317 Taniguchi, H., 308(236), 317 Taniguchi, J., 338(14), 354

Taniuchi, K., 50(27), 72 Tank, D. W . , 349(99), 357 Tanzawa, K., 182(26), 191(26), 211 Taparowsky, E., 377(144), 380 Tarlow, D. M . , 128(61), 129(61), 132(61), 134(I05), 145 Tamowski, W., 58(68), 73 Tashima, Y.,32(158), 45 Taunton, 0. D., 27(108, 109), 34(108, 109), 44, 50(31), 65, 68(109), 72, 74 Tavare, J. M., 141(124), 143(124), 146 Taya, Y., 191(82), 212 Taylor, D. A., 293(80), 313 Taylor, E. D., 385(14), 391(14), 413 Taylor, E. W., 391(57, 58, 59), 392(57, 58, 59), 393(57), 414 Taylor, K. A., 387(28), 393(28), 407(163), 414, 417, 443(123), 454 Taylor, M. K., 116(56), 120 Taylor, S. S., 321(6), 331 Taylor, W. M . , 88(51), 94 Teague, W. M., 82(22), 84(35, 37), 85(37), 86(35, 37), 88(37), 93, 94 Teichberg, V. I., 302(194), 303(194), 310(194), 316, 329(43), 332, 348(86, 87), 356 Tejwani, G. A., 5(12), 11(35), 27(96), 32(12, 156, 163), 41, 42, 43, 45 Tellez-Inon, M . T., 435(69), 453 Teplow, D. B., 377(143), 380 Terenzi, H. F., 435(68, 69), 453 Testa, U.,330(79), 333 Thampy, K. G., 141(128a), 146 Theibert, A., 447(149a), 455 Theurkauf, W. E., 291(52, 53). 306(217, 219), 312, 316 Thevelein, J. M., 433(53), 452 Thiemann, V., 58(68), 73 Thiessen, B. J . , 300(170), 315 Thoma, F., 440(100), 453 Thomas, A. A. M., 360(5), 377 Thomas, E. L., 70(126), 74 Thomas, G., 438(95), 453 Thomas, N. S . B., 364(58, 59), 367(58, 59). 368(58, 59), 369(59), 378 Thompoulos, P., 330(79), 333 Thompson, B., 162(80), 174 Thompson, M. P., 91(95), 95 Thompson, R. J., 291(50), 312 Thompson, W. J., 165(121), 175, 294(88), 313

498 Thrall, T., 130(78), 131(78), 145 Tichonicky, L., 69(116), 74 Tipper, J. P., 128(50), 129(68), 131(50), 133(50), 137(50), 138(50), 139(50, I16), 140(50, 121), 143(132),144, 145, 146, 165(146),176 Tipton, K. F., 32( 1 5 3 , 45 Titanji, V., 50(36), 51(37), 58(36), 60(37), 61(36), 72 Titanji, V. P. K.,53(47), 62(76, 84), 63(87), 64(89, 90),73, 74 Titchner, E. B., 124(9, lo), 143 Todaro, G. J., 330(60), 333 Tomita, Y., 188(55), 211 Tong, J. H., 273(181), 274(181), 282 Tonks, N. K., 296(123), 314 Tonomura, Y., 389(41), 390(41), 392(62), 406(144), 414, 417 Tooth, P. J., 395(70, 74), 415 Tormanen, C. D., 201(151), 214 Tornquist, H., 148(1, 4, 9), 149(1, 4), 150(1), 151(9), 153(1), 166(4), 168(4), 172 Toms, H. N., 435(68, 69, 70, 71), 453 Tomella, M., 427(35), 452 Tortora, P., 434(61), 453 Tourian, A., 219(18), 221(18), 222(18), 234(18), 236(18), 278 Tovar, G., 448(154), 455 Towle, H. C . , 187(47),211 Toyo-Oka, T., 207(181), 214 Toyoda, K.,36(197), 41(197), 46 Trachsel, H., 362(12, 36), 369(73), 371(12), 373(97), 375(12), 377, 378, 379 Traniello, S . , 14(48), 32(158), 42, 45 Traugh, J. A., 299(149), 303(149), 315 Trautwein, W., 338(14, 15, 16), 339(16), 340(29), 341(15), 342(139), 354, 355, 357 Treadwell, C. R., 169(169), 176 Trentalance, A., 205(168, 169), 214 Trevillyan, 436(74), 453 Trifaro, J. M., 297(130), 314 Triggle, D. J., 341(38), 355 Tritthart, H. A., 353(123), 357 Trotter, J. A,, 407(162), 408(167, 172), 417 Trujillo, J. L., 27(44), 44 Trundle, D., 19(56), 42 Truscott, R. J. W., 250(98), 258(98), 260(98), 270(163), 280, 281 Trybus, K.M., 387(30), 390(45b), 393(30, 65), 394(65), 395(65), 414, 415 Trzaskos, J. M., 194(96),212

AUTHOR INDEX Trzeciak, W. H.,169(161, 167, 170). 171(170), 176 Tsai, M. Y., 27(105), 44 Tsai, S.-C., 152(38), 173 Tsien, R. W . , 337(5, 130, 134). 338(12), 340(20, 23). 342(43), 344(43), 354, 355, 357 Tsou, K.,305(208), 316 Tsukamoto, T., 69(123), 70(127), 74 Tsuruhara, T., 162(90), 169(90), 170(90), 174 Tung, H. Y . L., 62(81), 73, 165(150), 176, 303(234), 308(234), 317 Turner, R. S., 298(142), 307(142, 224), 314, 316 Tutwiler, G. F., 34(175), 45 Twible, D. A,, 67(106), 74

U Udenfriend, S., 249(88, 89, 91). 252(91, 111, 114, 1 1 9 , 271(172), 279, 280, 282 Ueda, M., 165(127),175 Ueda, T., 295(99), 303(99), 305(209a), 313, 316 Ueno, T., 406(147), 417 Ui, M., 66(97), 74, 376(140), 380 Ulrich, J., 299(156), 315 Underwood, A. H., 27(107), 32(153), 44, 45 Unger, R. H., 244(77, 78), 245(78), 279 Uno, I., 431(48, 49), 432(48, 49, 50, 51), 433(54), 435(67), 439(97, 98), 452, 453 Urihe, E., 427(32), 452 Ushiro, H.,330(53), 332 Utter, M. F., 48(1), 50(1), 72 Uyeda, K.,5(3, 10, 16), 6(16, 21). 7(16), 12(16, 43), 13(43), 14(43), 15(16, 43), 21(16, 58, 59), 22(3, 16, 61, 63), 23(43), 27(94, 95), 28(3, 117), 29(3, 117, 123, 129, 130, 134), 30(140), 31(21), 36(198), 37(61, 199, 201), 40(21, 134), 41(21), 41, 42, 43, 44, 46

V Vaartjes, W. J., 130(91), 131(91), 132(91), 145 Vagelos, P. R., 124(3, 12), 125(12), 126(3), 128(3), 143, 152(44), 155(44), 156(44), 173 Vahouny, G. V.. 169(169), 176 Vainchenker, W., 330(79), 333 Valenzuela, P., 432(52), 452

499

AUTHOR INDEX Vall, R. J., 1 l(37). 42 Vallee, R. B., 291(52, 53), 300(167), 306(217, 219), 312, 315, 316 Vallejos, R. H., 427(34, 35), 452 Vanaman, T. C . , 295(108), 314 Van Berkel, T. J. C., 48(10), 50(25), 53(46), 55(25), 57(63), 65(92), 72, 73, 74 Vance, D. E., 124(4), 126(4), 143, 171(187), 177, 200(145), 201(145), 213 Vance, J. E., 171(186), 177 van de Meene, J. G.C., 442(114), 454 van den Berg, G.B., 53(46), 57(63), 73 van den Berg, J. W. O., 32(154), 45 Vanderhoff, G.,362(9), 377 Vanderhoff, G.A,, 362( 1I ) , 377 Vandlen, R. L., 329(41), 332, 348(94), 356 van Dongen, C . J., 309(241), 317 Van Dop, C . , 376(141), 377(146), 380 van Driel, R., 444(131, 136), 445(131), 454 van Golde, L. M. G.,200(146), 201(146), 213 Van, Jett, M.-F., 28( 116), 29( 116), 44 van Laere, A. J., 437(85), 453 Van Loere, A., 29(121), 44 Van Obberghen, E., 143(137), 146, 165(144, 147), 176 van Renswoude, J., 331(86), 333 Van Schaftingen, E., 5(2), 6(24), 14(46, 47). 15(2, 49), 22(2, 46, 47, 49, 62), 23(49, 64).24, 26(85), 27(85), 28(2, 116), 29(2, 24, 116, 119, 120, 121, 125), 32(165), 33(165, 171), 37(46, 49). 40(85), 41, 42. 43, 44, 45, 132(99), 145, 433(55, 56), 453 Varnado, C. E., 206(172), 214 Vary, T. C . , 116(51), I20 Vassort, G.,337(10), 354 Vaughan, M., 149(10), 152(38), 157(68), 162(82), 163(82, 109), 165(118, 119, 129, 130, 139), 171(189), 173, 174, 175, 177, 377(147), 380 Vaughan, R. L., 444(130), 454 Veech, R. L., 85(44), 94, 125(19, 20), 126(20), 129(20), 130(20), 144 Veerkamp, J. H., 109(51), 115(51, 54), 120 Veldhuis, H. D., 309(237), 317 Veldhuizen, J. A. M., 109(51), 115(51), 120 Veloso, D., 85(44), 94, 125(19), 144 Venkataramu, S. D., 16(50), 21(50), 42 Venkatesan, S., 187(49), 211 Venson, V., 298(134), 314 Venter, J . C . , 341(38), 355

Vergara, C., 342(42), 345(42), 355 Verger, R., 155(57, 58), 156(61), 174 Veron, M., 444(129, 135, 137), 445(129), 454

Veselovsky, N. S., 340(25), 355 Vibert, P., 390(50), 414 Vicalvi, J. J. Jr., 228(53), 279 Vigny, A,, 251(105), 280 Vining, R., 169(162), 176 Virmaux, N., 408(169), 417 Vissers, S., 29(125), 44 Vlahcevic, Z. R., 201(153), 214 Vogel, R. L., 276(195), 277(195), 282 Vogt, B., 126(34), 138(34), 144 Voloch, Z., 375(126), 380 Volpe, J. J., 124(3), 126(3), 128(3), 143, 182(34), 211 von Euler, U. S . , 252(109, 112), 280 von Saltza, M. H., 218(5), 236(5), 278 Voorma, H. O . , 360(5), 363(46), 364(57), 377, 378 Vorobetz, Z. D., 287(26), 289(26), 290(26), 303(26), 311, 340(30), 355 Vrana, K. E., 255(131), 260(131), 270(165), 271(165), 280, 281 Vu, N.-D., 393(63a), 408(63a), 415 Vulliet, P. R., 255(130), 262(130), 264(153), 266(153), 280, 281

W Wadzinski, I. M., 271(171), 282 Wagenmakers, A. J., M., 109(51), 115(51, 54). 120 Wagner, J. D., 7(30), 37(30), 42, 67(107), 74, 229(56), 241(56), 279 Wagner, P. D., 390(47), 393(63a), 407(47), 408(47, 63a), 414, 415 Wahba, A. J., 362(42), 363(42), 364(60), 366(60), 378 Waisman, H. A,, 271(171), 282 Wakabayashi, T., 393(66, 69), 395(66, 69), 415

Wakil, S. J., 124(6, 9, lo), 141(128a), 143, 146 Walaas, S. I., 286(8), 289(34), 290(8, 34, 35, 36), 291(35, 36), 294(35, 36, 96). 296(116), 297(35, 36, 116), 298(35, 36, 96, 147), 301(8, 35, 36, 96), 302(177, 178, 183), 303(8, 227), 304(8), 306(36), 307(177, 178, 227), 308(8, 35, 36, 96,

AUTHOR INDEX 147), 309(36, 147), 310(8), 311, 312, 313, 314. 315, 316 Walderhaug, M., 16(50), 21(50), 42 Waldo, G. L., 327(20), 332 Walker, J. E., 105(35), 108(35), 120 Walker, R. G . , 54(51), 73 Wallace, A. V., 165(143), 176 Wallace, R. W., 309(249), 317 Waller, C. A., 81(19), 93 Wallimann, T., 390(48, 50), 414 Wallis, M. H., 371(85), 379 Walser, M., 98(2), 99(2), 119 Walseth, T., 7(28), 42 Walsh, D. A., 82(24), 93, 152(40), 153(40), 173, 293(64), 312, 436(73), 453 Walsh, K., 425(27), 452 Walsh, M. P., 384(10), 396(89), 397(97), 398(89), 413. 415 Walter, P., 34(181), 45, 182(32), 211 Walter, R. D., 448(151, 152, 153), 455 Walter, U., 287(21), 288(21), 290(21, 38), 291(38, 47, 48, 56), 292(38), 293(38, 70, 75, 76, 77), 303(21), 306(217), 311, 312, 313, 316, 336(2), 354 Walton, G. M., 169(163), 176, 293(69), 313, 362(43), 363(43, 45), 378 Wancewicz, E., 171(186), 177 Wancewicz, E. V., 151(31), 170(31), 173 Wang, J. K.-T., 298(147), 299( 148), 308(147), 309(147), 314, 315 Wang, J. Y . J., 423(16, 17), 425(24), 452 Wardazala, L. J., 143(133), 146 Warner, R. C . , 127(40), 144 Warnette-Hammond, M. E., 34(177), 45 Watanabe, A., 32(162), 45 Watanabe, S., 386(26), 387(28), 392(62), 393(28, 64),414, 415 Watkins, P. A., 128(61, 62), 129(61, 62, 63), 130(63), 132(61, 62, 63), 145, 165(130), 175 Watson, D. C . , 51(38), 60(38), 72, 81(18), 93 Watson, J. A., 185(43), 187(52), 192(52, 85), 202(52), 211, 212 Watters, G. T., 126(29), 127(29), 144 Watterson, D. M., 401(110a), 416 Way, S. C . , 181(15), 210 Waymack, P. P.,116(55), 120 Waymire, J. C . , 255(132), 256(132, 136), 264(136), 280 Webb, W. W., 349(98, 99), 356, 357 Weber, A., 48(8), 72

Weber, G., 27(88), 43 Weber, H. W., 165(125), 175 Weber, L. A., 373(101), 379 Weber, W., 330(55), 332 Weeks, M. O., 321(7), 331 Weidemann, M. J., 165(140), 176 Weigers, S. E., 339(18), 340(18), 355 Weinberg, C. G . , 349(106), 357 Weiner, N., 252(110, 116, 117), 255(130), 262(130), 265(156), 266(157), 267(156, 157), 268(157, 160), 269(157), 280, 281 Weinstein, I. B., 330(56, 57), 332 Weiser, P. C . , 130(87), 131(87), 145 Weiss, C., 58(68), 73 Weiss, J., 338(13), 354 Weiss, L., 88(52), 94 Weissbach, H., 271(172), 282 Weller, M., 310(253), 317 Wells, W. W., 208(198(, 215 Weng, L., 31(144), 35(144), 44 Wernette-Hammond, M. E., 12(39, 40), 42 Werth, D. K., 386(19), 413 Westhead, E. W., 69(119, 120, 122), 70, 74 Westwood, S. A,, 404(125), 416 Wick, U., 443(124), 454 Wheeler, S., 253(122), 254(122), 280 Wheeler, T. J., 143(133), 146 White, D. A,, 192(88), 193(94), 212 White, H. D., 310(254), 317 White, M. F., 330(80), 333 Whitehead, T. R.,201(153), 214 Whitehouse, S . , 78(5), 84(34), 93. 94 Whittaker, J., 330(76), 333 Whittemore, S . R.,302(186), 303(186), 315 Wi,Zham, P. S . , 181(15), 210 Weczorek, C. M., 302(180), 315 Wieland, 0. H., 78(6), 85(40), 88(52, 53, 54), 89(6, 68), 90(78, 79), 91(84), 93, 94, 95, 129(64, 65), 143(138), 145, 146 Wigler, M., 377(144), 380 Wilce, P. A,, 192(84), 193(84), 205(167), 212, 214 Wilden, U., 328(30, 38), 332 Wiley, M. H., 191(76, 79), 198(133), 205(165), 212, 213, 214 Wilgus, H., 222(34), 223(34), 224(34), 225(44), 226(34, 44), 227(44, 50), 228(44, 52), 230(50), 231(50), 232(50), 239(50), 240(44), 241(50), 245(50), 248(44), 262(44), 278 Williams, B. R. G., 375(121), 380

AUTHOR INDEX Williams, K. R., 25(72), 43, 307(230), 316 Williams, P. E., 116(62), 121 Williams, R. C . , Jr., 292(60), 312 Williams, R. H.,165(121), 175 Williamson, D. H., 130(81, 84, 86, 96), 131(81, 84, 86), 132(84), 145 Williamson, J. R., 88(50), 89(66), 94, 98(2), 99(2), 119 Willner, J., 69(125), 74 Wilson, F. D., 287(18, 21), 288(18, 21). 290(18, 21), 303(18, 21), 311, 343(56),356 Wilson, J. M., 442(113), 454 Wilson, P. G . , 449(166), 455 Wilson, S. R . , 165(143), 176 Windisch, H., 353(123), 357 Windmueller, H. G . , 130(72, 73). 145 Wingender-Drissen, R., 434(62), 453 Winkelmann, D. A., 390(51), 414 Winkleman, L., 406(137), 416 Winton, B., 88(53), 94 Wirtz, K . W. A,, 309(241), 317 Wise, B. C . , 294(95, 97). 298(97, 140), 313, 314 Wise, K. L., 116(62), 121 Wise, L. S . , 166(157), 176 Wise, R . C . , 257(140), 281 Wiser, M. F., 448(156), 455 Withy, R. M., 351(111), 357 Witte, L. D., 191(78), 192(78), 193(78), 212 Witters, L., 62(79), 63(79), 73 Witters, L. A., 165(146), 176, 126(26, 30, 34), 127(46), 128(46, 48, 50), 129(68), 130(88, 92), 131(50, 88, 92), 132(26, 46, 88). 133(50, 92), 134(26), 135(26, 109), 137(26, 50, 92), 138(34, 50, 92), 139(50, 92, 116), 140(46, 50, 92, 121), 141(126, 128), 142(128), 143(132),144, 145, 146 Wittmann, H.G., 421(4), 451 Woese, C. R., 185(39),211, 445(138), 454 Woffendin, C., 443139, 140), 446(141), 454, 455 Wohlhueter, R. M., 115(52), 120 Wolf, D., 375(128), 380 Wolf, H.,400(109), 405(109), 416 Wolff, D. J., 292(62), 312 Wong, A. J., 407(158), 417 Wong, E. H. A., 162(85, 86, 87), 163(86, 87), 174 Wong, S., 362(38), 363(38), 364(38), 378 Wong, S. T., 367(69), 379 Wood, H. G . , 124(11), 143

Wood, W. A., 14(48),42 Woodgett, J. P., 264(150), 281 Woodgett, J. R., 264(153), 266(153), 281, 296(123, 124), 297(124), 314 Woodley, C. L., 362(42), 363(42), 364(60), 366(60), 378 Woody, C. D., 289(33), 290(33), 293(33), 312 Wrenn, R. W., 257(140), 281, 294(95, 97), 298(97), 313 Wretborn, M., 223(38), 278 Wu, E. S . , 349(99), 353(99), 357 WU,T.-L., 80(16), 85(42), 93, 94 WU,W. C.-S., 294(96), 298(96, 139), 301(96), 308(96, 139), 313, 314, 329(41), 332, 347(78), 348(94),356 Wurtman, R. J., 273(182), 274(182), 282 Wurtzburger, R . J., 250(95), 280 Wyngaarden, J. B., 249(89), 279

Y Yaghmai, B., 362(21, 23). 363(21), 377, 378 Yamamoto, H.,296(111), 300(169), 303(169), 306(169), 307(169), 314, 315 Yamamoto, K., 328(37), 332 Yamamoto, S., 171(189), 177 Yamamoto, T., 171(189), 177, 190(64), 211 Yamamura, H., 293(71), 297(133), 313, 314, 430(39), 431(39), 444(39), 452 Yamanaka, G., 376(141), 380 Yamanishi, J., 242(72), 279 Yamato, S., 207(173), 214 Yamauchi, T., 251(102), 255(128, 129), 256(128), 257(138, 139), 258(128), 263(138, 139, 147, 148), 264(128, 129, 147, 148, 149), 272(175), 276(196, 197, 198), 277(147, 148, 196, 198, 199), 280. 281, 282, 294(93), 296(115), 300(166, 168), 302(166), 303(166), 306(220), 307(220),313, 314, 315. 316 Yasuda, H., 441(104), 454 Yasuda, S . , 408(170), 417 Yatsunami, K., 377(145), 380 Yeaman, S. J., 81(18), 82(22, 27), 83(27), 84(35), 86(35), 92(97), 93, 94, 95, 100(15), 101(15), 102(15), 104(15, 29, 32, 33, 34, 3 3 , 105(15, 32, 33, 34, 3 9 , 107(33, 38), 108(35), 109(15, 29), 110(38), 120. 149(19), 150(19), 151(19, 30, 34, 3 3 , 169(34), 170(35), 173

AUTHOR INDEX Yeh, L.-A., 129(66, 67), 145 Yellen, G., 340(20), 355 Yellowlees, D., 154(53), 174 Yokayama, M., 5(3), 22(3, 61, 63), 28(3), 29(3), 37(61, 198, 199), 41, 43, 46 Yorek, M. A., 67(103), 74 Yorke, R. E., 125(17), 144 Yoshida, M., 444(125), 454 Yoshimasa, T., 325(18), 332 Yoshimura, N., 208(189), 215 Yost, D. A , , 402(113), 409(113), 416 Younathan, E. S . , 11(37), 27(100), 42, 43 Young, H. A., 321(7), 331 Young, J. D., 25(75), 43 Yu, B., 298(144), 314 Yumoto, N., 208(189), 215 " 6

Zahlten, R. N., 208(197), 215 Zallor, M. J., 321(6), 331 Zaltzman-Nirenberg, P., 252(114), 280 Zammit, V. A., 130(79, 83, 85). 131(79, 83,

85). 132(79, 83), 133(101), 137(101), 138(10l), 139(79), 140(101), 142(129), 145, 146, 204(163), 205(162, 163), 213 Zani, B. M., 351(148), 358 Zapalowski, C., 116(60), 121 Zaremba, T., 303(200), 304(200), 316 Zavoico, G. B., 329(47), 332 Zeeberg, B., 292(58), 312 Zetterqvist, 0.. 25(76, 82), 43, 50(36), 51(37, 41), 53(47), 58(36), 60(37), 61(36, 74), 62(74, 76), 64(89, 90, 91). 72, 73, 74, 200(142), 201(142), 213 Zick, Y., 143(136), 146, 330(54, 74, 75, 76, 77), 332. 333 Ziea, S . , 48(17), 72 Zilberstein, A., 372(90), 375(116), 379, 380 Zillig, W., 422(7), 451 Zimman, B., 165(120), 175 Zucker, W. V., 362(7), 377 Zwergel, E. E., 126(35), 144 Zwiers, H., 298(143), 309(143, 237, 239, 240), 314, 317 Zwiller, J., 302(187, 188), 303(187, 188), 315

Subject Index A

Acylglycerol lipase, 168 Acylphosphate, 422 Adenylate cyclase, 166, 435 desensitization, 323-324 Adipose tissue, brown, 170 Adipose tissue lipase see Lipase, hormonesensitive ADP-ATP exchange, 12-13 ADP-ribosyltransferase, 377 Adrenal cortex, 151 steriodogenesis, 169 Adrenal medulla, tyrosine hydroxylase, 24925 1 activation, 255 Adrenaline fatty acid synthesis, 129 lipolysis effect, 166- 167 pyruvate dehydrogenase complex, 89 Adrenergic agonists, 160 acetyl-CoA carboxylase, 131-132 desensitization heterologous, 321-327 homologous, 327-328 6-phosphohcto-I-kinase effect, 31 regulation, 320-321 Alanine, pyruvate kinase inhibitor, 50, 54, 61 Alzheimer’s disease, 299 Amino acid, branched-chain, regulation, 9899 Amino acid hydroxylase, aromatic, introduction, 218-222 cAMP compartmentalization, 163 cAMP phosphodiesterase, I65 Amphotericin B, 439 Angiotensin, 37 acetyl-CoA carboxylase, 13I - I32

Acanthamoeba castellanii, 443 myosin regulation, 406-407 Acetate, 128 Acetoacetyl-CoA, 1 17 Acetyl coenzyme A carbon-dioxide ligase, 124 pyruvate dehydrogenase kinase, 83-84 Acetyl-coenzyme A carboxylase, 117 allosteric regulators, 128-129 hormonal regulation, 130-134 introduction, 123- 125 molecular forms, 125-128 phosphorylation hormone effect, 135-138 reversible, 134- 135 protein kinases, 138-141 protein phosphatases, 141-142 Acetylcholine receptor desensitization, 350-35 I phosphorylation biochemical studies, 348-349 significance, 349-351 subunit phosphorylation, 350 ACTH, 309 hormone sensitive lipase, 160, 162 steriodogenesis, 169 Actin filament, structure, 382 Activator protein branched-chain ketoacid-dehydrogenase, 109-1 12 diet effect, 115 properties, 1 10- 1 12 purification and characterization, 1 10 Acyl-CoA:cholesterolacyltransferase, 180, 187, 209 503

504

SUBJECT INDEX

Angiotensin I1 phenylalanine hydroxylase, 241 pyruvate kinase, 66 Aplysia, neuron, 341, 343-345 function, 287-290 Ascaris suum, 6-phosphofructo- 1-kinase, 3 132 Ascorbic acid, 205-206 ATP, pyruvate dehydrogenase kinase, 83-84 Azatobacter vinelandii, pyruvate dehydrogenase, 80

B B-50 protein, 309 Bacteria intrinsic protein kinases, 422-424 isocitrate dehydrogenase, 425, 428 photosynthesis, 426-428 sugar transport, 425-426 Bacteriochlorophyll, 426 Bacteriophage, protein kinase encoded, 42 1422 Bacteriorhodopsin, 426-427 Bacillus stearothermophilus, pyruvate dehydrogenase, 80 Bile acid, binding of, 188 Biopterin, 218 Biotin, 124 Blastocladiella, 436 Blastocladiella emersonii, protein phosphorylation, 438-439 Brain phosphoprotein, 300-309 protein, phosphorylation see Protein phosphorylation, brain protein kinase, 290-300 protein phosphatases, 309-3 10 tryptophan hydroxylase properties, 274-275 regulation, 275-277 C Calciductin, 341 Calcineurin, 3 10 Calcium acetyl-CoA carboxylase, 141- 142

channel modulation evidence, 337-341 phosphorylation studies, 341-342 ionophore A23 187, 24 1 potassium channel activation, 345-346 pyruvate dehydrogenase phosphatase, 8 5 , 86, 90 pyruvate kinase, 70 smooth-muscle myosin, 388-389 tryptophan hydroxylase activation, 275-277 Calcium-calmodulin kinase 11, 305 Caldesmon, 397 Calmodulin, 325 myosin light chain kinase, 400-403 pyruvate kinase, 70 Calpain, 207-209 Candida utilis, 435 Carboxyamidomethylation, kinase stimulation, 19 Casein, 43 1, 446 Casein kinase, 141, 299 Catecholamines pyruvate kinase, 69 synthesis, 252, 256 Caulobacter crescentus, 422 Cuvia, neuronal function, 289 Cell division, blockage, 191 Cellular proliferation, HMG-CoA reductase, 205 Chloramphenicol, 423 Cholesterol biosynthesis, 168 feeding, 200-202 homeostasis determinants, 186- 187 HMG-CoA reductase, 185-188 secretion, 187 Cholesterol ester hydrolase, 151, 168 a-Cholesterol hydroxylase, 180, 187, 209 Cholestyramine, 188 Chromaffin cell, tyrosine hydroxylase, 255258 Chromatofocusing, pyruvate kinase, 57-58 Citrate, acetyl-coenzyme A carboxylase, 128129 Clofibrate, 116 Cloning, HMG-CoA reductase, 191-192 Compactin, HMG-CoA reductase, 19 I Contractile activity, regulation actin-linked, 382-384

SUBJECT INDEX

505

cytoplasmic myosin, 406-412 invertebrate, 406 myosin-linked, 382, 384 phosphorylation-dependent, 382, 384-385 smooth-muscle, 386-403 striated muscle, 404-406 Coprinus macrohizus, 439 Corpus luteum, 151 steroidogenesis, 170 CTP:phosphocholine cytidylytransferase, I71

D DARPP-32, 302, 307-308 Desensitization, 350-35 1 heterologous, 321-327 homologous, 32 1-327 Diabetes, 100 alloxan, 113 phenylalanine hydroxylase, 230 pyruvate-dehydrogenase complex, 88 Diacylglycerol lipase, 15 I Dibutyryl-CAMP, 268-269, 277, 343 tyrosine hydroxylase activation, 253 Dichloroacetate, I 16 Dictyostelium, 436 Dictyostelium discoideum myosin phosphorylation, 442-443 regulation, 406-407 protein phosphorylation, 442-443 Diet branched-chain ketoacid-dehydrogenase, 113-119 phenylalanine hydroxylase, effect, 246 pyruvate kinase effect, 65 Dihydrolipoly dehydrogenase, 101 Dihydropteridine reductase, 2 19 Dihydroxyphenalanine, 248 6,7-Dimethyltetrahydropterin,2 I8 2,3-Diphosphoglycerate, pyruvate kinase regulation, 69 Discoidin, 446 5,5’-Dithiobis(2-nitrobenzoicacid), 83 Dithiothreitol, 83 Diurnal variation, HMG-CoA reductase, 188, 192- 193 Dopa, biosynthesis, 248 Dopamine, 248

E EGTA, 325 Endocytosis, receptor-mediated, 190 Endoplasmic reticulum microenvironment, 194 reductase activity, 189 Enzyme activity, regulation, 437 yeast, 433-435 synthesis, regulation, 432-433 Epidermal growth factor, 304 Epidermal growth factor receptor, regulation, 329-330 Epinephrine acetyl-CoA carboxylase, 131- 134 phosphorylation, 136-138 biosynthesis, 248 phenylalanine hydroxylase, 239-240 6-phosphofructo-I-kinaseeffect, 3 1 6-phosphofructo-l-kinase-fnrctose-2,6bisphosphate control, 37 pyruvate kinase, 66-67 Ergocryptine, phenylalanine hydroxylase, 240 Erythrocyte, pyruvate kinase, 68-70 Escherichia coli protein phosphorylation, 42 I , 423-425 pyruvate dehydrogenase complex, 79-80 Esterdiol, 304 Eukaryotic initiation factor, 360 dephosphorylation, 367-368 phosphorylation effect, 364-366 recycling, 362-364 reversing factor-phosphate complex, 366368 Eukaryotic initiation factor kinase dsRNA-dependent, 37 1-373 heme-regulated, 369-371

F Fatty acid oxidation, 88 synthesis, hormonal regulation, 130-1 34 Fatty acid synthetase, 124 Felix, neuronal function, 289 3-Fluoromevalonate, 192 Follicle-stimulating hormone, 170 Forskolin, 162

506

SUBJECT INDEX

Fructose, 128 pyruvate kinase regulation, 48 Fructose-I ,6-bisphosphatase, 207, 434 phosphorylation site, 35-36 role, 4 specific activity, 5 Fructose 1,6-diphosphate, 426 pyruvate kinase effect, 50, 55, 59-61 Fructose-l,6-diphosphatase, 50 Fructose-2,6-bisphosphatase assay, 6-7 hormonal control, 37 low-molecular-weight effectors, 22 product-inhibitor specificity, 15 reaction mechanism, 16- 18 substrate inhibition, 15-16 specificity, 14 Fructose-6-phosphate, inhibition effect, 15- 16

G Glucagon, 27-28, 35 acetyl-CoA carboxylase, 125, 128- 130 phosphorylation, 136-138 HMG-CoA reductase activity, 202-205, 208 hormone-sensitive lipase, 160, 162 phenylalanine hydroxylase, 240-241, 243346 in vivo effect, 230-234 phosphorylation role, 226-230 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase, 37 pyruvate kinase, 65-67 regulation, 48 release, stimulation, 244 Glucagon-insulin ratio, HMG-CoA reductase, 192-193 Gluconeogenesis, 48 6-phosphofructo-2-kinase-fructose-2,6bisphosphatase, 37-40 pyruvate kinase regulation, 48 regulation, 48 yeast, 434 Glucose, 116 transport, 125 Glucosed-phosphate, protein synthesis, 371 Glutamate dehydrogenase, 207, 435 Glutathion triphosphate, 365-366 Glutathione, oxidized, 360

a-Glycerol-P, fructose-2,6-bisphosphataseeffect, 15-17 Glycogen phosphorylase, 108 Glycogen synthase, 296 Glycogen synthase kinase, calmodulin-dependent, 242 Glycolysis, 6-phosphofructo-2-kinase-fructose-2,6-bisphosphatase role, 37-40 Growth hormone, hormone-sensitive lipase, 160-161, 166 Guanine nucleotide-binding proteins, synthesis, 376-377

H H-7, 41 1 Halobacterium halobium, 42-427 Helix, neuronal function, 289 Helix roseneri, 346 Heme regulated initiation kinase, 369-37 1 biological significance, 373-376 inhibitors, 373-376 Heme-deficiency, protein synthesis, 360 Heme-regulation, eukaryotic initiation factor kinase, 369-371 Hemin, 370-373 Heparin, tyrosine hydroxylase activation, 25 I Hepatocyte, pyruvate kinase, 48 Hermissenda. neuronal function, 289 Hermissenda crassicorenis, 345 Histone kinase, 442 Histone H I , phosphorylation, 440-442 Histone phosphatase, 62 Hydrogen peroxide, 18, 90-91 Hydroxycholesterol, 189, 190 Hydroxymethylglutaryl-CoA reductase (HMGCoA reductase) cell lines, 191-192 cholesterol homeostasis, 185- I88 compactin, 191 degradation, 206-210 diurnal variation, 188, 192-193, 203-205 effectors, 194 introduction, 180 mevalonate, 190 mevinolin, 191 microenvironment, 194 modulation, 198- 199 phosphorylation intracellular, 199-206 reversible, 195- 199

SUBJECT INDEX

507

proteolysis, 207-209 quantity, 190 receptor-mediated endocytosis, 190 regulation, 188-195 topology, 180- I85 Hydroxysterols, HMG-CoA reductase, 190-191 5-Hydroxytryptophan, biosynthesis, 273 Hyperphenylalanemia, 243

I Immunoglobulin E, 304 Immunoglobulin E receptor, regulation, 33033 1 Insulin, 38, 116, 188,304 acetyl-CoA carboxylase, 128- 134 phosphorylation, 135- 138 HMG-CoA reductase activity, 202-205, 209 hormone-sensitive lipase, 158-166 phenylalanine hydroxylase, 230 pyruvate kinase, 63,65,68 regulation, 48 pyruvate-dehydrogenase complex, 89-92 Insulin receptor, regulation, 330 Insulin-glucagon ratio, HMG-CoA reductase, 192- 193 Interferon, 372 Intestine, pyruvate kinase, 68 Ion channel regulation acetylcholine receptor, 347-35 I calcium, 337-342 introduction, 335-337 potassium, 342-347 sodium, 351-356 Isocitrate, 128 Isocitrate dehydrogenase, bacterial, 425,428 Isocitrate dehydrogenase kinase-phosphatase, 425 Isocitrate dehydrogenase-phosphatase, 425 Isoleucine, regulation, 98-99 Isopentyl adenine, 191 Isoprenaline, 241 Isoproterenol, 162,341-342 I -(5-Isoquinolinesulfonyl)-2-methylpeperazine, 41 1

K Kemptide, 439 Ketoacid-decarboxylase, branched-chain activator protein, 109-1 12

coenzymes, 100-101 components, 101-103 diet effect, 113-1 19 discovery, 100 immunoassay, 118 introduction, 97-99 kinase reactions, 104- 107 kinetics, 106 molecular weight, 102 phosphatase reactions, 107- 109 protein inhibitor, 109-1 12 regulation, discovery, 103-104 reversible phosphorylation significance, 112-118 substrate, 100-101 a-Ketoacid dehydrogenase, complex, 78 a-Ketoacid dehydrogenase kinase, 92 a-Ketoacid dehydrogenase phosphatase, 92 Ketoisoleucine, 100 Ketoleucine, 1 16 Ketovaline, 100, 116 Kidney, pyruvate kinase, 48, 68 Kinase reaction, branched-chain ketoacid-dehydrogenase, 104-107 Kluyveromyces fragilis, 36, 43 1

L Lanosterol, 191 Leiotonin, 397 Leucine, 116 regulation, 98-99 Leupeptin, 209 Lipase diacylglycerol, 15 1 hormone-sensitive insulin effect, 158-166 location, 147-148, 150 multifunction role, 168-171 phosphorylation activity, 157- 160 properties, 148-151 reversible phosphorylation, 152- 156 short-term control, intact cell, 156-161 lipoprotein, 155 monoacylglycerol , 15 1 pancreatic, 155 Lipase kinase, 153 Lip01ysis control, I61- I66 regulation hormone-sensitive lipase, 156- 166

508

SUBJECT INDEX

Lipolysis, regulation (conr.) other mechanisms, 166-168 stimulation, 156 Lipolytic activation cascade, 152, 156, 162 Lipoprotein, very-low-density, 188 Liver, pyruvate kinase, 5 1-68 Logio, neuronal function, 289 Luteinizing hormone, 170

M Magnesium, 426 acetyl-CoA carboxylase, 128 pyruvate dehydrogenase kinase, 83 pyruvate dehydrogenase phosphatase, 84-85 pyruvate kinase, 53-54 tryptophan hydroxylase activation, 276-277 Malaria, 448 Malonyl-CoA, 125 Manganese, 426 pyruvate dehydrogenase kinase, 83 MAP-2, 296 Maple Syrup Urine Disease, 99 Meiosis, CAMP-dependent protein kinase, 43 1-432 Membrane replacement synthesis, 185 Memory, short-term, 347 p-Mercuribenzoate, kinase stimulation, 19 Meromyosin, 387 6-Methyltetrahydropterin, 2 18 Mevalonate HMG-CoA reductase, 190, 192 NADP+ oxidoreductase, 180 Mevalonolactone, 200-202 Mevinolin, 191 MgATPase, actin-activated, regulation, 386387 Microtubule-associated protein 2, (MAP-2), phosphorylation, 306-307 Mitosis, CAMP-dependent protein kinase, 431432 Monoglycerol lipase, 15 1 Mucor rowcii, protein phosphorylation, 436437 Muscle contraction calcium regulatory systems, 436-437 smooth-muscle, 386-403 sustained, 397-399 invertebrate, 406

latch state, 397-399 striated, phosphorylation role, 404-406 Myelin basic protein, phosphorylation, 307 Myelin protein kinase, 299 Myosin cytoplasmic protein kinase C, 409-4 I2 regulation, lower eukaryote, 406-407 vertebrate phosphorylation, 407-409 isozymes, 406 light chain, 296 phosphorylation, 387-388 phosphorylation, Dicryostelium discoideum, 442-443 smooth-muscle actin-activated MgATPase activity, 386387 calcium effect, 388-389 phosphorylation conformation, 393-396 intact studies, 396-399 kinetics, 391-393 sustained contraction, 397 striated muscle, phosphorylation, 404-405 structure, 382-384 two head cooperativity, 389-391 Myosin heavy chain phosphatase, 443 Myosin light chain kinase, 297, 386, 396, 41 1 phosphorylation, 399-403

N NADH, pyruvate dehydrogenase kinase, 83- 84 NADPH generation, 371 Nerve stimulation, tyrosine hydroxylase, 266269 Nerve terminal protein, calcium-phospholipidregulated, 308-309 Neuronal function, protein phosphorylation, 287-290 Neurospora, protein phosphorylation, 435-436 Neurotransmitter acetylcholine receptor, 347-35 1 potassium channel regulation, 343-345 release, phosphorylation role, 287-290 synthesis, 220 Nicotinic acetylcholine receptor, regulation, 329 Nitrendipine, 341 Noradrenaline. 346

SUBJECT INDEX hormone-sensitive lipase, 158 lipolysis activation, 166 Norepinephrine, 355 acetyl-CoA carboxylase, 131- 132 biosynthesis, 248 phenylalanine hydroxylase, 240-24I pyruvate kinase, 66 Nystatin, 439

0 Octanoate, 116,128 Oligo synthetase, 372 Omithine decarboxylase, 207 2,3-Oxidosqualene cyclase, 191 Oxysterols, 190-191 Oxytocin, 166

P Paramecium tetraurelia, 448-449

PEP carboxykinase, 48 Phenylalanine, pyruvate kinase inhibitor, 50 Phenylalanine hydroxylase activation, 222,232-233 variables, 233-234 angiotensin 11, 241 enzyme-product species, 237-238 epinephrine effect, 239-240 ergocryptine, 240 forms, interconversion, 236 glucagon activation, 243-246 effect, in vivo, 230-234 human liver regulation, 247-248 norepinephrine, 240-241 phosphorylation forms, 223-224 in vivo, 226-230 ligand effect, 235-239 phosphorylation-dephosphorylation regulation, 242-248 physical properties, 221-224 propranolol, 240-241 reactions catalyzed, 218-219 tetrahydrobiopterin effect, 235-239 vasopressin, 241 Phenylephrine, pyruvate kinase, 66 Phenylketonuria, 243 Phenylpyruvate, I16

Pheochrornocytoma, tyrosine hydroxylase, 251 activation, 255,264 Phorbol ester, 346 Phosphatase inhibitor-I , 307-308 Phosphatase reaction, branched-chain ketoaciddehydrogenase, 107- 109 Phosphate activation, 6-phosphofructo-2-kinase, 13- 14 fructose-2,6-bisphosphataseeffect, 15-18 3-Phosphistidine, 16 Phosphodiesterase calcium-calmodulin-sensitivecyclic n u cleotide, 296 cGMP, 328-329 inhibitors, 69,355 Phosphoenolpyruvate-sugarphosphotransferase system, 423,425 Phosphofructokinase, 50, 125 6-Phosphofructo-l-kinase,433 activators, 27 Ascaris mum, 31-32 heart, 31 role, 4 specific activity, 5 6-Phosphofructo-2-kinase, 433 assay, 6 hormonal control, 37 low-molecular-weight effectors, 21-22 phosphoryl-acceptor specificity, 9-12 phosphoryl-donor specificity, 12 reaction mechanism, 12-14 6-Phosphofructo-2-kinase-fructose-2,6bisphosphatase action, 5 catalytic centers, 18-21 iodoacetarnide effect, 19-20 kinase stimulation, 19 low-molecular-weight effectors, 21-22 purification, 5-6 structural properties, 7-8 Phosphoprotein bacterial, 422-424 brain neuronal classes, 302-304 phosphorylation, 305-309 regional distribution, 301-302 tyrosine hydroxylase, 254 Phosphoprotein phosphatase, 286,435 Phosphorylase kinase, 37-38,154,297

510

SUBJECT INDEX

Phosphorylation control fructose-l,6-bisphosphatase,34-35 fructose- I ,6-phosphofructo- I-kinase, 27 6-phosphofructose-l-kinase,27-32 insulin effect, 135 intracellular , 199-206 reversibile adipose tissue lipase, 152- 156 biological significance, 1 12- I18 feedback control, 200-202 HMG-CoA reductase, 195- I99 regulation, 103- I12 Phosphorylation-dephosphorylation,phenylalanine hydroxylase, 242-248 Phosphoserine, 423, 427 Phosphothreonine, 423, 427 Phosphotyrosine, 423, 427 Phosphotyrosyl phosphatase, 309 Phosphovitin, 446 Phosvitin, 43 I Photosynthesis, bacterial, 426-428 Phosphoprotein phosphatase, 435 Physarum polycephalum, histone H 1 phosphroylation, 430-435 Pineal gland, 272 Polylysine, 447 Polymixin B, 439 Potassium pyruvate kinase, 53-54 pyruvate dehydrogenase complex, 86 Potassium channel calcium activated, 345-346 neurotransmitter-regulated, 343-345 phosphorylation, 346-347 Progesterone, 304 Prokaryote, protein phosphorylation, 421-429 Prokaryotic initiation factor, 377 Propanolol, hormone-sensitive lipase, 158 Propionyl-CoA carboxylase, 125 Propranolol, phenylalanine hydroxylase, 24024 1 Prostaglandin E2, 69 Protamine, 439 Protease, Ca2 -activated, pyruvate kinase action, 56 Protein 111, 305-306 Protein kinase, 286 acetyl-CoA carboxykase, 138- 141 activation mechanism, 286-287 +

autophosphorylation, 302-304 bacterial intrinsic, 422-424 bacteriophage encoded, 421-422 Blastocladiella, 438-439 brain, 290-300 calcium-dependent, brain, 294-299 calcium-phospholipid-dependent, 290, 298299 calmodulin-dependent, 257, 263-265, 290, 294-298 cytoplasmic myosin, 409-4 12 Dictyostellium disocideum, 442-447 membrane-bound serine, 165 miscellaneous fungi, 439-440 Mucor rouxii, 436-437 Neurospora crassa, 435-436 pol yamhe-dependent , 442 second messenger-regulated. 292 T7, 421 tyrosine-specific, 299-300, 330 yeast, 430-43 1 Protein kinase, CAMP-dependent, 91, 139141, 153-154, 156-157, 162, 224, 227, 329, 342 cell division, 431-432 enzyme synthesis regulation, 432-433 hormone-sensitive lipase, 153-154, 156I57 isozymes, 291 light chain kinase, 399-403 lipolysis, 162- 168 neuronal function, 287-293 pyruvate kinase, 50-51, 59-62, 70 steriodogenesis, 169- 170 tyrosine hydroxylase, 253-256 Protein kinase C, 30, 38, 242, 257, 265, 290, 298, 329, 341, 346, 348 light chain kinase, 399-403 Protein kinase, cGMP-dependent, 154, 242 light chain kinase, 399-403 neuronal function, 290, 293-294 Protein phosphatase, 195, 198, 206, 286 acetyl-CoA carboxylase, 141-142 brain, 309-310 inhibitors, 303 cell-specific, 307-308 Protein phosphatase-I, 108-109, 164, 309310 Protein phosphatase-2, 108- 109, 309-310 Protein phosphatase2A, 62-65

SUBJECT INDEX Protein phosphatase-2C, 62-63 Protein phosphorylation Blastocladiella emersonii, 438-439 brain, system types, 286-287 ion channel, 335-357 Mucor rourii, 436-437 neuronal function, 287-290 Neurospora, 4356-436 photosynthesis, 426-428 prokaryote, 42 1-429 protozoa, 447-449 slime mold, 440-447 sugar transport, 425-426 yeast, 430-435 Protein synthesis see also Eukaryotic initiation factor guanine nucleotide-binding, 376-377 heme regulated initiation kinase, 369-371 inhibitors, 373-376 initiation, 360-36 1 Proteolysis intracellular, 208-209 phosphorylation-activated, 207-208 Pterin, 218 tyrosine hydroxylase cofactor, 261 Purkinje cell, 293 phosphoprotein, 302, 308 Putrescine, 85 Pyruvate carboxylase, 48, 132 Pyruvate dehydrogenase, 125 Pyruvate dehydrogenase kinase, 300 isolation, 82 physicochemical properties, 82-23 regulatory properties, 83-84 Pyruvate dehydrogenase phosphatase, 108 isolation, 84 physicochemical properties, 84 regulatory properties, 84-86 Pyruvate kinase, 35, 38, 125, 207 allosteric effectors, 59-60 erythrocyte, 68-70 inhibitors, 50, 54 intestinal, 68 introduction, 47-51 isozymes, 48-50 kidney, 68 liver CAMP-dependent protein kinase reaction, 59-62 chicken, 70-71

51 1 dephosphorylation, 62-65 hormonal regulation, 65-68 K,, 53-54, 62 phosphorylation kinetics, 51-55 proteolytic enzyme sensitivity, 55-59 Pyruvate-PEP cycle, regulation, 48 F'yruvate dehydrogenase complex insulin regulation, 89-92 localization, 88 regulation, 86-92 phosphorylation site, 81-82 structure. 79-8 1

R Receptor function desensitization heterologous, 321 -327 homologous, 327-328 epidermal growth factor, 329-330 nicotinic acetylcholine, 329 regulation, 3 19-333 P-adrenergic, 320-328 rhodopsin, 328-329 Reductase kinase, 195 Reductase kinase kinase, 195-196 Reversing factor action site, 368-369 eukaryotic initiation factor, 362-364 Rhodopsin, receptor regulation, 328-329 Rhodopsin kinase, 300 Rhodospirillium rubrum, 426-428 Ribosomal protein S 6, 296 RNA polymerase, 428-429, 432 Rous sarcoma virus, 70, 300

S Saccharomyces cerevisae fructose- I ,6-bisphosphatase, 36 protein phosphorylation, 430-435 Salmonella typhimurium, 423 Sequalene dioxide, 191 Serotonin, 343 Slime mold development, 443-446 protein phosphorylation, 440-447 Sodium channel, phosphorylation biochemical studies, 351-353 significance, 353-354 Sorbose-6-phosphate, 9, 11

SUBJECT INDEX

512 Spermidine, 85, 447 Spermine, 447 pyruvate dehydrogenase phosphatase, 8586 Starvation, 100 IIMO=CuA wtJu€taSc, I YL- I93 pyruvate dehydrogenase complex, 88 Steriodogenesis, 168- I70 Sterol-producing tissue, hormone-sensitive lipase, 168- 190 Streptococcus faecalis, 426 Streptococcus pyogenes, 449 Subtilisin, pyruvate kinase action, 56 Sugar transport, procaryote, 425-426 Synapsin I, 294-296 phosphorylation, 305-306 Synaptosome, 35 1-353

T Tau protein, 296 Tetrahydrobiopterin, 2 IS phenylalanine hydroxylase effect, 235-239 Tetrahydropterin, tryptophan hydroxylase cofactor, 272 Tetrahymena pyriformis, 449 Tetrahymena thermophila, 449 Tetradotoxin, 355 Thiamin diphosphate, 83 Thrombin, 411 Torpedo californica, 349 electric organ, 296, 300 Torpedo marmorata, 350 Transducin, 328, 376 Transferrin, 304 Transferrin receptor, 33 1 Trauma, 100 Trehalase activity, regulation, 433 Trehalose, 433 Triacylglycerol lipase, 170 Trifloorperazine, 396 Triglycerol hydrolase, 151 Tropomyosin, 390 phosphorylation, 405-406 Troponin, 207 phosphorylation, 405-406 Transferrin, 304 Transferrin receptor, 33 1 Trauma, 100 Trehalason, 275-277 Tryptophan hydroxylase, 296

physical properties, 274-275 role, 271-272 Tryptophan hYdronfdaton, 219 reaction catalyzed, a0-221 Tubulin. 2Q6 phosphorytstion. 306-30 Tyrocidin, 439 Tyrosine, serum level, 231-232 Tyrosine hydroxylase, 296, 302 activator protein, 263-264 CAMP-dependent protein kinase, 253-256, 26 1-263 brain, 250 catecholamine effect, 252 chromaffin cell, 255-258 dibutyryl-CAMP activation, 253 heparin effect, 251 nerve stimulation, 266-269 phosphoprotein, 254 phosphorylated stability, 270-27 1 phosphorylation activation, 25 1-269 physical properties, 249-25 1 pterin cofactor, 253 reaction, 248-249 catalyzed, 220-221 striatal, 258-260, 262 vas deferentia, 252-253 Tyrosine hydroxylation, 219 Tyrosine kinase, 349 Trypanosoma gambense. 448

U Ubiquitin, 207 Ustilago maydis, 439

v Valine, regulation, 98-99 Vas deferentia, tyrosine hydroxylase, 252-253 Vasopressin, 37 acetyl-CoA carboxylase, 131- 132 phenylalanine hydroxylase, 241 pyruvate kinase, 66 Veratridine, 268, 355

Y Yeast enzyme regulation, 433-435 enzyme synthesis regulation, 432-433 protein kinase, 430-43 1